Fungal Biology
Series Editors:
Vijai Kumar Gupta, PhD
Molecular Glycobiotechnology Group, Department of Biochemistry,
School of Natural Sciences, National University of Ireland Galway,
Galway, Ireland
Maria G. Tuohy, PhD
Molecular Glycobiotechnology Group, Department of Biochemistry,
School of Natural Sciences, National University of Ireland Galway,
Galway, Ireland
For further volumes:
http://www.springer.com/series/11224
Vijai Kumar Gupta • Maria G. Tuohy
Editors
Manimaran Ayyachamy
Anthonia O’Donovan • Kevin M. Turner
Associate Editors
Laboratory Protocols
in Fungal Biology
Current Methods in Fungal Biology
Editors
Vijai Kumar Gupta
Molecular Glycobiotechnology Group
Department of Biochemistry
School of Natural Sciences
National University of Ireland Galway
Galway, Ireland
Maria G. Tuohy
Molecular Glycobiotechnology Group
Department of Biochemistry
School of Natural Sciences
National University of Ireland Galway
Galway, Ireland
Assistant Professor of Biotechnology
Department of Science
MITS University
Lakshmangarh (Sikar), Rajasthan, India
Associate Editors
Manimaran Ayyachamy
Molecular Glycobiotechnology Group
Department of Biochemistry
School of Natural Sciences
National University of Ireland Galway
Galway, Ireland
Kevin M. Turner
Manufacturing Sciences and Technology
The Pfizer Biotech Campus at
Grange Castle Pfizer Ireland
Pharmaceuticals
Dublin, Ireland
Anthonia O’Donovan
Molecular Glycobiotechnology Group
Department of Biochemistry
School of Natural Sciences
National University of Ireland Galway
Galway, Ireland
ISBN 978-1-4614-2355-3
ISBN 978-1-4614-2356-0 (eBook)
DOI 10.1007/978-1-4614-2356-0
Springer New York Heidelberg Dordrecht London
Library of Congress Control Number: 2012951631
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Foreword
Fungi represent the fifth kingdom of organisms, which is characterized—second only to prokaryotes—by a huge number of diverse species. Even more,
fungi have developed a tremendous variety in lifestyles, biochemical properties, and morphological characters, the latter having been a permanent challenge for defining species and their identification. They have conquered
practically all habitats, from deep sea water to desert soil, and from prokaryotes to mammals, leading to an array of positive but also negative impacts on
mankind. On the negative side, fungi are known as pathogens of plants—a
situation which seriously affects crop plantations all around the earth—but
also of higher fungi, of lower eukaryotes, and of all animals up to mammals
and men. Also, their versatile metabolism provided them with efficient abilities to colonize almost all material, leading to biodeterioration of various
organic materials including paintings and covers, which allowed them to settle
in buildings and flats resulting in indoor contamination as a major problem of
today. Yet there are also numerous benefits: many fungi are known as beneficial
symbionts of plants, such as plant tissue endosymbionts and mycorrhizas. In
fact, the earth would be devoid of plants in the absence of the latter. Finally
several fungi have been domesticated by humans, either for their use in agriculture (such as for biocontrol of plant or invertebrate pathogens or in plant
growth protection and stimulation), for the preparation of feed- and foodstuff,
and as efficient producers of biotechnological products such as primary metabolites, numerous enzymes, and antibiotics. In the area of modern molecular
biotechnology, fungi such as Pichia pastoris have become important highthroughput hosts for the production of recombinant proteins of bacterial to
human origin. Last but not least, fungi like Saccharomyces cerevisiae,
Neurospora crassa, and Aspergillus nidulans have become model systems for
basic biochemical and genetic research, and an impressing amount of our textbook knowledge would not be available without them. In the current genomic
age, elucidation of the genome inventory of about 50 multicellular asco- and
basidiomycetes and the same number of yeasts has been completed and opened
new avenues for their investigation.
In view of this steadily increased interest in fungi, also the methods needed
for their isolation and identification, as well as their genetic manipulation and
monitoring of gene expression and protein production, have become refined
and complemented. This book aims at presenting an inventory of techniques
and methods that are currently in use for studying fungi: it contains 57 chapters dedicated to description of these techniques, starting from concepts of
v
vi
Foreword
cultivation, enumeration, and visualization of fungi; molecular approaches
for detection and quantification; measurement of relevant enzymes and methods for their application; and the use of bioinformatic tools to investigate
fungal genomes.
As a professional reference, this book is aimed at all people who work
with fungi and should be useful both to academic institutions and research
teams, as well as to teachers, graduate and postgraduate students.
Vienna, Austria
Prof. Christian P. Kubicek
Foreword
It gives me immense pleasure to write a foreword for Laboratory Protocols
in Fungal Biology of Springer, USA edited by Dr. Vijai Kumar Gupta and
Dr. Maria G. Tuohy. After going through the content of this laboratory protocol, I feel that it is a wonderful attempt done by Dr. Gupta to compile together
all the information about the subject that will be highly useful to all mycologists around the globe. I am sure that this volume will be highly useful to all
those concerned with fungi and their biology, including environmental and
public health officers and professionals in the field of interest. The volume is
really exhaustive covering almost all the aspects of fungal biology. It will also
be of interest to postgraduate students in this field and also for one and all
interested in Fungi. Additionally it will be of great market value. This effort of
Dr. Gupta’s is admirable.
Varanasi, India
Prof. R.S. Upadhyay
vii
Preface
The interaction between fungi and their environment is central to many natural processes that occur in the biosphere. The hosts and habitats of these
eukaryotic microorganisms are very diverse; fungi are present in every ecosystem on Earth. The fungal kingdom is equally diverse, consisting of seven
different known phyla. Yet detailed knowledge is limited to relatively few
species. The relationship between fungi and humans has been characterized
by the juxtaposed viewpoints of fungi as infectious agents of much dread and
their exploitation as highly versatile systems for a range of economically
important biotechnological applications. Understanding the biology of different fungi in diverse ecosystems as well as their interactions with living and
nonliving is essential to underpin effective and innovative technological
developments.
The tools and techniques of molecular biology, once reserved for mammalian and bacterial systems, have been adapted and optimized for the analysis of fungal species at the molecular level. Rapid screening techniques based
on screening specific regions in the DNA of fungi have been used in species
comparison and identification and are now being extended across fungal
phyla with the ultimate goal being the assembly of the “Fungal Tree of Life”
by the US National Science Foundation. Within a decade after the Human
Genome Sequence was published, genome sequencing technology has been
adapted to yield the complete genome sequences of not only fungi of commerce and medical relevance, but other more isoteric species. Post-genomics
approaches and systems biology are now also being applied to understanding
the details of fungal biology and the interactions between fungi, their hosts,
and their environment. The majority of fungi are multicellular eukaryotic systems and therefore may be excellent model systems by which to answer fundamental biological questions. A greater understanding of the cell biology of
these versatile eukaryotes will underpin efforts to engineer (e.g., “humanize”)
certain fungal species to provide novel cell factories for production of proteins for pharmaceutical applications. Finally, renewed interest in all aspects
of the biology and biotechnology of fungi may also enable the development
of “one pot” microbial cell factories to meet consumer energy needs into the
twenty first century. To realize this potential and to truly understand the diversity and biology of these eukaryotes, continued development of scientific
tools and techniques is essential.
This publication aims to provide a detailed compendium of analytical
methods used to investigate different aspects of mycology, including fungal
ix
x
Preface
biology and biochemistry, genetics, phylogenetics, genomics, proteomics,
molecular enzymology, and biotechnological applications, in a manner that
reflects the many recent developments of relevance to scientists investigating
the Kingdom of Fungi.
Galway, Ireland
Vijai Kumar Gupta
Maria G. Tuohy
Manimaran Ayyachamy
Anthonia O’Donovan
Kevin M. Turner
Contents
1
Safety Norms and Regulations in Handling
Fungal Specimens.........................................................................
Finola E. Cliffe
2
Methods of Cryopreservation in Fungi ......................................
Ladislav Homolka
3
Long-Term Preservation of Fungal Cultures in All-Russian
Collection of Microorganisms (VKM): Protocols
and Results....................................................................................
Svetlana M. Ozerskaya, Natalya E. Ivanushkina,
Galina A. Kochkina, Svetlana S. Eremina,
Alexander N. Vasilenko, and Nadezhda I. Chigineva
1
9
17
4
Fungal Specimen Collection and Processing .............................
Anthonia O’Donovan, Vijai Kumar Gupta,
and Maria G. Tuohy
5
Chemical and Molecular Methods for Detection
of Toxigenic Fungi and Their Mycotoxins
from Major Food Crops ..............................................................
S. Chandra Nayaka, M. Venkata Ramana, A.C. Udayashankar,
S.R. Niranjana, C.N. Mortensen, and H.S. Prakash
73
Identification Key for the Major Growth Forms
of Lichenized Fungi......................................................................
Jeyabalan Sangeetha and Devarajan Thangadurai
91
6
7
Microscopic Methods for Analytical Studies of Fungi ..............
De-Wei Li
8
Scanning Electron Microscopy for Fungal
Sample Examination ....................................................................
Eduardo Alves, Gilvaine Ciavareli Lucas,
Edson Ampélio Pozza, and Marcelo de Carvalho Alves
9
High-Resolution Imaging and Force Spectroscopy
of Fungal Hyphal Cells by Atomic Force Microscopy ..............
Biplab C. Paul, Hui Ma, Laelie A. Snook, and Tanya E.S. Dahms
67
113
133
151
xi
xii
10
11
Contents
Use of Fourier-Transform Infrared (FTIR)
Microscopy Method for Detection of Phyto-Fungal
Pathogens ......................................................................................
Vitaly Erukhimovitch and Mahmoud Huleihel
Diagnosis of Parasitic Fungi in the Plankton: Technique
for Identifying and Counting Infective Chytrids Using
Epifluorescence Microscopy ........................................................
Télesphore Sime-Ngando, Serena Rasconi,
and Mélanie Gerphagnon
161
169
12
Fungal Cell Wall Analysis ...........................................................
Pilar Pérez and Juan C. Ribas
13
Histopathological Technique for Detection of Fungal
Infections in Plants.......................................................................
Vijai Kumar Gupta and Brejesh Kumar Pandey
197
Development of Media for Growth and Enumeration
of Fungi from Water ....................................................................
Segula Masaphy
201
14
15
Sabouraud Agar for Fungal Growth ..........................................
Janelle M. Hare
16 A Method for the Formation of Candida Biofilms
in 96 Well Microtiter Plates and Its Application
to Antifungal Susceptibility Testing............................................
Christopher G. Pierce, Priya Uppuluri, and Jose L. Lopez-Ribot
175
211
217
17
Screening for Compounds Exerting Antifungal Activities .......
Jean-Paul Ouedraogo, Ellen L. Lagendijk,
Cees A.M.J.J. van den Hondel, Arthur F.J. Ram, and Vera Meyer
18
Fluorescence In Situ Hybridization
of Uncultured Zoosporic Fungi...................................................
Télesphore Sime-Ngando, Marlène Jobard, and Serena Rasconi
231
Staining Techniques and Biochemical Methods
for the Identification of Fungi .....................................................
Jeyabalan Sangeetha and Devarajan Thangadurai
237
19
20
Protocol for the In Vivo Quantification of Superoxide
Radical in Fungi ...........................................................................
Konstantinos Grintzalis, Ioannis Papapostolou,
and Christos Georgiou
225
259
21
Isolation of Intact RNA from Sorted S. cerevisiae Cells
for Differential Gene Expression Analysis ................................. 265
Jeannette Vogt, Frank Stahl, Thomas Scheper, and Susann Müller
22
Quantitative PCR Analysis of Double-Stranded
RNA-Mediated Gene Silencing in Fungi.................................... 279
José J. de Vega-Bartol, Vega Tello, Jonathan Niño, Virginia Casado,
and José M. Díaz-Mínguez
Contents
xiii
23
Semi-Nested PCR Approach to Amplify Large
18S rRNA Gene Fragments for PCR-DGGE Analysis
of Soil Fungal Communities ........................................................
Miruna Oros-Sichler and Kornelia Smalla
24
Proteomic Protocols for the Study of Filamentous Fungi ........
Raquel González Fernández and Jesús V. Jorrín Novo
25
Detection and Quantification of Endoprotease Activity
Using a Coomassie Dye-Binding Assay ......................................
Anthony J. O’Donoghue and Cathal S. Mahon
26
Protocol of a LightCycler™ PCR Assay for Detection
and Quantification of Aspergillus fumigatus DNA
in Clinical Samples of Neutropenic Patients..............................
Birgit Spiess and Dieter Buchheidt
27 Application of Polymerase Chain Reaction and PCR-Based
Methods Targeting Internal Transcribed Spacer Region
for Detection and Species-Level Identification of Fungi...........
K. Lily Therese, R. Bagyalakshmi, and H.N. Madhavan
28
Real-Time PCR Assay in Fungi ..................................................
Naomichi Yamamoto
29
Quantitative Sampling Methods for the Analysis
of Fungi: Air Sampling ................................................................
Mary C. O’Loughlin, Katherine D. Turner, and Kevin M. Turner
289
299
309
315
321
331
337
30 Transformation of Filamentous Fungi in Microtiter Plate.......
Bianca Gielesen and Marco van den Berg
343
31
Molecular Fingerprinting of Fungal Communities in Soil .......
Roberto A. Geremia and Lucie Zinger
349
32
Development of Microsatellite Markers from Fungal
DNA Based on Shotgun Pyrosequencing ...................................
Shaobin Zhong
33
34
35
Multiplex and Quantifiable Detection of Infectious
Fungi Using Padlock Probes, General qPCR,
and Suspension Microarray Readout.........................................
Magnus Jobs, Ronnie Eriksson and Jonas Blomberg
357
363
Rapid Deletion Plasmid Construction Methods
for Protoplast and Agrobacterium-based Fungal
Transformation Systems ..............................................................
María D. García-Pedrajas, Zahi Paz, David L. Andrews,
Lourdes Baeza-Montañez, and Scott E. Gold
375
Improved Transformation Method for Alternaria
Brassicicola and Its Applications ................................................
Yangrae Cho, Akhil Srivastava, and Christopher Nguyen
395
xiv
Contents
36
Methods for High-Quality DNA Extraction from Fungi ..........
Vijai Kumar Gupta, Maria G. Tuohy, and Rajeeva Gaur
37
Production of Recombinant Proteins from Pichia pastoris:
Interfacing Fermentation and Immobilized Metal Ion
Affinity Chromatography ............................................................
Berend Tolner, Gaurav Bhavsar, Bride Foster,
Kim Vigor, and Kerry Chester
407
Development of a Real-Time Quantitative PCR Assay
for the Assessment of Uncultured Zoosporic Fungi ..................
Télesphore Sime-Ngando and Marlène Jobard
421
38
39
40
41
42
Nucleic and Protein Extraction Methods for Fungal
Exopolysaccharide Producers .....................................................
Jochen Schmid, Dirk Mueller-Hagen, Volker Sieber,
and Vera Meyer
Directed Evolution of a Fungal Xylanase for Improvement
of Thermal and Alkaline Stability ..............................................
Dawn Elizabeth Stephens, Suren Singh, and Kugen Permaul
Genome Shuffling Protocol for the Pentose-Fermenting
Yeast Scheffersomyces stipitis ......................................................
Paramjit K. Bajwa, Nicole K. Harner, Terri L. Richardson,
Sukhdeep Sidhu, Marc B. Habash, Jack T. Trevors,
and Hung Lee
Detection and Identification of Fungal Microbial
Volatile Organic Compounds by HS-SPME-GC–MS...............
Bernhard Kluger, Susanne Zeilinger, Gerlinde Wiesenberger,
Denise Schöfbeck, and Rainer Schuhmacher
403
427
435
447
455
43 Transformation Methods for Slow-Growing Fungi ..................
Suman Mukherjee and Rebecca Creamer
467
44
Enzymatic Saccharification of Lignocellulosic Biomass...........
Manimaran Ayyachamy, Vijai Kumar Gupta,
Finola E. Cliffe, and Maria G. Tuohy
475
45
Protoplast Fusion Techniques in Fungi ......................................
Annie Juliet Gnanam
483
46
Large-Scale Production of Lignocellulolytic Enzymes
in Thermophilic Fungi .................................................................
Manimaran Ayyachamy, Mary Shier, and Maria G. Tuohy
489
Panfungal PCR Method for Detection
of Aflatoxigenic Molds .................................................................
Malik M. Ahmad, Pravej Alam, M.Z. Abdin, and Saleem Javed
495
47
48
Protocols for the Quantification of dsDNA
and Its Fragmentation Status in Fungi ......................................
Ioannis Papapostolou, Konstantinos Grintzalis,
and Christos Georgiou
501
Contents
xv
49
Rapid Identification and Detection of Pathogenic
Fungi by Padlock Probes .............................................................
Clement K.M. Tsui, Bin Wang, Cor D. Schoen,
and Richard C. Hamelin
505
50
Drug-Induced Permeabilization in Fungi ..................................
Maria D. Mayan, Alexandra McAleenan, and Priscilla Braglia
51
Extraction and Characterization of Taxol: An Anticancer
Drug from an Endophytic and Pathogenic Fungi .....................
M. Pandi, P. Rajapriya, and P.T. Manoharan
523
Identification of Mycotoxigenic Fungi Using
an Oligonucleotide Microarray...................................................
Eugenia Barros
529
DNA Microarray-Based Detection and Identification
of Fungal Specimens ....................................................................
Minna Mäki
535
Bioinformatic Protocols and the Knowledge-Base
for Secretomes in Fungi ...............................................................
Gengkon Lum and Xiang Jia Min
545
High-Throughput Functional Annotation and Data Mining
of Fungal Genomes to Identify Therapeutic Targets ................
Gagan Garg and Shoba Ranganathan
559
52
53
54
55
56 Application of Support Vector Machines in Fungal Genome
and Proteome Annotation............................................................
Sonal Modak, Shimantika Sharma, Prashant Prabhakar,
Akshay Yadav, and V.K. Jayaraman
57
519
565
Bioinformatics Tools for the Multilocus Phylogenetic
Analysis of Fungi ..........................................................................
Devarajan Thangadurai and Jeyabalan Sangeetha
579
Index ......................................................................................................
593
Contributors
M.Z. Abdin Department of Biotechnology, Jamia Hamdard University, New
Delhi, Delhi, India
Malik M. Ahmad Department of Biotechnology, Jamia Hamdard University,
New Delhi, Delhi, India
Pravej Alam Department of Biotechnology, Jamia Hamdard University,
New Delhi, Delhi, India
Eduardo Alves Department of Phytopathology, Federal University of
Lavras, Lavras, Minas Gerais, Brazil
David L. Andrews Department of Plant Pathology, University of Georgia,
Athens, GA, USA
Manimaran Ayyachamy Department of Biochemistry, School of Natural
Sciences, National University of Ireland, Galway, Ireland
Lourdes Baeza-Montañez Instituto de Hortofruticultura Subtropical y
Mediterránea “La Mayora”, Consejo Superior de Investigaciones Científicas
(IHSM-UMA-CSIC), Estación Experimental “La Mayora”, Algarrobo-Costa,
Málaga, Spain
R. Bagyalakshmi Sankara Nethralaya, Larsen and Toubro Microbiology
Research Centre, Chennai, Tamil Nadu, India
Paramjit K. Bajwa School of Environmental Sciences, University of
Guelph, Guelph, ON, Canada
Eugenia Barros Department of Biosciences, Council for Scientific and
Industrial Research (CSIR), Brummeria, Pretoria, South Africa
Gaurav Bhavsar Department of Oncology, University College London
Cancer Institute, London, UK
Jonas Blomberg Department of Medical Sciences, Uppsala Academic
Hospital, Uppsala University, Uppsala, Sweden
Priscilla Braglia Sir William Dunn School of Pathology, University of
Oxford, Oxford, UK
Dieter Buchheidt Third Department of Internal Medicine, Mannheim
University Hospital, Mannheim, Germany
xvii
xviii
Virginia Casado Department of Microbiologia y Genetica—CIALE,
Universidad de Salamanca, Salamanca, Spain
Kerry Chester Department of Oncology, University College London Cancer
Institute, London, UK
Nadezhda I. Chigineva All-Russian Collection of Microorganisms (VKM
IBPM RAS, Pushchino, Russia), G. K. Skryabin Institute of Biochemistry
and Physiology of Microorganisms, Russian Academy of Science, Pushchino,
Moscow Region, Russia
Yangrae Cho Department of Plant and Environmental Protection Sciences,
University of Hawaii at Manoa, Honolulu, HI, USA
Finola E. Cliffe Department of Biochemistry, School of Natural Sciences,
National University of Ireland Galway, Galway, Ireland
Rebecca Creamer Department of Entomology, Plant Pathology, and Weed
Science, New Mexico State University, Las Cruces, NM, USA
Tanya E.S. Dahms Department of Chemistry and Biochemistry, University
of Regina, Regina, SK, Canada
Marcelo de Carvalho Alves Department of Soil and Rural Engineering,
Campus of the Federal University of Mato Grosso, Federal University of
Mato Grosso, Cuiaba, Mato Grosso, Brazil
José J. de Vega-Bartol Department of Microbiologia y Genetica—CIALE,
Universidad de Salamanca, Salamanca, Spain
José M. Díaz-Mínguez Department of Microbiologia y Genetica—CIALE,
Centro Hispano Luso de Investigaciones Agrarias, Universidad de Salamanca,
Salamanca, Spain
Svetlana S. Eremina All-Russian Collection of Microorganisms (VKM
IBPM RAS, Pushchino, Russia), G. K. Skryabin Institute of Biochemistry
and Physiology of Microorganisms, Russian Academy of Science, Pushchino,
Moscow Region, Russia
Ronnie Eriksson Livsmedelsverket, Uppsala, Sweden
Vitaly Erukhimovitch Analytical Equipment Unit, Ben-Gurion University
of the Negev, Beer-Sheva, Israel
Raquel González Fernández Department of Biochemistry and Molecular
Biology, University of Córdoba, Córdoba, Spain
Bride Foster Department of Oncology, University College London Cancer
Institute, London, UK
María D. García-Pedrajas Instituto de Hortofruticultura Subtropical y
Mediterránea “La Mayora”, Consejo Superior de Investigaciones Científicas
(IHSM-UMA-CSIC), Estación Experimental “La Mayora”, Málaga, Spain
Gagan Garg Department of Chemistry and Biomolecular Sciences,
Macquarie University, Sydney, NSW, Australia
Contributors
Contributors
xix
Rajeeva Gaur Department of Microbiology, Dr. R.M.L. Avadh University,
Faizabad, Uttar Pradesh, India
Christos Georgiou Department of Biology, University of Patras, Patras,
Achaia, Greece
Roberto A. Geremia Laboratoire d’Ecologie Alpine, CNRS/UJF, Université
Joseph Fourier, Grenoble, France
Mélanie Gerphagnon Université Blaise Pascal, Aubière, France
Bianca Gielesen DSM Biotechnology Center, Delft, Zuid Holland, The
Netherlands
Annie Juliet Gnanam College of Natural Science, Institute for Cellular and
Molecular Biology, University of Texas at Austin, Austin, TX, USA
Scott E. Gold United States Department of Agriculture—Agricultural
Research Unit (USDA–ARS), Toxicology and Mycotoxin Research Unit,
Athens Georgia, USA
Konstantinos Grintzalis Department of Biology, University of Patras,
Patras, Achaia, Greece
Vijai Kumar Gupta Molecular Glycobiotechnology Group, Department of
Biochemistry, School of Natural Sciences, National University of Ireland
Galway, Galway, Ireland
Assistant Professor of Biotechnology, Department of Science, Faculty of
Arts, Science & Commerce, MITS University, Rajasthan, India
Marc B. Habash School of Environmental Sciences, University of Guelph,
Guelph, ON, Canada
Richard C. Hamelin Department of Forest Sciences, The University of
British Columbia, Vancouver, BC, Canada
Laurentian Forestry Centre, Natural Resources Canada, Quebec, QC,
Canada
Janelle M. Hare Department of Biology and Chemistry, Morehead State
University, KY, USA
Nicole K. Harner School of Environmental Sciences, University of Guelph,
Guelph, ON, Canada
Ladislav Homolka Department of Ecology of Microorganisms, Institute of
Microbiology, Academy of Sciences of the Czech Republic, Prague, Czech
Republic
Mahmoud Huleihel Department of Virology and Developmental Genetics,
Ben-Gurion University of the Negev, Beer-Sheva, Israel
Natalya E. Ivanushkina All-Russian Collection of Microorganisms (VKM
IBPM RAS, Pushchino, Russia), G. K. Skryabin Institute of Biochemistry
and Physiology of Microorganisms, Russian Academy of Science, Pushchino,
Moscow Region, Russia
xx
Saleem Javed Department of Biochemistry, Jamia Hamdard University,
New Delhi, India
V.K. Jayaraman Scientific and Engineering Computing Group (SECG),
Centre for Development of Advanced Computing (C-DAC), University of
Pune, Pune, Maharashtra, India
Marlène Jobard LMGE UMR CNRS, U.F.R. Sciences et Technologies,
Aubière Cedex, France
Magnus Jobs School of Health and Social Studies, Högskolan Dalarna,
Uppsala University, Falun, Sweden
Bernhard Kluger Department for Agrobiotechnology (IFA-Tulln),
University of Natural Resources and Life Sciences Vienna, Tulln, Austria
Galina A. Kochkina All-Russian Collection of Microorganisms (VKM
IBPM RAS, Pushchino, Russia), G. K. Skryabin Institute of Biochemistry
and Physiology of Microorganisms, Russian Academy of Science, Pushchino,
Moscow Region, Russia
Christian P. Kubicek Department of Chemical Engineering, Vienna
University of Technology, Vienna, Austria
Ellen L. Lagendijk Department of Molecular Microbiology and
Biotechnology, Leiden University, Leiden, The Netherlands
Hung Lee University of Guelph, School of Environmental Sciences, Guelph,
ON, Canada
De-Wei Li Valley Laboratory, The Connecticut Agricultural Experiment
Station, Windsor, CT, USA
Jose L. Lopez-Ribot Department of Biology, South Texas Center for
Emerging Infectious Diseases, The University of Texas at San Antonio, San
Antonio, TX, USA
Gilvaine Ciavareli Lucas Department of Phytopathology, Federal University
of Lavras, Lavras, Minas Gerais, Brazil
Departamento de Fitopatologia, Universidade Federal de Lavras, Caixa
postal, Lavras, Minas Gerais, Brazil
Gengkon Lum Department of Computer Science and Information Systems,
Youngstown State University, Youngstown, OH, USA
Hui Ma Department of Chemistry, National University of Singapore,
Singapore
Alexandra McAleenan Clinical Sciences Centre, Imperial College London,
London, UK
H.N. Madhavan Sankara Nethralaya, Larsen and Toubro Microbiology
Research Centre, Chennai, Tamil Nadu, India
Cathal S. Mahon Department of Pharmaceutical Chemistry, University of
California—San Francisco, San Francisco, CA, USA
Contributors
Contributors
xxi
Minna Mäki Program Leader, NAT, Orion Diagnostica Oy, Espoo, Finland
P.T. Manoharan Department of Botany, Vivekananda College, Madurai,
Tamil Nadu, India
Segula Masaphy Department of Applied Microbiology and Mycology,
MIGAL, Kiryat Shmona, Israel
Maria D. Mayan Fundación CHUAC, Biomedical Research Center—
INIBIC, A Coruña, Galicia, Spain
Vera Meyer Department of Applied and Molecular Microbiology, Berlin
University of Technology, Berlin, Germany
Xiang Jia Min Department of Biological Sciences, Center for Applied
Chemical Biology, Youngstown State University, Youngstown, OH, USA
Sonal Modak Bioinformatics Centre, University of Pune, Pune, Maharashtra,
India
C.N. Mortensen Department of Agriculture and Ecology, University of
Copenhagen, Copenhagan, Taastrup, Denmark
Dirk Mueller-Hagen Department of Applied and Molecular Microbiology,
Technische Universität Berlin, Berlin, Germany
Suman Mukherjee Laboratory of Biochemistry and Genetics, NIDDK,
National Institutes of Health, Bethesda, MD, USA
Susann Müller Department of Environmental Microbiology, Helmholtz
Centre for Environmental Research—UFZ, Leipzig, Saxonia, Germany
S. Chandra Nayaka Department of Studies in Biotechnology, Asian Seed
Health Centre, University of Mysore, Mysore, Karnataka, India
Christopher Nguyen Department of Plant and Environmental Protection
Sciences, University of Hawaii at Manoa, Honolulu, HI, USA
Jonathan Niño Department of Microbiologia y Genetica—CIALE,
Universidad de Salamanca, Villamayor, Salamanca, Spain
S.R. Niranjana Department of Studies in Biotechnology, University of
Mysore, Mysore, Karnataka, India
Jesús V. Jorrín Novo Department of Biochemistry and Molecular Biology,
University of Córdoba, Córdoba, Spain
Anthony J. O’Donoghue Department of Pharmaceutical Chemistry,
University of California—San Francisco, San Francisco, CA, USA
Anthonia O’Donovan Discipline of Biochemistry, School of Natural
Sciences, National University of Ireland, Galway, Ireland
Mary C. O’Loughlin Department of Life Sciences, University of Limerick,
Castletroy, Limerick, Ireland
Miruna Oros-Sichler Institute for Epidemiology and Pathogen Diagnostics,
Julius Kühn Institut, Braunschweig, Lower Saxony, Germany
xxii
Jean-Paul Ouedraogo Department Applied and Molecular Microbiology,
Institute of Biotechnology, Berlin University of Technology, Berlin,
Germany
Svetlana M. Ozerskaya All-Russian Collection of Microorganisms (VKM
IBPM RAS, Pushchino, Russia), G. K. Skryabin Institute of Biochemistry
and Physiology of Microorganisms, Russian Academy of Science, Pushchino,
Moscow Region, Russia
Brejesh Kumar Pandey Molecular Plant Pathology Laboratory, Central
Institute for Subtropical Horticulture, Indian Council of Agricultural
Research, Lucknow, Uttar Pradesh, India
M. Pandi Department of Molecular Microbiology, School of Biotechnology,
Madurai Kamaraj University, Madurai, Tamil Nadu, India
Ioannis Papapostolou Department of Biology, University of Patras, Patras,
Achaia, Greece
Biplab C. Paul Department of Chemistry and Biochemistry, University of
Regina, Regina, SK, Canada
Zahi Paz Department of Plant Pathology, University of Georgia, Athens,
GA, USA
Pilar Pérez Departamento de Microbiología CSIC/Universidad de
Salamanca, Instituto de Biología Funcional y Genómica (IBFG), Salamanca,
Spain
Kugen Permaul Department of Biotechnology and Food Technology,
Durban University of Technology, Durban, Kwa-Zulu-Natal, South Africa
Christopher G. Pierce Department of Biology, South Texas Center for
Emerging Infectious Diseases, The University of Texas at San Antonio, San
Antonio, TX, USA
Edson Ampélio Pozza Departamento de Fitopatologia, Universidade
Federal de Lavras, Caixa postal, Lavras, Minas Gerais, Brazil
Prashant Prabhakar Department of Biotechnology, Dr. D.Y. Patil
University, Pune, Maharashtra, India
H.S. Prakash Department of Studies in Biotechnology, Asian Seed Health
Centre, University of Mysore, Mysore, Karnataka, India
P. Rajapriya Department of Microbiology, Srinivasan College of Arts and
Science, Perambalur, Tamil Nadu, India
Arthur F.J. Ram Department of Molecular Microbiology and Biotechnology,
Leiden University, Leiden, BE, The Netherlands
M. Venkata Ramana Department of Studies in Microbiology, University of
Mysore, Mysore, Karnataka, India
Shoba Ranganathan Department of Chemistry and Biomolecular Sciences,
Macquarie University, Sydney, NSW, Australia
Contributors
Contributors
xxiii
Department of Biochemistry, Yong Loo Lin School of Medicine, National
University of Singapore, Singapore, Singapore
Serena Rasconi Department of Biology, University of Oslo, Oslo, Norway
Juan C. Ribas Departamento de Microbiología CSIC/Universidad de
Salamanca, Senior Scientist from the Spanish Research Council (Consejo
Superior de Investigaciones Científicas, CSIC), Instituto de Biología
Funcional y Genómica (IBFG), Salamanca, Spain
Terri L. Richardson School of Environmental Sciences, University of
Guelph, Guelph, ON, Canada
Jeyabalan Sangeetha Department of Zoology , Karnataka University,
580003, Dharwad, Karnataka, India
Thomas Scheper Chip Technology Institute for Technical Chemistry,
University of Hannover, Hannover, Lower Saxony, Germany
Jochen Schmid Department of Chemistry of Biogenic Resources, Technische
Universität München, Straubing, Bavaria, Germany
Cor D. Schoen Department of Bio-Interactions and Plant Health, Plant
Research International B. V, Wageningen, The Netherlands
Denise Schöfbeck Department for Agrobiotechnology (IFA-Tulln),
University of Natural Resources and Life Sciences Vienna, Tulln, Austria
Rainer Schuhmacher Department for Agrobiotechnology (IFA-Tulln),
University of Natural Resources and Life Sciences Vienna, Tulln, Austria
Shimantika Sharma Department of Biotechnology, Dr. D.Y. Patil University,
Pune, Maharashtra, India
Mary Shier Department of Biochemistry, National University of Ireland,
Galway, Ireland
Sukhdeep Sidhu School of Environmental Sciences, University of Guelph,
Guelph, ON, Canada
Volker Sieber Chemistry of Biogenic Resources, Technische Universität
München, Straubing, Bavaria, Germany
Télesphore Sime-Ngando UMR CNRS 6023, Université Blaise Pascal,
Clermont II, Aubière, Cedex, France
Suren Singh Department of Biotechnology and Food Technology, Durban
University of Technology, Durban, Kwa-Zulu-Natal, South Africa
Kornelia Smalla Julius Kühn Institut, Federal Research Centre for
Cultivated Plants, Institute for Epidemiology and Pathogen Diagnostics,
Braunschweig, Lower Saxony, Germany
Laelie A. Snook Department of Human Health and Nutritional Sciences,
Guelph, Ontario, Canada
xxiv
Birgit Spiess Third Department of Internal Medicine, Mannheim University
Hospital, Mannheim, Germany
Akhil Srivastava Department of Plant and Environmental Protection
Sciences, University of Hawaii at Manoa, Honolulu, HI, USA
Frank Stahl Chip Technology Institute for Technical Chemistry, University
of Hannover, Hannover, Germany
Dawn Elizabeth Stephens Department of Biotechnology and Food
Technology, Durban University of Technology, Durban, Kwa-Zulu-Natal,
South Africa
Vega Tello Department of Microbiologia y Genetica—CIALE, Universidad
de Salamanca, Salamanca, Spain
Devarajan Thangadurai Department of Botany, Karnataka University,
Dharwad, Karnataka, India
K. Lily Therese Sankara Nethralaya, Larsen and Toubro Microbiology
Research Centre, Vision Research Foundation, Chennai, Tamil Nadu, India
Berend Tolner Department of Oncology, University College London Cancer
Institute, London, UK
Jack T. Trevors School of Environmental Sciences, University of Guelph,
Guelph, ON, Canada
Clement K.M. Tsui Department of Forest Sciences, The University of
British Columbia, Vancouver, BC, Canada
Maria G. Tuohy Department of Biochemistry, School of Natural Sciences,
National University of Ireland, Galway, Ireland
Katherine D. Turner School of Natural Sciences, Centre for Chromosome
Biology, National University of Ireland Galway, Galway, Ireland
Kevin M. Turner Manufacturing Sciences and Technology, Pfizer Ireland
Pharmaceuticals, The Pfizer Biotech Campus at Grange Castle, Dublin,
Ireland
A.C. Udayashankar Department of Studies in Biotechnology, Asian Seed
Health Centre, University of Mysore, Mysore, Karnataka, India
R.S. Upadhyay Department of Botany, Centre of Advanced Study, Banaras
Hindu University, Varanasi, Uttar Pradesh, India
Priya Uppuluri Department of Biology, South Texas Center for Emerging
Infectious Diseases, The University of Texas at San Antonio, San Antonio,
TX, USA
Marco van den Berg Applied Biochemistry and Screening, DSM
Biotechnology Center, Delft, Zuid-Holland, The Netherlands
Cees A.M.J.J. van den Hondel Department of Molecular Microbiology
and Biotechnology, Leiden University, Leiden, BE, The Netherlands
Contributors
Contributors
xxv
Alexander N. Vasilenko All-Russian Collection of Microorganisms (VKM
IBPM RAS, Pushchino, Russia), G. K. Skryabin Institute of Biochemistry
and Physiology of Microorganisms, Russian Academy of Science, Pushchino,
Moscow Region, Russia
Kim Vigor Department of Oncology, University College London Cancer
Institute, London, UK
Jeannette Vogt Department of Environmental Microbiology, Helmholtz
Centre for Environmental Research—UFZ, Leipzig, Saxonia, Germany
Bin Wang Westmead Hospital, Centre of Virus Research, Westmead
Millennium Institute, University of Sydney, Westmead, NSW, Australia
Gerlinde Wiesenberger Institute of Applied Genetics and Cell Biology,
University of Natural Resources and Life Sciences Vienna, Tulln, Austria
Akshay Yadav Scientific and Engineering Computing Group (SECG),
Centre for Development of Advanced Computing (C-DAC), University of
Pune, Pune, Maharashtra, India
Naomichi Yamamoto Department of Environmental Health, Graduate
School of Public Health, Seoul National University, 1 Gwanak-ro, Gwanak-gu,
Seoul, Korea
Susanne Zeilinger Research Area Gene Technology and Applied
Biochemistry, Institute for Chemical Engineering, Vienna University of
Technology, Vienna, Austria
Shaobin Zhong Department of Plant Pathology, North Dakota State
University, Fargo, ND, USA
Lucie Zinger Laboratoire d’Ecologie Alpine, CNRS/UJF, Université Joseph
Fourier, Grenoble, France
1
Safety Norms and Regulations
in Handling Fungal Specimens
Finola E. Cliffe
Abstract
This chapter provides basic safety information required when handling
fungal cultures and when performing the procedures outlined in this manual. Several topics are discussed including routine precautions when working with fungal organisms.
Keywords
Fungi • Mycology • Health and safety • Biosafety • Biosafety levels
Introduction
Biosafety measures designed to ensure the safety
of laboratory workers include the use of various
primary and secondary barriers, many of which
are due to the advent of new technologies in the
fields of materials science and engineering.
Personnel undertaking the protocols in this manual may come across potentially hazardous
materials such as pathogenic and infectious
biological fungal agents, in addition to toxic
F.E. Cliffe (*)
Department of Biochemistry,
Molecular Glycobiotechnology Group, School of Natural
Sciences, National University of Ireland Galway,
University Road, Galway, Ireland
e-mail: fcliffe@gmail.com
chemicals and carcinogenic, mutagenic, or
teratogenic reagents. In the case of fungal specimens, it has long been acknowledged that laboratory workers can attain infections from the
agents they work with.
There have been many reported cases of laboratory-acquired infection, with countless more
cases undoubtedly left unreported. Inhalation
appears to be the most prominent route of exposure. Fungal hyphae in nature develop structures
such as conidia on fruiting bodies or hyphal elements that develop into transmissible subsegments, which are ultimately designed for optimum
dispersal in air. These elements are designed to
be readily discharged, resistant to desiccation,
and to remain aloft for long periods of time. Once
inhaled by a host, the conidia develop into the
yeast phase and can be found in the tissue of
infected hosts [1]. Even with the advances in biosafety training and education, laboratory-acquired
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_1, © Springer Science+Business Media, LLC 2013
1
2
fungal infections continue to occur. The dimorphic
fungi Blastomyces dermatitidis, Coccidioides
immitis, and Histoplamsa caspsulatum, for example, were found to be responsible for the majority
of laboratory-acquired fungal infections in the
United States [2–4]. Laboratory-associated pulmonary infections have occurred following the
inhalation of conidia from mold-form cultures of
B. dermatitidis [5, 6] and local infections from
the accidental parenteral inoculation with infected
tissues or cultures containing yeast forms of the
fungus [7, 8] have been documented. Various
reports of laboratory-associated C. immitis are
reported in the literature prior to 1980 [9–11]
including a case recorded by Nabarro [12] where
a biochemist developed an intense acute infection after working with a colonial growth.
Laboratory-associated histoplasmosis occurs
mainly through inhalation of conidia produced
by the mold form of the fungus [4, 13]; however,
cutaneous infections have occurred due to accidental inoculation [14, 15].
These incidences indicate the ongoing occurrences of laboratory-acquired infections as a
result of simple and preventable laboratory errors.
As mentioned, the bulk of laboratory-acquired
fungal infections are caused by inhalation of
infectious conidia from the mold form, resulting
in pulmonary infections; for example, the simple
processes of opening of a culture plate lid can
result in the release of large numbers of conidia
[16]. To reduce the risk of infection it is practical
to handle all fungal cultures under the conditions
of biosafety laboratory containment BSL-2 or
BSL-3 [17].
New biosafety technologies and associated
guidelines have been developed to considerably
improve ways to safely use fungal material. An
enhanced understanding of the risks associated
with various manipulations of many agents transmissible by different routes has enabled laboratory workers to apply appropriate biosafety
practices to specific laboratory areas. These
safety guidelines include engineering controls,
management policies, work practices and procedures, as well as medical interventions. However,
users must always progress with the caution
F.E. Cliffe
associated with good laboratory practice, under
the supervision of personnel responsible for
implementing laboratory safety programs at their
institutions.
Biosafety Levels
Several biosafety levels (BSL) have been developed for laboratories to provide increasing levels of staff and environmental protection. BSLs
are guidelines that describe appropriate containment equipment, facilities, and procedures for
use by laboratory workers. The BSLs range from
biosafety level 1 (BSL 1) to biosafety level 4
(BSL 4), and each BSL is based on the increased
risk associated with the pathogenicity of the
microorganisms encountered. Most clinical
microbiology laboratories follow BSL 2 practices. When working with highly infectious
agents for which the risk of aerosol transmission
is greater, laboratories should follow BSL 3
practices.
BSL-1 is suitable for working with fungal
agents that are not known to cause disease in
healthy humans. BSL-1 practices, safety equipment, and facility design and construction are
appropriate for undergraduate and secondary
educational training and teaching laboratories,
and for other laboratories in which work is done
with defined and characterized strains of viable
microorganisms not known to consistently cause
disease in healthy adult humans. It is important to
remember, however, that many agents not ordinarily associated with disease processes in
humans are opportunistic pathogens and may
cause infection in the young, the aged, and
immunodeficient or immunosuppressed individuals. BSL-1 represents a basic level of containment that relies on standard microbiological
practices with no special primary or secondary
barriers recommended, other than a sink for hand
washing.
BSL-2 should be used for work involving
fungal agents that pose a moderate potential
hazard to laboratory workers. These agents
include the large group of opportunistic fungal
1
Safety Norms and Regulations in Handling Fungal Specimens
pathogens such as Aspergillus spp. and Fusarium
spp. Some protocols can be carried out on an
open bench providing the potential for aerosol
production is low [17]. Although organisms regularly employed at Biosafety Level 2 are not
known to be transmissible by the aerosol route,
procedures with aerosol or high splash potential
that may increase the risk of such personnel
exposure must be conducted in primary containment equipment, or in devices such as a biological safety cabinet (BSC) or safety centrifuge
cups. Personal protective equipment (PPE), such
as splash shields, face protection, gowns, and
gloves should be used as appropriate. In addition, secondary barriers such as hand-washing
sinks and waste decontamination facilities must
be accessible to decrease the chance of environmental contamination [18].
BSL-3 is appropriate for work with infectious
agents, which may cause serious or potentially
lethal diseases as a result of inhalation. The fungal pathogens C. immitis and H. capsulatum fall
into this group. Autoinoculation, ingestion, and
exposure to infectious aerosols are the main hazards to personnel working with these organisms.
All laboratory operations should be performed in
a BSC or other enclosed apparatus, such as a gastight aerosol generation chamber. Secondary barriers for this level include controlled access to the
laboratory and ventilation requirements that minimize the release of infectious aerosols from the
laboratory. Within this level, primary and secondary barriers to protect personnel in contiguous
areas, the community, and the environment from
exposure to potentially infectious aerosols have
been highlighted [18].
At present, no fungal agents have been
classified for use at BSL-4. BSL-4 practices,
safety equipment, and facility design and construction are applicable for work with hazardous
and exotic agents that pose a high individual
risk of life-threatening disease, which may be
transmitted via the aerosol route, and for
which there is no available vaccine or therapy.
The primary hazards to personnel working
with Biosafety Level 4 agents are respiratory
3
exposure to infectious aerosols, mucous membrane or broken skin exposure to infectious
droplets, and autoinoculation. All manipulations
of potentially infectious diagnostic materials,
isolates, and naturally or experimentally infected
animals pose a high risk of exposure and infection to laboratory personnel, the community,
and the environment. The laboratory worker’s
complete isolation from aerosolized infectious
materials is accomplished primarily by working
in a Class III BSC or in a full-body, air-supplied,
positive-pressure personnel suit. The BSL-4
facility itself is generally a separate building or
completely isolated zone with complex, specialized ventilation requirements and waste management systems to prevent release of viable
agents to the environment [18].
The safety plan of a laboratory should address
general considerations, chemical safety, and
section-specific safety. In the case of mycology
laboratories, as with all laboratories, each section requires a site-specific risk assessment to
address biohazard considerations and to outline
measures for staff protection. Table 1.1 outlines
an example of the type of assessment that should
be performed.
Materials (See Note 1)
1. Sterile distilled water
2. PPE such as coats, gowns, gloves, masks,
face shields, safety glasses
3. Ethanol (70%)
4. Biosafety cabinet
5. Eyewash station
6. Hand washing sinks
7. HEPA filtered respirators or masks
8. Plasticwear (substitute for glass)
9. Centrifuge safety cups
10. Containers for transport of specimens, waste,
and sharps.
11. Biohazard bags
12. Biohazard labels
13. Automatic or mechanical pipetting devices
4
F.E. Cliffe
Table 1.1 Example of a risk assessment for mycology (abridged list of tasks) [19]
Task
Plating/inoculation of specimens
Place specimens on media
Inoculate primary culture media with specimens
Storage/disposal/retrieval of specimens
Culture reading
Prepare fungal wet mounts (KOH-calcofluor)
and/or India ink preps on isolates
Read fungal wet mounts (KOH-calcofluor) and/
or India ink preps on isolates
Examine fixed smears
Examine sealed cultures
Manipulation of yeast isolated in culture,
subculture of colonies, preparation of wet
mounts/smears
Manipulation of yeast, preparation of suspension
Manipulation of molds isolated in culture,
subculture of colonies or broth, preparation of
wet mounts/smears/fixed slides
Transfer of molds between BSC and incubators
and/or storage areas
Nucleic acid probe or antigen detection
performed on mold isolates
Transport of fungal blood cultures between
bench/incubator
Subculture/inoculate fungal blood culture
Exposure risk
Biosafety level
Personal protective equipment
Low
Moderate
Low
BSL-1
BSL-2
BSL-1
Lab coat, gloves, face shield
Lab coat, gloves, BSC
Lab coat, gloves
Moderate
BSL-2
Low
BSL-2
Lab coat, gloves, biosafety
cabinet (BSC)
Lab coat/gloves
Low
Low
Low
BSL-1
BSL-1
BSL-1
Lab coat
Lab coat
Lab coat
Low to
moderate
Moderate
BSL-1
BSL-2
Lab coat, gloves, face shield,
or BSC
Lab coat, gloves, BSC
Moderate
BSL-1
Lab coat
Moderate
BSL-2
Lab coat, gloves, BSC
Moderate
BSL-1
Lab coat/gloves
High
BSL-2
Lab coat, gloves, BSC
Methods
Routine Precautions When Working
with Fungal Cultures
The following practices are recommended for all
laboratories handling potentially dangerous fungal agents:
1. Limit access to work areas. Close doors during work with research materials and lock
when staff are not present in the laboratory.
2. Decontaminate all work surfaces after each
working day using an appropriate disinfectant
and fungicide such as 2% amphyl solution.
Decontaminate all spills of viable material
immediately and all liquid or solid wastes that
have come in contact with viable material.
3. Aerosol-containment safety carriers much be
used in the centrifuge for use with all infectious and potentially infectious materials.
4. Do not pipette by mouth.
5. Do not allow eating, drinking, smoking, or
application of cosmetics in the work area. Do
not store food in refrigerators that contain
laboratory supplies.
6. Wash hands with soap or detergent after handling viable materials or removing gloves,
and before leaving the laboratory. Do not
handle telephones, doorknobs, or other common utensils without washing hands.
7. When handling viable materials, minimize
creation of aerosols. For example, aerosolcontainment safety carriers must be used in
the centrifuge for use with all infectious and
potentially infectious materials.
8. Wear laboratory coats (preferably disposable) when in work area, but do not wear
them away from the work area. Observers
who are not handling infectious and potentially infectious material, but are present in
the laboratory where such an activity is in
1
Safety Norms and Regulations in Handling Fungal Specimens
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
progress, will wear a protective gown and
mask if necessary.
Wear disposable gloves when handling viable materials. These should be disposed of as
biohazardous waste. Change gloves if they
are directly contaminated. Do not wear
gloves away from the work area.
Any potentially contaminated material
should be autoclaved for 30 min at 121 °C
prior to reuse.
Use sharps only when no alternatives (e.g.,
safety devices or nonsharps) exist.
Wear eye/face protection if splashes or sprays
are anticipated.
Transport materials outside of the laboratory
using secondary containment.
Transfer materials to other facilities according to federal and international regulations.
Be acquainted with written instructions for
laboratory procedures and proper responses
to emergencies.
Report spills, exposures, illnesses, and injuries immediately.
Control pest populations. Windows in the
laboratory that can be opened must be
equipped with screens to exclude insects.
Use furniture that is easy to clean (i.e., with
smooth, waterproof surfaces) and as few
seams as possible.
Keep biohazard waste in covered containers
free from leaks. Use biohazard bags as
required by institutional procedure. Dispose
of according to institutional procedure.
When C. immitis or H. capsulatum is used,
all isolates should be maintained on slants to
prevent aerosolization when cultures are
manipulated.
Seal all plated media with a gas-permeable
tape before removing them from the biosafety cabinet.
All excess materials are to be discarded in
biohazard bags secured with tape and placed
in containers designated for disposal of biohazardous waste.
All laboratory surfaces that come in contact
with fungal specimens or cultures should be
decontaminated with 10% bleach that is
freshly prepared each day [19].
5
Experimentation
Administrative controls established prior to
commencing experimentations should be used.
These include
1. Substitution of hazardous materials with less
hazardous materials in experiments.
2. Documentation of laboratory procedures that
state safe work practices and the use of PPE.
Engineering controls that are mechanical in
nature must also be used. Such controls employ a
mechanical way to isolate the worker from the
hazard. In fungal culturing such controls
include:
1. The availability of needles or blades that are
self-sheathing or automatically retracted and
are known as “safer sharps.”
2. The use of plastic instead of glass for vials,
flasks, beakers, etc.
3. Containers for waste collection should be
sturdy, compatible with the waste to be collected, labeled suitably, and kept closed when
not in use.
4. Biological safety cabinets should be employed
as often as possible. These cabinets protect the
worker by scrubbing the particulate-laden air
currents using high efficient particulate air
(HEPA) filters.
5. PPE is employed to firstly provide a barrier to
possible routes of entry on a workers body and
secondly to protect clothing and shoes so that
laboratory contaminants are not transferred
from the laboratory to external areas. PPE
must be selected as appropriate to the work
being undertaken.
6. The building ventilation system must be so
constructed that air from the laboratory is not
recirculated to other areas within the building. Air may be high-efficiency particulate
air (HEPA) filtered, reconditioned, and recirculated within that laboratory. When exhaust
air from the laboratory is discharged to the
outside of the building, it must be dispersed
away from occupied buildings and air
intakes. Depending on the agents in use, this
air may be discharged through HEPA filters.
A heating, ventilation, and air-conditioning
(HVAC) control system may be installed to
6
prevent sustained positive pressurization of
the laboratory.
7. An autoclave for the decontamination of contaminated waste material should be available
in the containment laboratory. If waste has to
be transferred from the containment laboratory for decontamination and disposal, it must
be transported in sealed, unbreakable, and
leakproof vessels according to national or
international regulations [20].
Accidents
1. Workers must vacate the work area in the
event of contamination (e.g., breakage of culture tubes, etc.) until the whole area has been
decontaminated.
2. If a culture tube/plate is broken, several steps
must be taken
(a) All staff must exit the room and close door
as they leave.
(b) To decontaminate the room, the appropriate PPE must be worn (mask, gown,
gloves, shoes, and hair cover).
(c) Enter the room and cover spill with paper
towels soaked thoroughly with disinfectant such as 2% amphyl solution. Leave
the towels to stand for 1 h before cleaning
up the spill and keep the area wet with
amphyl to prevent dried particles from
becoming airborne. If required, disinfect
all contaminated equipment, i.e., specimen containers, culture tubes, pens, etc.
(d) Exit area and do not re-enter for 1 h.
(e) Contaminated clothing must be discarded
or autoclaved for 1 h at 121 °C.
3. The laboratory supervisor must be informed
of the accident and an incident report must be
completed.
4. For accidents involving the eye,
(a) Go to the eye wash station and call for
help.
(b) Eyes must be washed carefully and
thoroughly for at least 15 min to remove
chemicals or particles.
(c) Seek medical attention.
(d) The laboratory supervisor must be informed
and an incident report must be completed.
F.E. Cliffe
Emergency Preparedness and Response
Emergencies and disruptions to the normal
working environment, such as hurricanes and
other disasters, can occur. Possible disruptions
can include spills, exposures, injuries, power or
water loss, equipment failure, fire, or flooding. The
response to each of these emergencies will depend
on the individual and institutional circumstances,
which are too varied to discuss in detail. However,
a written emergency and evacuation plan should
be put in place and communicated to all personnel for such situations to circumvent employee
injury or contamination via fungal agents.
To prepare the mycology laboratory for an emergency the following steps should be taken:
1. Cover all external windows with wood.
2. Ensure all refrigerators and freezers are connected to emergency electrical outlets.
3. Incubators should be locked to prevent breakage and subsequent dispersion of cultures
containing potentially pathogenic fungi.
4. Computer terminals and electronic equipment
should be disconnected and moved to one
room and covered with plastic.
5. Work and log books and other essential paperwork should also be moved to one room and
covered with plastic.
Information and Training
Findings from several studies have suggested that
formal training in mycology laboratories is inadequate and must be supplemented. All of the
information discussed in this chapter must be
communicated to laboratory personnel by management. Lack of communication of safety guidelines can result in health and safety incidences.
1. Every laboratory worker should know the
location and proper use of PPE and first aid
equipment (e.g., eye wash stations, etc.).
2. Emergency telephone numbers for emergency
services and personnel should be clearly
posted and known to all workers.
3. At least one employee should be trained in
first aid procedures.
4. Specific training for use of hazardous fungal
agents should be available for all personnel.
1
Safety Norms and Regulations in Handling Fungal Specimens
5. A written training plan complemented with
completed employee training records is strongly
recommended for every laboratory to ensure
awareness of routine laboratory practice as well
as safe mycology laboratory practice [18, 21].
Notes
Work surfaces must be disinfected prior to commencing work, after work, and in the instance of
spillages. Various types of disinfectants and in the
case of mycology, fungicides, are available for use
and can be determined as appropriate to the work
being undertaken. In particular, aldehydes such as
formaldehyde and glutaraldehyde, halogens such
as iodine and chlorine compounds, and phenol
derivative proprietary products such as Amphyl,
Lysol, and Vesphene are effective fungicidal agents
[22]. Many other commercial disinfectants are
also available; however, their product information sheet and material safety data sheet (MSDS)
must be consulted to ensure appropriate use.
References
1. Gilchrist MJR, Fleming DO (2000) Biosafety precautions for Mycobacterium tuberculosis and other airborne pathogens. In: Fleming DO, Hunt DL (eds)
Biological safety: principles and practices. ASM
Press, Washington, DC, pp 209–221
2. Pike RM (1976) Laboratory-associated infections:
summary and analysis of 3921 cases. Health Lab Sci
13:105–114
3. Pike RM (1978) Past and present hazards of working
with infectious agents. Arch Pathol Lab Med 102:
333–336
4. Sewell D (1995) Laboratory-associated infections and
biosafety. Clin Microbiol Rev 8(3):389–405
5. Baum GL, Lerner PI (1970) Primary pulmonary blastomycosis: a laboratory-acquired infection. Ann Intern
Med 73(2):263–265
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6. Denton JF, Di Salvo AF, Hirsch ML (1967)
Laboratory-acquired North American blastomycosis.
JAMA 199(12):935–936
7. Graham WR Jr, Callaway JL (1982) Primary inoculation blastomycosis in a veterinarian. J Am Acad
Dermatol 7(6):785–786
8. Larsh HW, Schwarz J (1977) Accidental inoculation
blasmycosis. Cutis 19:334–336
9. Smith DT, Harrell ER (1948) Fatal coccidioidomycosis—a case of a laboratory infection. Am Rev Tuberc
57(4):368–374
10. Wilson JW, Smith CE, Plunkett OA (1953) Primary
cutaneous coccidioidomycosis; the criteria for diagnosis and a report of a case. Calif Med 79(3):
233–239
11. Fiese MJ (1958) Coccidioidomycosis. CC Thomas,
Illinois
12. Nabarro JD (1948) Primary pulmonary coccidioidomycosis; case of laboratory infection in England.
Lancet I:982–984
13. Murray JF, Howard D (1964) Laboratory-acquired
histoplasmosis. Am Rev Respir Dis 89(5):631–640
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histoplasmosis. N Engl J Med 275(11):597–599
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Primary cutaneous histoplasmosis—report of case.
Ann Intern Med 114(1):118–119
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Infect Dis 49(1):142–147
17. Warnock DW (2000) Mycotic agents of human disease. In: Fleming DO, Hunt DL (eds) Biological safety:
principles and practices. ASM Press, Washington, pp
111–120
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guidelines for working with pathogenic and infectious
microorganisms. Current protocols in microbiology.
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Biosafety Manual. WHO, Geneva. [cited 2011 6th
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bacteriology and mycology, 6th edn. Wiley-Blackwell,
Ames, Iowa
2
Methods of Cryopreservation
in Fungi
Ladislav Homolka
Abstract
Traditional method of the routine subculturing by transfer of fungal
cultures from staled to fresh media is not a very practical means of storing
large numbers of fungal cultures. It is time-consuming, prone to contamination, and does not prevent genetic and physiological changes. At present, besides freeze-drying (lyophilization), cryopreservation seems to be
the best preservation technique available for fungi.
Keywords
Cryopreservation • Fungi collections • Liquid nitrogen • Perlite
• Subculturing • Fungi storage
Introduction
Serious mycological (and generally biological)
work requires a reliable source of cultures (i.e.,
well-defined and taxonomically determined starting material), which is ensured by its safe longterm storage. This implies the fundamental and
growing importance of culture collections not
only for preservation of the endangered genofond (and consequently the biodiversity), but
also as a principal source of material for biotechnological processes, research, and teaching. The
L. Homolka (*)
Department of Ecology of Microorganisms,
Institute of Microbiology, Academy
of Sciences of the Czech Republic, Vídeňská 1083,
Prague 4 142 20, Czech Republic
e-mail: homolka@biomed.cas.cz
first and most important problem to be solved is
the long-term maintenance of this material.
Collections of fungi were originally kept by
serial transfers from staled to fresh media. This
routine subculturing is not a very practical method
for storing large numbers of fungal cultures. It is
time-consuming, prone to contamination, and
does not prevent genetic and physiological
changes (degeneration, aging) during long-term
and frequent subculturing [1]. Over the years,
various storage methods have been developed in
order to eliminate these disadvantages. Their
common feature is at least partial suppression of
growth and metabolism of the cultures. Among
them, keeping fungal cultures in sterile water
[2–8] was surprisingly efficient (especially with
lower fungi, but also with some basidiomycetes)
and experiences its revival. In some fungi, preservation under a layer of mineral oil, in silica
gel, soil, or sand [9–13] was successful. These
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_2, © Springer Science+Business Media, LLC 2013
9
10
methods enabled a reduction in fungal growth
and extended the time intervals between transfers
to fresh media. Nevertheless, the method of serial
subculturing is still used in collections with limited financial support or in the majority of other
collections as a backup preservation method.
Searching for improved or new methods
resulted through different intermediate steps in
the introduction of lyophilization and cryopreservation of fungal cultures [14]. Application of the
most extended method of culture preservation—
freeze-drying (lyophilization), tested for sporulating fungi as early as 1945 [15]—is rather
limited in the case of basidiomycetes and other
fungi nonsporulating in vitro [16, 17]. Most
attempts to revitalize dehydrated hyphae of fungi
have failed, except for some successes [18–20];
nevertheless, the real absence of spores must be
always carefully checked. Despite this, several
attempts have recently been made using modified
protocols [21–23] and the growing interest in this
technique can be seen at present. An important
role played in the whole process, besides the
freeze-drying, is also the freezing rate and the
lyoprotectant used [24, 25]. Freeze-drying of
fungi has several important advantages over all
other maintenance methods. Cultures can be
stored easily in dense packing without any special requirements and need not be revived on agar
slants prior to dispatch. The product is light, inactive, and dry, enabling easy distribution by mail.
The new or modified methods have been frequently used and evaluated [5, 6, 10, 26–31], but
they are not generally applicable. Often a specific
preservation protocol is necessary even for individual strains of the same species. At present,
besides freeze-drying (lyophilization), cryopreservation seems to be the best preservation
technique available for filamentous fungi [14,
26, 32]. A very comprehensive and detailed
overview of the methods and results of the cryopreservation in microorganisms was published
by Hubálek [33].
Cryogenic technique for long-term storage of
large numbers of fungal species was introduced
to ATCC in 1960 and the results have been very
satisfactory [34, 35]. The technique was consecutively introduced to many other prominent
L. Homolka
collections, e.g., CAB International Mycological
Institute [1], etc. In certain collections—e.g.,
IFO (Institute for Fermentation Osaka)—
nonsporulating cultures of basidiomycetes are
stored by cryopreservation at −80°C in electric
freezers [36].
Mycelium and/or spore suspensions with or
without a cryoprotectant in sealed glass ampoules
were originally used for cryopreservation of
filamentous fungi. Later, glass ampoules were
replaced with safer polypropylene cryovials
and/or straws. Agar blocks immersed in an
appropriate cryoprotectant were originally used
as carriers of fungal mycelium for the cryopreservation process [37]. A useful straw technique with agar miniblocks for the preservation
of fungi in liquid nitrogen was developed by
Elliott in 1976 [38] and improved by Stalpers
et al. [39]. Another technique using straws in
cryotubes without a cryoprotectant solution was
described by Hoffmann [40]. A modified
Hoffmann’s technique was compared with the
original agar block one in our paper [29].
Commercial preservation systems with polystyrene beads as carriers were used for cyopreservation of conidia of enthomopathogenic fungi
[41] and of sporulating Aspergillus fumigatus
cultures at −80°C [42]. Porous ceramic beads
were employed for cryopreservation of several
sporulating fungal cultures and for a
Saccharomyces cerevisiae culture at −70°C by
Palágyi et al. [43]. It is symptomatic that these
techniques have not been used for nonsporulating filamentous fungi. In this context it should
be mentioned that as early as 1978 Feltham et al.
described a method of preservation of bacteria
on glass beads at −76°C [44]. Some reports [45–
47] indicate that cryopreservation at −80°C is
suitable for many fungal cultures, including
basidiomycetes. Nevertheless, Leeson et al. [48]
state that to completely stabilize frozen cultures,
the temperature must be sufficiently reduced to
both minimize metabolism and prevent ice crystal formation, which can cause physical damage
during storage. The temperature limit securing
prevention of formation of such ice crystals is
−139°C. This is why at present many culture
collections start to keep their cultures at −150°C
2
Methods of Cryopreservation in Fungi
in ultralow-temperature electric freezers, which
are sometimes equipped with liquid nitrogen
supply.
The cryopreservation process includes freezing and thawing and the protocol of these procedures plays an important role [49, 50]. In
principle, there are two kinds of freezing protocols: a slow (controlled) one and a fast (uncontrolled) one, which both have been used for
cryopreservation of fungi [51, 52]. Generally,
too low freezing rates cause excessive dehydration and concentration of the solution leading to
cell damage; on the contrary, too fast freezing
leads to insufficient dehydration and formation
of abundant ice crystals with lethal consequences.
Nevertheless, different fungal cultures exhibit
different sensitivities to freezing conditions and
to the presence and concentration of cryoprotectants. A freezing rate of 1°C per minute is usually used for cryopreservation of fungi; in the
author’s experiments with sensitive mutant
strains of Agaricus bisporus, the freezing rate
0.5°C per minute was successfully used.1 Lately,
cryomicroscopic methods have been used to
study the process of freezing and thawing of fungal cultures [14, 24, 53]. Successful cryopreservation depends on the cryoprotectant used [54].
At present, dimethylsulfoxid and glycerol are the
most widespread [55]. The method of cryogenic
culture maintenance seems to be mostly successful also in nonsporulating cultures [14, 32, 56,
57]. An overview study on the influence of the
cryopreservation process on survival of taxonomically very broad spectrum of fungi published by Smith [14] showed that there was no
obvious link between taxonomic grouping and
the response of the fungi to freezing and thawing. This was confirmed in our study [31]. Similar
studies were carried out also in edible mushrooms Lentinus edodes [58] and the genus
Pleurotus [59]. Davell and Flygh [60] showed
that even an ectomycorrhizal fungus
Cantharellus cibarius can be successfully cryopreserved when a sufficient number of cryoprotocols is tested. Cryopreservation of spores of
1
Homolka, unpublished results.
11
vesicular–arbuscular mycorrhizal fungi was
described by Douds and Schenck [61].
Beyond survival, another principal requirement for the successful preservation of fungal
strains is maintenance of their genetic and physiological features, such as growth, morphology,
and metabolite production. In our experiments
with some white-rot basidiomycetes, no negative
effect of cryopreservation or the used cryoprotective on production of ligninolytic enzymes was
found [62]. The complete revival of cryopreserved cultures (evaluated mostly by measuring
the colony diameter) is generally still uncertain.
The survival rate varies between 60 and 100%
[21, 58, 59, 63, 64]. Only a few studies of the
genetic stability of cryopreserved fungi have been
performed. Singh et al. [47] confirmed the genetic
stability of 11 cryogenically preserved edible
mushroom strains by comparing random
amplified polymorphic DNA (RAPD) and internal transcribed spacer (ITS) profiles. Using polymerase chain reaction (PCR) fingerprinting, Ryan
et al. [65] checked the genetic stability of several
isolates of Fusarium oxysporum and Metarhizium
anisopliae. Other studies include confirmation of
the genetic stability of Uncinula necator conidia
after storage at −80°C [45] and investigation of
the influence of mid-term cryopreservation at
−80°C on 15 isolates of 10 basidiomycete species, for which the DNA fingerprint patterns were
unchanged [66]. All of these reports were solely
based on fingerprinting methods, which are not
suitable for the detection of minute yet important
changes in the genome, such as point or short
indel mutations. Rather, sequencing approaches
are required to successfully detect these mutations. This approach was used in our recent study
[67]. Considering the above data, there is a continuous need for developing, improving, optimizing, and combining of preservation procedures,
because the present methods are not applicable to
all fungal cultures. Although many of these fungi
can be grown in pure cultures on solid media,
their growth is often attenuated and their morphology and other characteristics changed, which
can result in their complete loss. The number of
characteristics evaluating the success of preservation should be increased.
12
As mentioned previously, cryopreservation,
namely in liquid nitrogen, seems to be the most
reliable, safe, and prospective method of a
long-term maintenance of most fungal species,
especially those not amenable to freeze-drying.
It is probably the only storage technique that
can ensure genomic and phenotypic stability.
But not even the aforementioned cryopreservation method is applicable to preservation of all
fungal cultures in the present form. According
to the literature as well as the author’s personal
experience, especially the maintenance of
basidiomycetes is challenging. Many of these
fungi do not form asexual spores, their dominant life form, the vegetative mycelium, is sensitive to environmental conditions and therefore
not amenable to freeze-drying.
To address these issues, a method of cryopreservation using perlite as a carrier for fungal
mycelia was developed in the author’s laboratory
(perlite protocol or PP) [28] and then successfully verified for 442 basidiomycete strains [30].
Perlite is a unique aluminosilicate volcanic mineral that retains substantial amounts of water that
can be released when needed—a feature that
seems to have a dominant effect on cryopreservation success. The PP can be used for cryopreservation of taxonomically different groups of fungi,
including yeasts [31], and works relatively well
for fungi that cannot survive other routine preservation procedures. Expanded perlite was used as
a solid support in solid-state fermentations [68];
otherwise it is used in many applications, particularly in the construction, horticulture, and other
various industrial fields. It is recommended as an
efficient purifying agent and as a carrier for pesticides, feed concentrates, herbicides, and other
similar applications.
L. Homolka
Materials
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
Perlite Protocol (PP)
The protocol is suitable for maintaining a broad
spectrum of fungal cultures of different origin. It
was verified in several culture collections (e.g.,
in Finland, the Netherlands, USA, Czech
Republic, etc.) with great success and now it is
routinely used there.
Distilled water.
Agar Difco.
Glycerol p.a. Sigma.
Isopropyl
alcohol
100%
Sigma
(alternatively).
Agricultural-grade
perlite
(Agroperlit,
GrowMarket s.r.o., Prague, Czech Republic,
http://www.growmarket.cz/produkt/agroperlit-8l)—particles 1–2 mm.
Dried wort extract Sladovit, Malthouse
Bruntál, Bruntál, Czech Republic, diluted to
a density of 4° Balling scale with distilled
water (further wort in the text); or preferably
MYA medium: Malt-extract Difco 25 g,
Yeast-extract Difco 2 g, Glycerol p.a., Serva
50 g, distilled water ad 1,000 mL. pH adjusted
to 6.5 with 1 M KOH solution.
Ethanol (70%).
Liquid nitrogen (LN).
Nunc CryoTube Vials 1.8 mL screw-capped
(Nalgene/Nunc, Rochester, USA).
Cork borer.
Lancet and small spoon.
Rule for measuring of colony diameter.
Water bath.
Balling
hydrometer
(saccharometer)
(alternatively).
Hot-air sterilizer.
Autoclave.
Refrigerator (about 4°C).
Deep-freezer (−80°C).
Microscope equipped with phase contrast.
Laminar flow box.
Thermostat (incubator) for 24°C.
Container for storing samples in liquid nitrogen (e.g., HARSCO TW-5K container,
Harsco, Camp Hill, USA).
Programmable freezer for controlled freezing of cryovials with mycelia (e.g., IceCube
1800 freezer, SY-LAB Geraete GmbH.,
Neupurkersdorf, Austria); or alternatively
Cryo 1°C Freezing Container “Mr. Frosty”
(Nalgene Labware)2.
All chemicals and devices named can be replaced with
other ones produced by other renowned companies.
2
2
Methods of Cryopreservation in Fungi
Methods
The methods given below describe general
procedures for cryopreservation of fungi using
the perlite protocol (PP).
Strains (Cultures)
The starting cultures are kept on wort agar slants
or dishes (wort 4° Balling, 1.5% agar Difco) at
4°C or other media suitable for the growth of the
strains destined for the procedure (e.g., MYA
medium with 1.5% agar, etc.) and transferred to
the fresh medium every 6 months.
1. Prepare an agar medium, sterilize it in an autoclave (121°C, 20 min.), pour it into sterile
plastic Petri dishes (diameter 100 mm, 30 mL
per dish), and let it cool down in a laminar
flow box.
2. In a laminar flow box cut out an agar plug
(6 mm diameter) from the actively growing
part of a colony on a Petri dish with a cork
borer, place it on a Petri dish with fresh medium
using a sterile lancet and then let the dish incubate for 14 days at 24°C. Then put the dish(es)
into a refrigerator and keep it at about 4°C.
13
on a Petri dish using a cork borer, place it
using a sterile lancet on the surface of perlite
in the cryovial, close the vial firmly and let it
incubate for 14 days at 24°C.
2. Freeze the cryovials with perlite overgrown by
mycelium in a programmable freezer (or alternatively in a “Mr. Frosty” container in a deepfreezer) to −70°C at a freezing rate of 1°C per
minute. Then place them in LN in a container.
3. Take the stored frozen cultures in cryovials out
of the LN container, transfer them to a water
bath (37°C), and leave them there until the ice
is completely thawed (thawing—reactivation
of cultures). Prior to opening, disinfect the surface of cryovials with 70% ethanol.
Viability Test
1. After thawing, separate at least partially the
perlite particles overgrown with mycelium by
shaking, the content of the cryovials (two parallels of each strain) divide into three approximately equal aliquots each and these plate onto
wort (or an other) agar medium in Petri dishes
(diameter 100 mm) using a small sterile spoon.
2. Incubate the cultures in Petri dishes at 24°C
for 14 days. Strains exhibiting survival of at
least four out of six separate aliquots are considered viable.
Culture Preparation and Freezing–
Thawing Protocols
Fungal cultures are grown directly in firmly
closed sterile plastic cryovials (1.8 mL) with
200 mg of perlite (Agroperlit, agricultural grade)
moistened with 1 mL of wort (4° Balling) or other
medium (e.g., MYA) enriched with 5% glycerol
as a cryoprotectant. For sterilization of the cork
borer, lancet, and small spoon use a hot-air sterilizer (150°C, 30 min.).
1. Distribute perlite into cryovials (200 mg per
vial), flood it with 1.8 mL of the medium
enriched with 5% of glycerol, and sterilize
vials in autoclave (121°C, 20 min.). In a laminar flow box cut out an agar plug (6 mm diameter) from the actively growing part of a colony
Growth Estimation and Morphological
Analysis
1. Growth of cultures measure as a mean diameter increase of a growth-covered zone (in mm)
during a 14-day incubation at 24°C on the
respective agar medium in Petri dishes (diameter 100 mm) inoculated with perlite aliquots
from cryovials before freezing and after reactivation. Measure six zones (three aliquots
from two cryovials) for each strain. The first
occurrence of growth varies between frozen
cultures, with some strains showing signs of
re-growth within 2 days but most strains reactivating within 7 days after plating.
14
2. Use the same procedure except for freezing
and thawing for growth measurement of the
control.
3. Carry out the morphological analysis on control cultures and on those arising from the
viability tests. Check the selected macroscopic
features (colony color, reverse color, texture
of the mycelium) and microscopic features
(hyphal branching, presence/absence of clamp
connections, presence/absence of hyphal vacuolization, etc.) using a microscope.
4. If possible, estimate also other characteristics
of the resulting cultures (e.g., enzyme or
metabolite production, etc.) according to your
consideration.
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46. Kitamoto Y, Suzuki A, Shimada S, Yamanaka K (2002)
A new method for the preservation of fungus stock cultures by deepfreezing. Mycoscience 43:143–149
47. Singh SK, Upadhyay RC, Kamal S, Tiwari M (2004)
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yield and genetic stability. CryoLetters 25:23–32
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3
Long-Term Preservation of Fungal
Cultures in All-Russian Collection
of Microorganisms (VKM): Protocols
and Results
Svetlana M. Ozerskaya, Natalya E. Ivanushkina,
Galina A. Kochkina, Svetlana S. Eremina,
Alexander N. Vasilenko, and Nadezhda I. Chigineva
Abstract
Results of successful preservation experience are given for the taxonomic
groups of fungi preserved in All-Russian Collection of Microorganisms
(VKM): the species names, conservation methods, storage time
estimates.
Keywords
Fungi • Culture collection • Storage time • Survival • Lyophilization •
Freeze-drying • Cryopreservation • Sterile soil
Introduction
Preservation and long-term storage of type,
authentic, and other kinds of fungal cultures in a
living state is of high importance both for the fundamental and practical mycology.
Long-time storage of strains is performed in
microbiological culture collections (biological
resource centers). Various methods of preservation of fungal cultures have been reported [1–3].
Freeze-drying (lyophilization) and cryopreservation methods are utilized for thousands of fungal strains in microbial collections all over the
S.M. Ozerskaya () • N.E. Ivanushkina • G.A. Kochkina
S.S. Eremina • A.N. Vasilenko • N.I. Chigineva
G. K. Skryabin Institute of Biochemistry
and Physiology of Microorganisms, Russian Academy
of Science, prospekt Nauki, 5, Pushchino,
Moscow Region 142290, Russia
e-mail: smo@dol.ru
world [4–6]. Nevertheless, it is clear that the fungal
strains of different species vary in ability to survive
after the long-time storage preservation under laboratory conditions. Some of them are very difficult
to maintain ex situ, whereas others could be easily
and successfully preserved alive by using almost
any conservation technique. Available information
on the maximal time periods in which the reliable
storage of different fungal species are ensured does
not cover those for the diversity of fungi maintained
in culture collections. This chapter presents the
methods of cryopreservation, freeze-drying, and
preservation in sterile soil that are utilized in VKM
fungal collection, accompanied by data on maximal storage time registered. The methods take into
consideration the special features of cultures preserved as well as the equipment used.
VKM fungal collection (All-Russian
Collection of Microorganisms, Russia) was
established in 1955 and has a long-term experience for preservation and storage of fungal
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_3, © Springer Science+Business Media, LLC 2013
17
18
cultures. Collection of filamentous fungi is currently composed of approximately 5,000 strains
(545 genera, 1,450 species) belonging to species
of the kingdoms Chromista (Oomycetes) and
Fungi
(Zygomycetes,
Ascomycetes,
and
Basidiomycetes). The current use of different
preservation methods for more than 3,800 strains
maintained in VKM for more than 40 years was
analyzed using a specially designed database.
The database keeps the protocols of conservation
methods, storage conditions, the calculated time
of reliable storage, special requirements of
growth, and other information related to the issue.
Data presented in this chapter are derived from
this database. The information on preservation
methods is also available in VKM catalogue
(http://www.vkm.ru/Catalogue.htm), in data
sheet for each strain.
Cryopreservation of Filamentous
Fungi
According to published data, the fast cooling
rates followed by storage in liquid nitrogen at
−196°C allow secure and long-term preservation
of some fungal cultures [7]. However, the ability
to resist damage by freezing and warming differs
considerably among genera/species and depends
on their particular features (presence and type of
sporulation, chemical composition of cytoplasmic membrane and cell wall, physiological state,
etc.). Selection of optimal cryoprotectants, rates
of cooling and warming has enabled increasing
the number and diversity of taxa preserved by
this method [8, 9].
More than 70% filamentous fungal of VKM
(2,714 strains belonging to 1,148 species and 405
genera) are stored using various cryopreservation
protocols. Cultures with abundant sexual and
nonsexual sporulation usually were preserved by
using fast cooling rates followed by storage either
in liquid nitrogen or in ultra-low temperature
freezers at −80°C.
It was noticed that some cultures of
Zygomycetes belonging to the genera Mortierella,
Basidiobolus, Coemansia, and Lobosporangium
S.M. Ozerskaya et al.
(syn. Echinosporangium) do not survive the ultrarapid freezing procedure even if they have abundant sporulation. Successful preservation of such
strains was achieved by modification of the cryopreservation regime, for example using slow programmed freezing. The same method was used
either for nonsporulating fungi or zoosporeforming ones (Basidiomycetes, Oomycetes).
According to our data, some part of strains of
Oomycetes (20%), Basidiomycetes (4%),
Zygomycetes (1%), and Ascomycetes (1%) did
not survive cryopreservation at all freezing
regimes and modification applied [10]. The
strains most difficult to maintain belong to genera Brevilegnia, Dictyuchus, Phytophthora, and
to some species of Achlya and Saprolegnia.
Similar situations have also been seen with some
species of Basidiomycetes (Suillus, Amanita,
Dictyophora, Mutinus, etc.). They are usually
maintained by subculturing and preservation
under mineral oil.
It has been suggested that those microbial
cultures that are able to survive the freezing and
a short storage will permanently stay in the vital
state after any length of storage [11]. According
to our data this is not quite true: some strains of
Achlya colorata, Antrodia serialis, Armillaria
cepistipes, Athelia rolfsii, Ceratellopsis equiseticola, Choanephora conjuncta, Clitocybe
nuda, Coemansia aciculifera, Collybia butyracea, Conidiobolus thromboides, Exobasidium
karstenii, E. splendidum, Kickxella alabastrina,
Lactarius deliciosus, Marasmius oreades,
Mortierella gamsii, M. humilis, Mycena pura,
Phallus impudicus, Rhizoctonia solani,
Sclerotium tuliparum, Suillus variegatus, and
Ustilago scabiosae have lost their ability to
grow after 5–7 years of storage in liquid nitrogen, although they were in the vital state after
24 h of storage. The reason is not yet known.
Nevertheless, the viability test showed that 350
strains of fungi remain alive after 19.5 years of
storage (Table 3.1).
The cooling equipment being used in VKM is
storage tanks “Bioproducts-0.5” with capacity of
500 liters of liquid nitrogen and ultra-low temperature freezers (−80°C, Sanyo, Japan).
Name of species
Absidia blakesleeana Lendner 1924
A. coerulea Bainier 1889
A. cuneospora G.F.Orr et Plunkett 1959
A. cylindrospora Hagem 1908
A. glauca Hagem 1908
A. hyalospora (Saito 1906) Lendner 1908
A. repens van Tieghem 1878
A. spinosa Lendner 1907
A. bisexualis Coker et Couch 1927
Achlya bonariensis Beroqui 1969
A. colorata Pringsheim 1882
A. intricata Beneke 1948
A. sparrowii Reischer 1949
Acladium curvatum Bonorden 1851
Acremonium alternatum Link 1809
A. arxii W.Gams 1971
A. atrogriseum (Panasenko 1964) W.Gams 1971
A. bacillisporum (Onions et G.L. Barron 1967) W.Gams 1971
A. bactrocephalum W.Gams 1971
A. berkeleyanum (P.Karsten 1891) W.Gams 1982
A. biseptum W.Gams 1971
A. breve (Sukapure et Thirumalachar 1966) W.Gams 1971
A. cavaraeanum (Jasevoli 1924) W.Gams 1971
A. charticola (J.Lindau 1907) W.Gams 1971
A. crotocinigenum (Schol-Schwarz 1965) W.Gams 1971
A. cymosum W.Gams 1971
A. domschii W.Gams 1971
A. egyptiacum (J.F.H.Beyma 1933) W.Gams 1971
Cryopreservation
Number
Storage
of strains
time (years)
1
19.30
2
19.69
1
19.69
4
1
1
1
2
1
2
1
1
19.37
19.37
19.24
12.28
0.51
0.16
6.33
0.15
1.11
1
1
1
0.58
19.44
17.46
1
19.49
3
1
2
19.29
19.92
19.56
1
6.52
1
19.77
Freeze-Drying
Number
of strains
4
5
1
2
10
1
1
2
Storage
time (years)
32.91
29.67
27.38
24.02
36.13
31.10
22.36
19.46
1
2
3
2
1
3
3
1
4
1
3
4
1
2
1
32.40
27.04
27.32
32.65
15.50
24.26
30.12
25.30
27.78
6.05
25.98
32.50
28.44
28.36
29.72
Soil
Number
of strains
3
5
0
2
5
1
2
Storage
time (years)
27.14
12.42
16.24
28.45
15.58
11.64
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
Sr. No.
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
3
Table 3.1 Storage time of VKM fungal cultures
(continued)
19
20
Table 3.1 (continued)
Sr. No.
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
51
52
53
54
55
2
1
3
19.75
17.71
20.27
2
2
1
19.36
19.92
8.01
1
5.09
14
19.79
1
1
2
1
17.48
19.58
19.34
19.28
2
19
1
20.03
20.93
19.84
1
1
1
1
1
19.28
19.76
19.27
19.78
19.76
Freeze-Drying
Number
of strains
1
1
1
3
1
4
1
2
3
3
3
3
Storage
time (years)
7.06
31.29
3.23
27.42
25.80
26.11
24.26
30.12
25.34
19.10
2.70
24.26
20
1
1
1
8
1
7
39.59
6.25
6.36
18.18
27.50
22.40
35.32
1
13.21
11
29.77
2
1
1
24.42
16.13
24.95
Soil
Number
of strains
Storage
time (years)
1
9.71
3
13.56
7
28.66
1
7.45
S.M. Ozerskaya et al.
48
49
50
Name of species
A. gamsii Tichelaar 1971
A. hennebertii W.Gams 1971
A. hyalinulum (Saccardo 1879) W.Gams 1971
A. implicatum (J.C.Gilman et E.V.Abbott 1927) W.Gams 1975
A. incrustatum W.Gams 1971
A. kiliense Gruetz 1925
A. lichenicola W.Gams 1971
A. persicinum (Nicot 1958) W.Gams 1971
A. polychromum (J.F.H.Beyma 1928) W.Gams 1971
A. rutilum W.Gams 1971
A. salmoneum W.Gams et Lodha 1975
A. sclerotigenum (Moreau et R.Moreau 1941 ex Valenta 1948)
W.Gams 1971
A. strictum W.Gams 1971
A. tubakii W. Gams 1971
Acrophialophora fusispora (S.B.Saksena 1953) Samson 1970
Acrostalagmus albus Preuss 1851
A. luteoalbus (Link 1809) Zare et al. 2004
Acrothecium robustum J.C.Gilman et E.V.Abbott 1927
Actinomucor elegans (Eidam 1884) C.R.Benjamin et
Hesseltine 1957
Agaricus arvensis Schaeffer 1774
A. bisporus (J.Lange 1926) Imbach 1946
Albonectria rigidiuscula (Berkeley et Broome 1875)
Rossman et Samuels 1999
Alternaria alternata (Fries 1832) Keissler 1912
A. brassicae (Berkeley 1836) Saccardo 1880
A. brassicicola (Schweinitz 1832) Wiltshire 1947
A. cheiranthi (Libert 1827) P.C.Bolle 1924
A. chrysanthemi E.G.Simmons et Crosier 1965
Cryopreservation
Number
Storage
of strains
time (years)
1
19.78
1
1
1
1
1
1
1
1
1
1
19.78
19.76
12.56
12.56
12.56
17.79
19.76
17.59
19.30
19.76
1
1
1
1
1
1
1
1
1
1
1
1
3
1
6.18
15.33
19.52
12.57
12.18
1
1
1
1
1
15.93
19.19
12.44
12.01
0.79
12.28
12.57
12.37
12.57
1.64
20.37
17.71
9.93
2
1
14.87
11.28
1
1
1
2
1
1
1
1
1
1
2
2
1
1
22.55
27.64
11.82
8.64
17.43
31.28
5.32
11.42
13.21
16.55
13.12
18.13
18.54
10.41
1
3
20.62
24.20
1
1.63
1
1
20.57
20.62
1
12.18
21
(continued)
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
57
58
59
60
61
62
63
64
65
66
67
68
69
70
71
72
73
74
75
76
77
78
79
80
81
82
83
84
85
86
87
A. cucumerina (Ellis et Everhart 1895) J.A.Elliott 1917
var. cucumerina
A. dauci (J.G.Kuehn 1855) J.W.Groves et Skolko 1944
A. dianthicola Neergaard 1945
A. geophila Daszewska 1912
A. godetiae (Neergaard 1933) Neergaard 1945
A. macrospora Zimmermann 1904
A. multirostrata E.G.Simmons et C.R.Jackson 1968
A. nobilis (Vize 1877) E.G. Simmons 2002
A. radicina Meier et al. 1922
A. raphani J.W.Groves et Skolko 1944
A. solani Sorauer 1896
A. tenuissima (Kunze 1818) Wiltshire 1933
Amauroascus aureus (Eidam 1887) von Arx 1971
Amblyosporium botrytis Fresenius 1863
Amerosporium concinnum Petrak 1953
Ampelomyces artemisiae (Voglino 1905) Rudakov 1979
A. heraclei (Dejeva 1967) Rudakov 1979
A. humuli (Fautrey 1890) Rudakov 1979
A. polygoni (Potebnia 1907) Rudakov 1979
A. quisqualis Cesati 1852
A. ulicis (Adams 1907) Rudakov 1979
A. uncinulae (Fautrey 1893) Rudakov 1979
Anthurus archeri (Berkeley 1859) E. Fisch. 1886
Aphanoascus fulvescens (Cooke 1879) Apinis 1968
Aphanocladium album (Preuss 1848) W.Gams 1971
Aphanomyces helicoides Minden 1915
Apiospora montagnei Saccardo 1875
Aplanes treleaseanus (Humphrey 1893) Coker 1927
Aposphaeria caespitosa (Fuckel 1869) Jaczewski 1917
Arachniotus aurantiacus (Kamyschko 1967) von Arx 1971
Armillaria bulbosa (Barla 1887) Kill et Watling 1983
A. gallica H.Marxmueller et Romagn. 1987
3
56
Sr. No.
A. mellea (Vahl 1792) Kummer 1871
Arthrinium arundinis (Corda 1838) Dyko et Sutton 1981
A. saccaricola F. Stevens 1917
A. sphaerospermum Fuckel 1874
Arthrobotrys arthrobotryoides (Berlese 1888) J.Lindau 1907
A. cladodes Drechsler 1937
A. conoides Drechsler 1937
A. longa Mekhtieva 1973
A. longispora Press 1853
A. oligospora Fresenius 1850
A. robusta Duddington 1951
A. superba Corda 1839
Ascochyta bohemica Kabat et Bubak 1905
A. boltshauseri Saccardo 1891
A. cucumeris Fautrey et Roumeguere 1891
A. malvicola Saccardo 1878
A. pinodes L.K.Jones 1927
A. pisi Libert 1830
A. viciae Libert 1837
Ascotricha chartarum Berkeley 1838
Aspergillus alliaceus Thom et Church 1926
A. amylovorus Panasenko 1964 ex Samson 1979
A. aureolatus Muntanola-Cvetkovic et Bata 1964
A. awamori Nakazawa 1915
A. awamori Nakazawa 1915 var. fumeus Nakazawa et al. 1936
A. brasiliensis Varga et al. 2007
A. caespitosus Raper et Thom 1944
A. candidus Link 1809
A. carbonarius (Bainier 1880) Thom 1916
A. carneus Blochwitz 1933
Cryopreservation
Number
Storage
of strains
time (years)
3
1
19.93
12.57
1
1
1
2
1
1
4
1
1
1
1
1
1
1
1
1
1
19.19
19.30
15.73
19.30
10.64
15.63
21.21
19.40
19.25
20.32
20.32
20.32
19.21
19.21
19.44
20.30
18.95
1
12.45
Freeze-Drying
Number
Storage
of strains
time (years)
3
1
1
10.13
16.91
20.82
1
1
1
23.73
9.44
9.42
5
1
1
22.24
27.06
18.25
1
1
27.54
27.91
1
1
1
1
3
1
1
6
1
1
1
5
2
1
22.41
23.61
26.76
10.75
21.91
14.96
9.61
28.56
17.96
31.96
9.66
38.93
20.02
16.12
Soil
Number
of strains
Storage
time (years)
1
1
1
2
3.85
3.91
3.91
3.91
1
1
1
2
3.91
2.55
3.91
3.91
1
8.67
2
1
1
4
1
1
1
5
1
20.77
20.73
9.58
20.73
20.91
22.13
9.58
20.87
20.87
S.M. Ozerskaya et al.
88
89
90
91
92
93
94
95
96
97
98
99
100
101
102
103
104
105
106
107
108
109
110
111
112
113
114
115
116
117
Name of species
22
Table 3.1 (continued)
1
12.45
7
1
1
4
4
13
1
1
12
2
1
1
1
4
2
3
26
3
1
8
21
1
30.13
9.60
13.22
27.20
31.64
39.07
26.98
14.18
38.91
16.87
15.87
15.90
36.04
24.93
39.62
39.08
27.30
28.54
37.39
24.89
30.35
12.34
7
1
2
3
4
9
1
1
1
1
2
2
1
18
2
1
7
19
1
11.41
20.92
20.66
16.06
1.91
27.41
37.78
11.74
20.89
23.11
9.58
1
1
3
1
1
1
9
1
2
1
18.59
37.39
38.97
14.94
36.19
9.87
38.67
15.70
21.13
27.29
1
1
2
1
21.90
4.65
21.72
7.66
1
8
1
1
1
4.32
27.41
2.24
2.05
8.08
(continued)
23
2
27.41
9.58
1.53
27.41
20.90
20.87
8.69
8.08
28.72
17.90
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
140
141
142
143
144
145
146
147
148
149
A. clavatus Desmazieres 1834
A. echinulatus (Delacroix 1893) Thom et Church 1926
A. ficuum (Reichardt 1867) Thom et Currie 1916
A. fischeri Wehmer 1907
A. flavipes (Bainier et R.Sartory 1911) Thom et Church 1926
A. flavus Link 1809
A. flavus Link 1809 var. columnaris Raper et Fennell 1965
A. foetidus Thom et Raper 1945
A. fumigatus Fresenius 1863
A. giganteus Wehmer 1901
A. heteromorphus Batista et H. Maia 1957
A. insuetus (Bainier 1908) Thom et Church 1929
A. janus Raper et Thom 1944
A. japonicus Saito 1906
A. kanagawaensis Nehira 1951
A. melleus Yukawa 1911
A. niger van Tieghem 1867
A. niveus Blochwitz 1929
A. nutans McLennan et Ducker 1954
A. ochraceus G.Wilhelm 1877
A. oryzae (Ahlburg 1878) E.Cohn 1884
A. oryzae (Ahlburg 1878) E.Cohn 1884 var. effusus
(Tiraboschi 1908) Y.Ohara 1951
A. pallidus Kamyschko 1963
A. parvulus G.Smith 1961
A. penicilliformis Kamyschko 1963
A. phoenicis (Corda 1840) Thom et Currie 1916
A. pseudodeflectus Samson et Mouchacca 1975
A. puniceus Kwon-Chung et Fennell 1965
A. repens (Corda 1842) Saccardo 1882
A. restrictus G. Smith 1931
A. sclerotiorum G.A.Huber 1933
A. silvaticus Fennell et Raper 1955
3
118
119
120
121
122
123
124
125
126
127
128
129
130
131
132
133
134
135
136
137
138
139
24
Table 3.1 (continued)
Sr. No.
150
151
152
153
154
155
156
157
158
159
160
161
162
163
164
165
166
167
168
A. subsessilis Raper et Fennell 1965
A. sulphureus (Fresenius 1863) Thom et Church 1926
A. sydowii (Bainier et R.Sartory 1913) Thom et Church 1926
A. tamarii Kita 1913
A. terreus Thom 1918
A. terreus Thom 1918 var. africanus Fennell et Raper 1955
A. terreus Thom 1918 var. aureus Thom 1918 Thom et
Raper 1945
A. terricola Marchal et É.J. Marchal 1893
A. terricola Marchal et É.J. Marchal 1893 var. americanus
Marchal et É.J. Marchal 1921
A. tubingensis Mosseray 1934
A. umbrosus Bainier et Sartory 1912
A. unguis (Weill et L.Gaudin 1919) Thom et Raper 1939
A. ustus (Bainier 1881) Thom et Church 1926
A. varians Wehmer 1897
A. versicolor (Vuillemin 1903) Tiraboschi 1908
A. wentii Wehmer 1896
Athelia rolfsii (Curzi 1932) C.C.Tu et Kimbrough 1978
Aureobasidium microstictum (Bubak 1907) W.B.Cooke 1962
A. pullulans (de Bary 1866) G.Arnaud 1918 var. melanigenum
Hermanides-Nijhof 1977
A. pullulans (de Bary 1866) G.Arnaud 1918 var. pullulans
Backusella circina J.J. Ellis et Hesseltine 1969
B. lamprospora (Lendner 1908) Benny et R.K.Benjamin 1975
Bactridium equiseticola Milko et Dunaev
Basidiobolus magnus Drechsler 1964
B. meristosporus Drechsler 1955
Beauveria bassiana (Balsamo-Crivelli 1835) Vuillemin 1912
B. brongniartii (Saccardo 1892) Petch 1924
Freeze-Drying
Number
Storage
of strains
time (years)
Soil
Number
of strains
Storage
time (years)
1
12.43
1
1
6.67
1
1
6
1
17
1
1
24.42
17.11
25.86
38.72
35.75
14.73
14.73
1
1
4
1
15
1
1
10.82
22.65
21.01
20.73
34.79
7.74
7.22
3
1
17.41
21.47
3
1
20.92
7.74
1
3
5
11
1
11
3
7.19
38.97
22.22
24.75
18.57
29.25
26.56
1
2
5
6
21.03
16.58
20.89
11
3
22.34
7.69
1
6
26.42
27.81
2
12.60
10
2
4
1
31.03
31.44
39.34
10.24
1
3.76
2
8.78
6
5
30.68
31.16
1
10.92
1
12.45
1
1
1
8.63
19.84
6.76
1
20.20
2
19.63
1
1
3
4
20.05
20.20
20.39
20.39
S.M. Ozerskaya et al.
169
170
171
172
173
174
175
176
Name of species
Cryopreservation
Number
Storage
of strains
time (years)
195
196
197
198
199
200
201
202
203
204
205
206
1
19.54
2
1
16.91
26.61
2
24.89
2
19.28
8.53
19.11
1
8.43
1
19.34
1
1
2
1
1
2
8
2
1
17.46
19.31
19.15
17.44
20.20
8.31
19.56
8.60
19.96
19.06
6.61
25.93
7.04
5.45
19.77
5.40
16.28
0.66
27.07
8.96
15.09
2
1
5
1
3
1
1
3
1
1
1
2
2
1
17
3
1
40.03
8.36
26.19
4
11.74
1
18.95
1
19.21
1
1
19.29
19.19
1
19.39
1
1
1
4
2
2
1
1
15
2
2.01
5.39
19.35
27.64
28.41
17.59
22.52
6.88
22.56
3.56
1
1
19.21
19.39
2
17.62
25
(continued)
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
179
180
181
182
183
184
185
186
187
188
189
190
191
192
193
194
Benjaminiella poitrasii (R.K.Benjamin 1960) von Arx 1981
Bionectria ochroleuca (Schweinitz 1832) Schroers et
Samuels 1997
Bipolaris australiensis (M.B.Ellis 1971) Tsuda et Ueyama 1981
B. bicolor (Mitra 1931) Shoemaker 1959
B. cynodontis (Marignoni 1909) Shoemaker 1959
B. kusanoi (Y.Nisikado 1928) Shoemaker 1959
B. nodulosa (Berkeley et M.A.Curtis 1886) Shoemaker 1959
B. sorokiniana (Saccardo 1890) Shoemaker 1959
B. spicifera (Bainier 1908) Subramanian 1971
B. victoriae (F. Meehan et H.C. Murphy 1946) Shoemaker 1959
Biscogniauxia nummularia (Bulliard 1790) Kuntze 1891
Bispora antennata (Persoon 1801) E.W. Mason 1953
B. betulina (Corda 1838) S.Hughes 1958
B. effusa Peck 1891
Bjerkandera adusta (Willdenow 1787) P.Karsten 1879
Blakeslea trispora Thaxter 1914
Blumeriella jaapii (Rehm 1907) Arx 1961
Botryosphaeria rhodina (Berkeley et M.A.Curtis 1889)
von Arx 1970
Botryosporium longibrachiatum (Oudemans 1890) Maire 1903
Botryotinia narcissicola (P.H.Gregory 1941) N.F.Buchwald 1949
B. polyblastis (P.H.Gregory 1938) N.F.Buchwald 1949
Botryotrichum piluliferum Saccardo et Marchal 1885
Botryoxylon geniculatum (Corda 1839) Ciferri 1962
Botrytis aclada Fresenius 1850
B. anthophila Bondartsev 1913
B. bifurcata J.H. Mill., Giddens et A.A. Foster 1958
B. cinerea Persoon 1794
B. convallariae (Klebahn 1930) Ondrej 1972 ex Boerema et
Hamers 1988
B. convoluta Whetzel et Drayton 1932
B. elliptica (Berkeley 1881) Cooke 1901
3
177
178
Sr. No.
B. fabae Sardina 1929
B. galanthina (Berkeley et Broome 1873) Saccardo 1886
B. gladiolorum Timmermans 1941
B. hyacinthi Westerdijk et J.F.H.Beyma 1928
B. lutescens Saccardo et Roumeguere 1882
B. squamosa J.C.Walker 1925
B. tulipae (Libert 1830) Lind 1913
Brachysporium nigrum (Link 1824) S. Hughes 1958
Burgoa anomala (Hotson 1912) Goidanich 1937
Byssochlamys nivea Westling 1909
Cadophora fastigiata Lagerberg et Melin 1928
C. malorum (Kidd et Beaumont 1924) W. Gams 2000
C. melinii Nannfeldt 1934
Calcarisporium arbuscula Preuss 1851
C. griseum Spegazzini 1902
Calocera viscosa (Persoon 1794) Fries 1828
Ceratellopsis aquiseticola (Boudier 1917) Corner 1950
Ceratocystis paradoxa (Dade 1928) C.Moreau 1952
C. pilifera (Fries 1822) C.Moreau 1952
Cercospora armoraciae Saccardo 1876
C. beticola Saccardo 1876
C. carotae (Passerini 1887) Kaznowski et Siemaszko 1929
C. rosicola Passerini 1875
C. violae Saccardo 1876
Ceriporiopsis gilvescens (Bresadola 1908) Domanski 1963
Cerrena unicolor (Bulliard 1788) Murrill 1903
Chaetocladium brefeldii van Tieghem et G.Le Monnier 1873
C. jonesii (Berkeley et Broome 1854) Fresenius 1863
Chaetocytostroma sp.
Chaetomidium pilosum (C.Booth et Shipton 1966) von Arx 1975
Cryopreservation
Number
Storage
of strains
time (years)
1
1
1
1
1
1
1
19.11
19.21
19.29
19.19
19.19
19.31
19.48
1
2
1
4
1
1
1
1
1
2
1
1
19.86
19.42
19.39
19.74
19.47
1.93
7.62
0.10
12.06
19.84
18.85
20.32
1
1
20.32
16.65
1
1
1
1
12.57
12.58
17.58
17.58
1
14.44
Freeze-Drying
Number
Storage
of strains
time (years)
1
2
9.62
23.47
1
1
1
2
2
4
1
3
2
13.16
26.08
8.98
30.50
21.47
19.27
22.91
28.45
27.88
2
1
1
2
1
17.43
25.82
11.89
15.29
12.70
1
12.67
2
1
1
1
33.55
27.42
20.53
18.28
Soil
Number
of strains
Storage
time (years)
1
1
22.83
1
6.47
1
16.98
1
3.11
S.M. Ozerskaya et al.
207
208
209
210
211
212
213
214
215
216
217
218
219
220
221
222
223
224
225
226
227
228
229
230
231
232
233
234
235
236
Name of species
26
Table 3.1 (continued)
258
259
260
261
262
263
264
265
266
1
1
2
1
3
19.27
19.27
20.50
19.27
20.50
19.66
1
1
2
1
3
1
1
11
1
2
2
1
1
1
1
1
1
1
1
1
31.01
17.37
15.45
20.34
27.68
9.05
26.15
31.21
10.66
17.66
17.39
35.25
35.29
17.35
11.07
35.29
25.61
11.80
2.01
21.35
1
7
1
1
2
1
1
1
1
1
1
19.27
19.42
19.32
19.27
19.27
18.85
19.27
19.27
19.27
19.27
20.37
1
1
20.06
1
27.36
2
1
1
1
5.81
20.19
19.07
20.09
1
1
1
8.64
20.93
26.29
1
1
2
1
5
19.81
22.10
19.07
31.11
8.59
1
9.56
4
5.61
1
1
11.03
10.68
3
10.68
1
9.02
1
1
5.95
5.87
27
(continued)
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
257
Chaetomium amesii Sergeeva 1965
C. angustispirale Sergeeva 1956
C. aureum Chivers 1912
C. crispatum (Fuckel 1867) Fuckel 1870
C. elatum Kunze 1817
C. fieberi Corda 1837
C. funicola Cooke 1873
C. globosum Kunze 1817
C. homopilatum Omvik 1953
C. indicum Corda 1840
C. megalocarpum Bainier 1910
C. nozdrenkoae Sergeeva 1961
C. perlucidum Sergeeva 1956
C. semenis-citrulli Sergeeva 1956
C. spirale Zopf 1881
C. subaffine Sergeeva 1961
C. subspirilliferum Sergeeva 1960
C. trilaterale Chivers 1912
Chaunopycnis alba W. Gams 1979
Chloridium virescens (Persoon 1797) W.Gams et
Holubova-Jechova 1976 var. caudigerum (Hoehnel 1903)
W.Gams et Holubova-Jechova 1976
Choanephora circinans (H.Naganishi et N.Kawakami 1955)
Hesseltine et C.R.Benjamin 1957
C. conjuncta Couch 1925
C. cucurbitarum (Berkeley et Ravenel 1875) Thaxter 1903
C. infundibulifera (Currey 1873) Saccardo 1891
Chondrostereum purpureum (Persoon 1794) Pouzar 1959
Chrysonilia sitophila (Mont. 1843) Arx 1981
Chrysosporium carmichaelii Oorschot 1980
C. keratinophilum D.Frey 1959 ex J.W.Carmichael 1962
C. lobatum Scharapov 1978
C. lucknowense Garg 1966
3
237
238
239
240
241
242
243
244
245
246
247
248
249
250
251
252
253
254
255
256
28
Table 3.1 (continued)
Sr. No.
267
268
269
270
271
272
273
274
275
276
277
278
C. merdarium (Link 1818 ex Greville 1823) J.W.Carmichael 1962
C. queenslandicum Apinis et R.G.Rees 1976
C. tropicum J.W.Carmichael 1962
Circinella muscae (Sorokin 1870) Berlese et de Toni 1888
C. rigida G.Smith 1951
C. umbellata van Tieghem et G.Le Monnier 1873
Cladobotryum binatum Preuss 1851
C. dendroides (Bulliard 1791) W.Gams et Hoozemans 1970
C. multiseptatum de Hoog 1978
C. varium Nees 1816
C. verticillatum (Link 1809) S. Hughes 1958
Cladophialophora chaetospira (Grove 1886) Crous et
Arzanlou 2007
Cladosporium aecidiicola Thuemen 1876
C. brevicompactum Pidoplichko et Deniak 1941
C. bruhnei Linder 1947
C. cladosporioides (Fresenius 1850) G. A. de Vries 1952
C. colocasiae Sawada 1916
C. cucumerinum Ellis et Arthur 1889
C. elegantulum Pidoplichko et Deniak 1938
C. gossypiicola Pidoplichko et Deniak 1941
C. halotolerans Zalar et al. 2007
C. herbarum (Persoon 1794) Link 1816
C. macrocarpum Preuss 1848
C. sphaerospermum Penzig 1882
C. straminicola Pidoplichko et Deniak 1938
C. transchelii Pidoplichko et Deniak 1938
Clavariadelphus pistillaris (Fries 1753) Donk 1933
Claviceps paspali F.Stevens et J.G.Hall 1910
C. purpurea (Fries 1823) Tulasne 1853
2
1
1
1
1
19.81
9.08
2.81
19.67
19.67
1
1.20
1
19.19
2
1
19.51
19.34
1
1
18.99
19.29
2
3
3
2.16
18.93
18.93
Freeze-Drying
Number
Storage
of strains
time (years)
2
2
1
5
1
1
1
3
1
3
2
1
25.63
31.15
24.13
28.07
26.96
6.91
29.01
25.88
9.00
27.38
28.41
17.82
1
2
1
15
1
1
2
2
1
31
3
7
1
1
5.95
27.73
9.30
37.72
18.35
18.62
26.47
34.83
13.43
36.58
17.54
26.65
26.05
13.10
1
4.55
Soil
Number
of strains
Storage
time (years)
1
1
5.87
6.07
4
1
1
28.66
16.39
19.35
3
6.72
1
7.58
8
5.76
2
14.36
S.M. Ozerskaya et al.
279
280
281
282
283
284
285
286
287
288
289
290
291
292
293
294
295
Name of species
Cryopreservation
Number
Storage
of strains
time (years)
305
306
307
308
309
310
311
312
313
314
315
316
317
318
319
320
321
322
323
324
325
1
14
0.01
19.85
1
1
2
1
18.89
8.85
20.14
16.86
1
19.32
1
1
19.30
2.82
4
1
1
15.93
6.78
18.95
4
1
12.01
18.99
1
1
1
1
2
1
1
1
2
1
1
1
17.68
19.39
18.99
6.11
12.05
3.82
20.14
18.86
18.82
18.82
18.86
20.37
23
37.79
3
32.42
3
1
3
2
1
27.13
9.68
6.41
37.54
9.42
1
24.90
1
10.61
1
14.79
1
1
1
2
1
12.63
9.65
16.15
16.28
16.61
1
39.91
16
19.96
1
22.74
29
(continued)
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
298
299
300
301
302
303
304
Clitocybe odora (Bulliard 1784) P.Kummer 1871
Clonostachys rosea (Link 1816) Schroers, Samuels,
Seifert et W.Gams 1999
C. solani (Harting 1846) Schroers et W.Gams 2001
Coemansia aciculifera Linder 1943
Cokeromyces recurvatus Poitras 1950
Colletoconis aecidiophila (Spegazzini 1886) de Hoog et al. 1978
Colletotrichum coccodes (Wallroth 1833) S.Hughes 1958
C. gloeosporioides (Penzig 1880) Saccardo 1882
C. lindemuthianum (Saccardo et Magnus 1878) Briosi et
Cavara 1889
C. musae (Berkeley et M.A.Curtis 1874) Arx 1957
Collybia butyracea (Bulliard 1792) P.Kummer 1871
Colpoma quercinum (Persoon 1796) Wallroth 1823
Conidiobolus coronatus (Costantin 1897) Batko 1964
C. thromboides Drechsler 1953
Coniochaeta verticillata (van Emden 1973) Dania García,
Stchigel et Guarro 2006
Coniophora puteana (Schumacher 1803) P.Karsten 1868
Coniothyrium concentricum (Desmazieres 1840) Saccardo 1878
C. fuckelii Saccardo 1878
C. hellebori Cooke et Massee 1886
C. rosarum Cooke et Harkness 1882
C. wernsdorffiae Laubert 1905
Coprinus atramentarius (Bulliard 1783) Fries 1838
C. comatus (O.F. Mueller 1780) Persoon 1797
C. disseminatus (Persoon 1801) Gray 1821
C. domesticus (Bolton 1788) Gray 1821
C. ephemerus (Bulliard 1786) Fries 1838
C. micaceus (Bulliard 1785) Fries 1838
C. radians (Desmazieres 1828) Fries 1838
C. sterquilinus (Fries 1821) Fries 1838
Corynascus sepedonium (C.W.Emmons 1932) von Arx 1973
3
296
297
Sr. No.
326
327
328
329
330
331
332
333
334
335
336
337
338
339
340
341
342
343
Cryphonectria parasitica (Murrill 1906) M.E.Barr 1978
Cunninghamella blakesleeana Lendner 1927
C. echinulata (Thaxter 1891) Thaxter 1905
C. homothallica Kominami et Tubaki 1952
C. japonica (Saito 1905) Pidoplichko et Milko 1971
C. vesiculosa P.C.Misra 1966
Curvularia clavata B.L.Jain 1962
C. comoriensis Bouriquet et Jauffret 1955 ex M.B.Ellis 1966
C. geniculata (Tracy et Earle 1896) Boedijn 1933
C. inaequalis (Shear 1907) Boedijn 1933
C. lunata (Wakker 1898) Boedijn 1933
C. protuberata Nelson et Hodges 1965
Cyathus olla (Batsch 1783) Persoon 1801
Cylindrium cordae Grove 1886
Cylindrocarpon album (Saccardo 1877) Wollenweber 1917
C. chlamydospora Schischkina et Tzanava 1973
C. congoense J.A.Meyer 1958
C. destructans (Zinssmeister 1918) Scholten 1964 var.
destructans
C. didymum (Hartig 1846) Wollenweber 1926
C. gracile Bugnicourt 1939
C. heteronema (Berkeley et Broome 1865) Wollenweber 1928
C. ianthothele Wollenweber 1917 var. minus Reinking 1938
C. magnusianum Wollenweber 1928
C. obtusisporum (Cooke et Harkness 1884) Wollenweber 1926
C. peronosporae (Fautrey et Lambotte 1896) Rudakov 1981
C. stilbophilum (Corda 1838) Rudakov 1981
C. theobromicola C. Booth 1966
Cylindrocephalum stellatum (Harz 1871) Saccardo 1886
Cylindrophora alba Bonorden 1851
Cryopreservation
Number
Storage
of strains
time (years)
1
2
7
1
4
1
19.34
14.09
19.42
20.07
20.19
14.09
1
2
3
1
17.79
19.28
19.74
19.34
1
0.46
1
1
1
1
17.66
19.96
19.54
15.66
1
1
1
1
1
1
19.54
15.58
19.51
19.51
15.58
19.54
Freeze-Drying
Number
Storage
of strains
time (years)
Soil
Number
of strains
Storage
time (years)
1
18.90
15
35.33
2
12
13.81
29.23
6
33.07
1
28.66
1
1
3
3
4
1
0.65
13.83
19.15
20.46
29.21
1.33
1
4.33
1
1
1
1
1
27.86
29.98
11.89
18.45
14.82
1
2
1
9.05
11.58
18.45
1
1
1
2
3
1
1
14.78
18.94
32.16
29.51
7.89
27.87
19.78
1
1
10.86
S.M. Ozerskaya et al.
344
345
346
347
348
349
350
351
352
353
354
Name of species
30
Table 3.1 (continued)
380
381
382
383
384
1
1
1
1
17.71
6.62
16.08
19.28
1
28.06
1
1
23.58
0.08
2
1
1
20.11
19.31
6.03
1
1
1
1
2
1
1
28.29
14.23
25.87
24.14
15.59
18.88
19.72
1
1
1
1
1
1
1
1
19.26
18.95
2.39
5.74
12.56
19.19
19.39
6.68
1
10.34
4.03
1
1
1
1
1
1
2
2
1
2
12.36
22.93
12.32
28.44
8.30
10.82
28.45
27.21
17.71
17.71
1
1
10.64
17.79
1
1
1
1
18.86
20.30
19.32
1
1
1
17.71
14.78
39.74
1
1
12.31
19.43
1
22.90
1
5.65
1
4.79
1
1.51
31
(continued)
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
359
360
361
362
363
364
365
366
367
368
369
370
371
372
373
374
375
376
377
378
379
C. hoffmannii Daszewska 1912
Dacrymyces stillatus Nees 1817
Dactylaria dimorphospora Veenbaas-Rijks 1973
Dactylellina asthenopaga (Drechsler 1937) M. Scholler,
Hagedorn et A. Rubner 1999
Daedalea quercina (Linnaeus 1753) Persoon 1801
Dendrodochium toxicum Pidoplichko et Bilai 1947
Dendrostilbella macrospora W.Bally 1917
D. mycophila (Persoon 1822) Seifert 1985
Dendryphion nanum (Nees 1816) S.Hughes 1958
D. penicillatum (Corda 1838) Fries 1849
Dichobotrys sp.
Dichotomomyces cejpii (Milko 1964) D.B. Scott 1970
Dictyophora duplicata (Bosc 1811) E.Fischer 1888
Dictyostelium discoideum (Bosc 1811) E.Fischer 1888
Dictyuchus monosporus Leitgeb 1869
Dicyma ampullifera Boulanger 1897
D. olivacea (Emoto et Tubaki 1970) Arx 1982
D. ovalispora (S.Hughes 1951) Arx 1982
Didymopsis helvellae (Corda 1854) Saccardo et Marchall 1885
Dimargaris bacillispora R.K.Benjamin 1959
Dinemasporium strigosum (Persoon 1801) Saccardo 1881
Diplocladium majus Bonorden 1851
D. penicillioides Saccardo 1886
Dipodascopsis tothii (Zsolt 1963) L.R.Batra et Millner 1978
D. uninucleata (Biggs 1937) L.R.Batra et Millner 1978 var.
uninucleata
Dipodascus aggregatus Francke-Grosmann 1952
Discula brunneotingens E.I.Meyer 1953
D. pinicola (Naumov 1926) Petrak 1927 var. mammosa
Lagerberg et al. 1927
Dispira cornuta van Tieghem 1875
Dissoacremoniella silvatica Kirilenko 1970
3
355
356
357
358
32
Table 3.1 (continued)
Sr. No.
385
386
387
388
389
409
1
19.39
Freeze-Drying
Number
Storage
of strains
time (years)
1
20.31
3
1
24.58
13.88
2
22.21
1
19.28
1
12.26
1
19.37
2
1
5.32
18.20
1
19.30
1
19.79
1
1
11
3
4
1
7
2
1
1
9
1
1
1
3
25.93
12.81
39.08
26.90
37.09
26.90
32.76
21.67
21.45
14.81
32.50
22.93
20.39
21.59
11.56
3
1
1
1
10
1
20.43
19.32
19.32
14.87
20.41
20.43
1
19.32
1
13.91
1
0.16
Soil
Number
of strains
Storage
time (years)
1
5.68
1
4.33
9
2
4
1
27.41
11.42
16.74
8.68
1
9.64
S.M. Ozerskaya et al.
390
391
392
393
394
395
396
397
398
399
400
401
402
403
404
405
406
407
408
Name of species
Doratomyces purpureofuscus (Schweinitz 1832) F.J.Morton et
G.Smith 1963
D. stemonitis (Persoon 1801) F.J.Morton et G.Smith 1963
Drechmeria coniospora (Drechsler 1941) W. Gams et H.-B.
Jansson 1985
Drechslera avenacea (M.A.Curtis ex Cooke 1889) Shoemaker
1959
D. biseptata (Saccardo et Roumeguere 1881) M.J.Richardson et
E.M.Fraser 1968
D. campanulata (Leveille 1841) B.Sutton 1976
D. poae (Baudys 1916) Shoemaker 1962
Duddingtonia flagrans (Duddington 1949) R.C.Cooke 1969
Echinobotryum rubrum Sorokin ex Jaczewski 1917
Eladia saccula (E.Dale 1926) G.Smith 1961
Emericella nidulans (Eidam 1883) Vuillemin 1927
E. quadrilineata (Thom et Raper 1939) C.R. Benjamin 1955
E. rugulosa (Thom et Raper 1939) C.R. Benjamin 1955
E. variecolor Berkeley et Broome 1857
Emericellopsis donezkii Beliakova 1974
E. glabra (J.F.H.Beyma 1940) Backus et Orpurt 1961
E. humicola (Cain 1956) Gilman 1956
E. maritima Beliakova 1970
E. minima Stolk 1955
E. pallida Beliakova 1974
E. robusta van Emden et W.Gams 1971
E. terricola J.F.H.Beyma 1940
Engyodontium album (Limber 1940) de Hoog 1978
Entomophthora dipterigena (Thaxter 1888) Saccardo et
Traverso 1891
E. pyriformis Thoizon 1967
Cryopreservation
Number
Storage
of strains
time (years)
1
19.38
433
434
435
436
437
438
2
1
1
1
3
3
1
12.05
12.34
19.80
18.85
17.31
16.31
18.30
1
3.99
1
18.86
1
19.32
1.00
2
1
1
2
1
1
1
9.02
8.17
2.04
12.19
18.48
20.38
19.53
1
1
1
1
2
1
1.64
21.78
5.73
1
20.18
1
24.83
8
5
1
1
7
1
8
4
32.38
20.75
1
5
1
21.57
11.24
19.77
1
38.97
36.93
28.92
37.95
32.45
39.91
15.54
1.64
1
1.64
1
1
2
1
1
1.81
1.64
16.34
21.63
7.93
17.77
19.46
1
1
1
7.93
5.95
18.88
19.63
1
1
1
26.00
10.72
15.02
1
25.98
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
417
418
419
420
421
422
423
424
425
426
427
428
429
430
431
432
E. thaxteriana I.M.Hall et J.Bell 1963
Entyloma gaillardianum Vanky 1982
Epicoccum nigrum Link 1815
Eremascus fertilis Stoppel 1907
Eremothecium ashbyi Guilliermond 1935
E. gossypii (S.F.Ashby et W.Nowell 1926) Kurtzman 1995
Eupenicillium javanicum (J.F.H.Beyma 1929) Stolk et
D.B.Scott 1967 var. javanicum
Eurotium amstelodami (Mangin 1909) Thom et Church 1926
E. chevalieri L. Mangin 1909
E. halophilicum C.M.Christensen et al. 1959
E. herbariorum (F.H.Wiggers 1780) Link 1809
E. rubrum Jos. König et al. 1901
E. tonophilum Ohtsuki 1962
Evlachovaea kintrischica B.Borisov et Tarasov 1999
Exobasidium bisporum Sawada 1950
E. karstenii Saccardo et Trotter 1912
E. myrtilli Siegmund 1879
E. pachysporum Nannfeldt 1981
E. vaccinii (Fuckel 1861) Woronin 1867
E. warmingii Rostrup 1888
Exophiala castellanii Iwatsu et al. 1999
E. heteromorpha (Nannfeldt 1934) de Hoog et Haase 2003
E. lecanii-corni (Benedek et Specht 1933) Haase et
de Hoog 1999
E. moniliae de Hoog 1977
E. salmonis J.W.Carmichael 1966
Exserohilum pedicellatum (A.W. Henry 1924) K.J. Leonard et
Suggs 1974
Farlowiella carmichaeliana (Berkeley 1836) Saccardo 1891
Farrowia seminuda (L.M.Ames 1949) D. Hawksworth 1975
Fennellomyces linderi (Hesseltine et Fennell 1955) Benny et
R.K.Benjamin 1975
3
410
411
412
413
414
415
416
(continued)
33
34
Table 3.1 (continued)
Sr. No.
439
440
441
442
443
444
445
446
447
448
449
450
451
452
Fibroporia vaillantii (de Candolle 1815) Parmasto 1968
Filobasidiella depauperata (Petch 1932) R.A.Samson et al. 1983
Flammulina velutipes (Curtis 1777) Singer 1951
Fomes fomentarius (Linnaeus 1753) Fries 1849
Fomitopsis pinicola (Swartz 1810) P.Karsten 1889
F. rosea (Albertini et Schweinitz 1805) P.Karsten 1881
Fonsecaea pedrosoi (Brumpt 1922) Negroni 1936
Funalia trogii (Berkeley 1850) Bondartsev et Singer 1941
Fusarium agaricorum Sarrazin 1887
F. aquaeductuum (Rabenhorst et Radlkofer 1863)
Lagerheim 1891
F. aquaeductuum (Rabenhorst et Radlkofer 1863) Lagerheim
1891 var. medium Wollenweber 1931
F. arthrosporioides Sherbakoff 1915
F. avenaceum (Fries 1832) Saccardo 1886
F. avenaceum (Fries 1832) Saccardo 1886 var. herbarum
(Corda 1839) Saccardo 1886
F. cerealis (Cooke 1878) Saccardo 1886
F. chlamydosporum Wollenweber et Reinking 1925
F. concolor Reinking 1935
F. culmorum (W.G.Smith 1884) Saccardo 1895
F. decemcellulare Brick 1908
F. episphaeria (Tode 1791) Snyder et Hansen 1945
F. epistroma (Hoehnel 1909) C.Booth 1971
F. equiseti (Corda 1838) Saccardo 1886
F. expansum Schlechtendal 1824
F. fujikuroi Nirenberg 1976
F. graminearum Schwabe 1839
F. heterosporum Nees et T. Nees 1818
F. heterosporum Nees et T. Nees 1818 var. pucciniophilum
1
1
5
1
4
1
1
2
1
1
20.09
7.57
20.01
20.05
20.03
20.09
19.71
20.05
17.66
0.19
2
19.98
1
2.19
2
1
2
1.88
17.66
17.46
2
19.26
1
2
7.64
15.66
1
17.71
Freeze-Drying
Number
Storage
of strains
time (years)
Soil
Number
of strains
Storage
time (years)
1
16.12
1
2
27.21
33.19
1
23.04
1
5.64
1
3
1
10.16
33.20
26.39
2
14.34
1
2
1
3
2
1
2
6
1
1
4
2
1
22.49
29.57
5.10
25.41
33.20
15.12
32.43
29.63
29.47
21.06
38.91
29.86
23.19
2
2
1
11.43
9.66
6.65
2
23.56
1
2
1
21.87
7.09
7.09
S.M. Ozerskaya et al.
453
454
455
456
457
458
459
460
461
462
463
464
465
Name of species
Cryopreservation
Number
Storage
of strains
time (years)
490
491
492
493
494
495
496
3
2
3
1
16.56
19.85
19.78
7.68
2
9
2
17.66
19.86
17.71
3
7.68
1
2
1
1
2
4.98
16.15
17.71
17.71
17.46
15
1
19.98
17.66
1
1
1
3
19.81
10.01
5.61
19.26
1
3
1
2
19.26
12.63
0.47
19.45
1
18.85
4
2
6
2
1
3
20
3
3
9
1
28.49
32.16
32.78
33.20
0.57
27.70
33.20
31.70
14.98
30.03
1.16
1
14
4
2
4
3
21
1
1
21.31
32.19
32.12
31.70
32.19
29.13
38.76
23.96
33.02
3
27.06
3
2
1
6.59
21.85
1
8
1
5.97
23.93
12.68
3
13.96
4
1
1
1
2
16
16.90
15.85
12.84
4.75
12.68
23.65
21.10
1
3.45
1
20.12
1
3.45
14
1
1
1
33.38
20.84
21.00
11.15
1
1
5.31
15.07
35
(continued)
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
477
478
479
480
481
482
483
484
485
486
487
488
489
F. incarnatum (Roberge 1849) Saccardo 1886
F. javanicum Koorders 1907
F. lateritium Nees 1816
F. merismoides Corda 1838
F. merismoides Corda 1838 var. merismoides
F. nivale (Fries 1849) Cesati 1860 ex Saccardo 1886
F. oxysporum Schlechtendal 1824
F. poae (Peck 1903) Wollenweber 1913
F. redolens Wollenweber 1913
F. sambucinum Fuckel 1869
F. sambucinum Fuckel 1869 var. ossicola (Berkeley et
M.A.Curtis 1875) Bilai 1955
F. sarcochroum (Desmazieres 1850) Saccardo 1879
F. solani (Martius 1842) Saccardo 1881
F. sporotrichioides Sherbakoff 1915
F. sporotrichioides Sherbakoff 1915 var. sporotrichioides
F. tricinctum (Corda 1838) Saccardo 1886
F. ventricosum Appel et Wollenweber 1913
F. verticillioides (Saccardo 1882) Nirenberg 1976
F. viride (Lechm.) Wollenweber 1917
F. wolgense Rodigin 1942
Fusicladium pomi (Fries 1825) Lind 1913
Gabarnaudia betae (Delacroix 1897) Samson et W.Gams 1974
Gaeumannomyces caricis J.Walker 1980
Galactomyces geotrichum (E.E.Butler et L.J.Petersen 1972)
Redhead et Malloch 1977
G. reessii (van der Walt 1959) Redhead et Malloch 1977
Ganoderma applanatum (Persoon 1799) Patouillard 1889
Geastrum fimbriatum Fries 1829
Geomyces pannorum (Link 1824) Sigler et J.W.Carmichael 1976
Geosmithia lavendula (Raper et Fennell 1948) Pitt 1980
G. namyslowskii (K.M.Zalessky 1927) Pitt 1980
Geotrichum amycelicum Redaelli et Ciferri 1935
3
466
467
468
469
470
471
472
473
474
475
476
36
Table 3.1 (continued)
Sr. No.
497
498
499
500
501
502
503
504
505
506
519
520
521
522
523
G. bipunctatum Rolland et Fautrey 1894
G. candidum Link 1809
G. fragrans (Berkhout 1923) Morenz 1960 ex Morenz 1964
G. klebahnii (Stautz 1931) Morenz 1964
Gibberella fujikuroi (Sawada 1917) Wollenweber 1931
G. zeae (Schweinitz 1821) Petch 1936
Gibellulopsis nigrescens (Pethybridge 1919)
Zare. W. Gams et Summerbell 2007
Gilbertella persicaria (E.D.Eddy 1925) Hesseltine 1960
Gliocephalotrichum bulbilium J.J.Ellis et Hesseltine 1962
Gliocladiopsis tenuis (Bugnicourt 1939) Crous et
M.J. Wingfeld 1993
Gliocladium album (Preuss 1851) Petch 1926
G. ammoniphilum Pidoplichko et Bilai 1953
G. aurifilum (W. Gerard 1874) Seifert, Samuels et W. Gams 1985
G. cholodnyi Pidoplichko 1931
G. comtus Rudakov 1981
G. deliquescens Sopp 1912
G. penicillioides Corda 1840
G. viride Matruchot 1893
Gliomastix cerealis (P.Karsten 1887) C.H.Dickinson 1968
G. inflata C.H.Dickinson 1968
G. luzulae (Fuckel 1870) E.W. Mason 1953 ex S.Hughes 1958
G. murorum (Corda 1839) S. Hughes 1958 var. felina (Marchal
1895) S.Hughes 1958
G. murorum (Corda 1838) S. Hughes 1958 var. murorum
Gloeophyllum odoratum (von Wulfen 1788) Imazeki 1943
G. sepiarium (von Wulfen 1786) P.Karsten 1879
Gongronella butleri (Lendner 1926) Peyronel et Dal Vesko 1955
G. lacrispora Hesseltine et J.J.Ellis 1961
19
3
2
2
20.39
19.85
19.77
19.31
2
19.31
1
11.41
1
1
2
19.25
0.54
16.14
1
2
7.80
7.80
1
4
6.22
19.90
6
1
5
2
1
19.88
12.39
20.09
19.69
15.37
Freeze-Drying
Number
Storage
of strains
time (years)
1
23
4
3
2
2
5
28.38
31.16
31.67
29.62
5.25
8.72
32.31
1
1
1
27.13
11.92
14.46
2
1
1
2
1
1
2
4
3
2
3
5
26.90
28.55
14.41
26.27
30.99
17.27
28.51
32.23
25.16
23.17
29.44
32.53
10
6
1
Soil
Number
of strains
Storage
time (years)
1
1.59
1
1
2
27.17
5.60
4.80
1
27.09
1
9.57
2
9.13
1
10.97
1
9.25
33.64
1
9.49
32.12
18.86
1
5.42
S.M. Ozerskaya et al.
507
508
509
510
511
512
513
514
515
516
517
518
Name of species
Cryopreservation
Number
Storage
of strains
time (years)
551
552
553
554
1
17.66
1
1
12.58
12.06
1
19.43
1
1
2
1
1
1
12.55
19.37
19.56
4.09
19.28
19.47
1
1
19.29
17.58
1
19.59
4
2
1
1
12.56
12.63
6.83
20.09
1
1
1
27.64
6.36
23.88
1
2
36.56
26.42
1
1
1
2
1
1
3
1
1
1
2
1
1
27.00
14.08
15.65
28.92
28.59
17.19
27.73
3.02
11.44
39.15
27.11
9.56
39.58
1
15.88
15.26
23.26
34.15
26.36
27.42
22.56
27.50
2
19.47
1
1
12
1
1
1
1
19.80
16.92
19.56
19.28
1
1
2
2
1
1
15.39
16.63
1
2.64
2
17.28
37
(continued)
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
531
532
533
534
535
536
537
538
539
540
541
542
543
544
545
546
547
548
549
550
Gonytrichum caesium Nees 1818
G. macrocladum (Saccardo 1880) S.Hughes 1951
Graphium putredinis (Corda 1839) S. Hughes 1958
Grifola frondosa (Dickson 1785) Gray 1821
Guepiniopsis buccina (Persoon 1801) L.L.Kennedy 1958
Gymnoascus reessii Baranetzky 1872
Hansfordia pulvinata (Berkeley et M.A.Curtis 1875)
S.Hughes 1958
H. triumfettae (Hahsford 1943) S. Hughes 1952
Haplaria repens Bonorden 1851
Haplographium delicatum Berkeley et Broome 1859
Haplotrichum capitatum (Link 1809) Link 1824
Hapsidospora milkoi Beliakova 1975
Harposporium lilliputanum Dixon 1952
Harzia acremonioides (Harz 1871) Costantin 1888
Harziella capitata Costantin et Matr. 1899
Helicodendron tubulosum (Riess 1853) Linder 1929
Helicostylum elegans Corda 1842
H. pulchrum (Preuss 1851) Pidoplichko et Milko 1971
Helminthosporium solani Durieu et Montagne 1849
Hemicarpenteles ornatum (Subramanian 1972) Arx 1974
Hericium coralloides (Scopoli 1772) Persoon 1794
H. erinaceus (Bulliard 1791) Persoon 1797
Hesseltinella vesiculosa H.P.Upadhyay 1970
Heterobasidion annosum (Fries 1821) Brefeld 1888
Hirsutella thompsonii F.E. Fischer 1950
Hormiactis alba Preuss 1851
Hormoconis resinae (Lindau 1906) von Arx et G.A.
de Vries 1973
Hormonema macrosporum L.Voronin 1986
H. prunorum (Dennis et Buhagiar 1973) Hermanides-Nijhof 1977
Humicola fuscoatra Traaen 1914
H. grisea Traaen 1914
3
524
525
526
527
528
529
530
38
Table 3.1 (continued)
Sr. No.
555
556
557
558
559
H. grisea Traaen 1914 var. thermoidea Cooney et Emerson 1964
H. insolens Cooney et R. Emerson 1964
Hymenochaete tabacina (Sowerby 1796) Levielle 1846
Hyphozyma sanguinea (C.Ramirez 1952) de Hoog et
M.T.Smith 1981
H. variabilis de Hoog et M.T.Smith 1981 var. odora de Hoog et
M.T.Smith 1981
H. variabilis de Hoog et M.T.Smith 1981 var. variabilis
Hypomyces ochraceus (Persoon 1801) Tulasne et C. Tulasne 1865
Hypsizygus ulmarius (Bulliard 1790) Redhead 1984
Inonotus dryophilus (Berkeley 1847) Murrill 1904
I. obliquus (Ach. ex Persoon 1801) Pilat 1942
I. rheades (Persoon 1825) Bondartsev et Singer 1941
Irpex lacteus (Fries 1818) Fries 1828
Isaria farinosa (Holmskjold 1781) Fries 1832
I. fumosorosea Wize 1904
Itersonilia perplexans Derx 1948
Kickxella alabastrina Coemans 1862
Kuehneromyces lignicola (Peck 1872) Redhead 1984
K. mutabilis (Schaeffer 1774) Singer et A.H.Smith 1946
Laccaria bicolor (Maire 1937) P.D.Orton 1960
Lactarius helvus (Fries 1821) Fries 1838
Laetiporus sulphureus (Bulliard 1788) Murrill 1920
Lasiodiplodia theobromae (Pat. 1892) Griffon et Maublanc 1909
Lecanicillium fungicola (Preus 1851) Zare et W. Gams 2008
L. fusisporum (W.Gams 1971) Zare et W.Gams 2001
L. lecanii (Zimmermann 1898) Zare et W. Gams 2001
L. muscarium (Petch 1931) Zare et W. Gams 2001
L. psalliotae (Treschew 1941) Zare et W. Gams 2001
Leccinum scabrum (Bulliard 1783) Gray 1821
Freeze-Drying
Number
Storage
of strains
time (years)
1
1
11.62
11.62
1
1
19.55
19.28
1
20.45
1
19.28
1
21.64
1
19.28
1
2
20.45
18.58
1
1
1
2
1
3
2
2
1
1
2
1
1
3
1
1
8.54
19.41
8.58
12.29
12.39
19.56
19.51
6.72
6.69
12.07
12.39
0.84
12.58
19.97
7.36
4.90
5
5
1
1
22.64
27.38
9.95
6.46
2
3
1
1
19.83
19.86
19.28
12.07
3
1
2
6
9
27.16
25.03
31.78
29.49
29.57
Soil
Number
of strains
Storage
time (years)
3
1
18.38
2.32
S.M. Ozerskaya et al.
560
561
562
563
564
565
566
567
568
569
570
571
572
573
574
575
576
577
578
579
580
581
582
Name of species
Cryopreservation
Number
Storage
of strains
time (years)
597
598
599
600
601
602
603
604
605
606
607
608
609
610
611
1
20.38
1
21.28
1
1
1
2
2
1
3
1
1
2
20.38
20.51
20.38
8.52
20.09
12.12
20.96
12.32
0.21
8.25
1
2
1
19.35
26.59
25.14
1
1
18.95
3.04
1
1
1.56
6.03
1
1
6.92
7.16
1
17.15
1
1
1
1
1
1
1
8.02
20.03
12.20
12.31
12.06
12.01
0.97
1
11.56
1
24.07
1
22.25
5
1
1
32.69
6.27
5.84
1
2
8.29
19.40
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
584
585
586
587
588
589
590
591
592
593
594
595
596
Lecythophora decumbens (J.F.H.Beyma 1942)
E. Weber et al. 2002
L. fasciculata (J.F.H.Beyma 1939) E. Weber et al. 2002
L. hoffmannii (J.F.H.Beyma 1939) W. Gams et McGinnis 1983
L. mutabilis (J.F.H. Beyma 1944) Gams et McGinnis 1983
Lentinula edodes (Berkeley 1878) Pegler 1975
Lentinus lepideus (Fries 1815) Fries 1825
L. sulcatus Berkeley 1845
L. tigrinus (Bulliard 1781) Fries 1825
Lenzites betulina (Linnaeus 1753) Fries 1838
Lepista luscina (Fries 1818) Singer 1951
L. nuda (Bulliard 1790) Cooke 1871
Leptographium lundbergii Lagerberg et Melin 1927
Leptosphaeria coniothyrium (Fuckel 1870) Saccardo 1875
Leucoagaricus leucothites (Vittadini 1835) M.M. Moser ex
Bon 1977
Linderina pennispora Raper et Fennell 1952
Lobosporangium transversale (Malloch) M.Blackwell et
Benny 2004
Lycoperdon perlatum Persoon 1796
L. pyriforme Schaeffer 1763
Macrolepiota gracilenta (Krombholz 1836) Wasser 1978
M. procera (Scopoli 1772) Singer 1948
M. puellaris (Fries 1863) M.M.Moser 1967
M. rhacodes (Vittadini 1833) Singer 1948
Macrophoma mantegazziana (Penzig 1882) Berlese et
Voglino 1886
Magnusiomyces magnusii (F.Ludwig 1886) de Hoog et M.T.
Smith 2004
Malbranchea cinnamomea (Libert) Oorschot et de Hoog 1984
Marasmius oreades (Bolton 1792) Fries 1836
Mariannaea elegans (Corda 1838) Samson 1974
Melanconium apiocarpum Link 1825
M. bicolor Nees 1817
3
583
2
9.56
39
(continued)
40
Table 3.1 (continued)
Sr. No.
612
613
614
615
616
617
618
619
620
621
622
623
635
636
Melanocarpus albomyces (Cooney et R.Emerson 1964)
von Arx 1975
Melanospora betae Panasenko 1938
M. damnosa (Saccardo 1895) Lindau 1897
M. kurssanoviana (Beliakova 1954) Czerepanova 1962
M. phaseoli Roll-Hansen 1948
Memnoniella echinata (Rivolta 1884) Galloway 1933
Menispora ciliata Corda 1837
Merimbla ingelheimense (J.F.H. Beyma 1942) Pitt 1980
Metarhizium anisopliae (Metschnikoff 1879) Sorokin 1883
Microascus cirrosus Curzi 1930
M. trigonosporus C.W.Emmons et B.O.Dodge var. terreus
Kamyschko 1966
Microbotryum silenes-inflatae (de Candolle 1815 ex Liro 1924)
G.Deml et Oberwinkler 1982
M. violaceum (Persoon 1797) G.Deml et Oberwinkler 1982
Microdiplodia pruni Diedicke 1914
Microsphaeropsis olivacea (Bonorden 1869) Höhnell 1917
Mirandina corticola G.Arnaud 1952 ex Matsushima 1975
Monascus sp.
Monilia brunnea J.C.Gilman et E.V.Abbott 1927
M. diversispora J.F.H.Beyma 1933
M. medoacensis (Saccardo 1913) J.F.H. Beyma 1933
M. megalospora (Berkeley et M.A.Curtis 1869) Saccardo 1886
M. shawi P.Filho
Moniliella suaveolens (Lindner 1895 ex Lindner 1906) von Arx
1972 var. nigra (Burri et Staub 1909) de Hoog 1979
M. suaveolens (Lindner 1895 ex Lindner 1906) von Arx 1972
var. suaveolens
Monilinia fructigena (Aderhold et Ruhland 1905) Honey 1936
Freeze-Drying
Number
Storage
of strains
time (years)
2
18.60
1
34.85
1
1
1
1
2
1
4.06
19.92
16.04
0.12
20.53
19.23
1
1
21.47
9.99
2
1
1
19.54
19.31
20.37
1
2
1
1
2
21.13
12.53
16.73
15.95
22.25
1
36.69
3
12.22
1
1.84
2
1
1
1
12.18
19.89
19.39
19.25
1
1
1
15.21
15.21
19.43
1
4
19.56
19.54
1
1
1
1
1
1
1
1
3
12.05
23.42
5.27
27.24
35.69
28.21
26.72
29.68
23.99
1
18.95
1
10.76
1
20.02
Soil
Number
of strains
Storage
time (years)
1
1
18.03
15.82
1
9.18
S.M. Ozerskaya et al.
624
625
626
627
628
629
630
631
632
633
634
Name of species
Cryopreservation
Number
Storage
of strains
time (years)
1
1
16.31
19.83
1
1
1
1
2
1
1
1
2
1
1
2
2
3
7
3
1
2
2
5
3
3
1
2
4
1
1
1
5
17.77
17.66
3.81
13.86
13.93
11.73
14.34
12.12
11.19
13.86
6.98
14.09
15.97
13.93
15.91
12.12
13.86
13.14
13.99
14.09
13.99
13.28
13.96
19.42
13.99
6.84
3.17
13.99
19.07
1
1
1
2
2
1
2
17.12
15.10
22.59
24.20
21.23
11.72
17.75
2
25.87
21.97
2
21.91
1
1
1
2.74
21.28
16.35
1
1
13.30
1.08
1
2
4.40
22.78
2
12.73
2
1.73
1
2.76
1
1.61
2
3
1
4
1
1
13.93
32.53
39.33
35.06
14.31
14.53
2
1.08
1
4
1
23.94
28.66
1.00
1
5
2.52
22.80
1
1
1.08
0.79
(continued)
41
1
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
Monocillium dimorphosporum W.Gams 1971
M. indicum S.B.Saksena 1955
M. nordinii (Bourchier 1961) W.Gams 1971
M. tenue W.Gams 1971
Monodictys levis (Wiltshire 1938) S.Hughes 1958
M. paradoxa (Corda 1938) S.Hughes 1958
Monographella cucumerina (Lindfors 1919) Arx 1984
Mortierella alliacea Linnemann 1953
M. alpina Peyronel 1913
M. ambigua B.S.Mehrotra 1963
M. angusta Linnemann 1969
M. beljakovae Milko 1973
M. bisporalis (Thaxter 1914) Bjoerling 1936
M. capitata Marchal 1891
M. dichotoma Linnemann 1936 ex W.Gams 1977
M. elasson Sideris et G.E.Paxton 1929
M. elongata Linnemann 1941
M. exigua Linnemann 1941
M. gamsii Milko 1974
M. gemmifera M.Ellis 1940
M. globalpina W.Gams et Veenbaas-Rijks 1976
M. globulifera O.Rostrup 1916
M. horticola Linnemann 1941
M. humilis Linnemann 1936 ex W.Gams 1977
M. hyalina Harz 1871 var. hyalina
M. jenkinii (A.L.Smith 1898) Naumov 1935
M. lignicola (G.W.Martin 1937) W.Gams et R.Moreau 1959
M. longicollis Dixon-Stewart 1932
M. minutissima van Tieghem 1878
M. mutabilis Linnemann 1941
M. nigrescens van Tieghem 1878
M. oligospora Bjoerling 1936
M. parvispora Linnemann 1941
3
637
638
639
640
641
642
643
644
645
646
647
648
649
650
651
652
653
654
655
656
657
658
659
660
661
662
663
664
665
666
667
668
669
42
Table 3.1 (continued)
Sr. No.
670
671
672
673
674
675
676
677
678
679
680
681
682
683
684
685
686
687
688
689
690
691
693
694
695
696
M. polycephala Coemans 1863
M. pulchella Linnemann 1941
M. pusilla Oudemans 1902
M. reticulata van Tieghem et G.Le Monnier 1873
M. sarnyensis Milko 1973
M. sclerotiella Milko 1967
M. simplex van Tieghem et G.Le Monnier 1873
M. strangulata van Tieghem 1875
M. stylospora Dixon-Stewart 1932
M. turficola Y.Ling 1930
M. verticillata Linnemann 1941
M. zonata Linnemann 1936 ex W.Gams 1977
M. zychae Linnemann 1941
Mucobasispora tarikii Moustafa et Abdul-Wahid 1990
Mucor abundans Povah 1917
M. aligarensis B.S.Mehrotra et B.R.Mehrotra 1969
M. amphibiorum Shipper 1978
M. bacilliformis Hesseltine 1954
M. bainieri B.S.Mehrotra et Baijal 1963
M. circinelloides van Tieghem 1875 var. circinelloides
M. circinelloides van Tieghem 1875 var. griseocyanus (Hagem
1908) Schipper 1976
M. circinelloides van Tieghem 1875 var. janssenii
(Lendner 1907) Schipper 1976
M. circinelloides van Tieghem 1875 var. lusitanicus (Bruderlein
1916) Schipper 1976
M. flavus Bainier 1903
M. fragilis Bainier 1884
M. fuscus Bainier 1903
M. genevensis Lendner 1908
1
1
1
1
1
1
1
13.99
16.82
6.98
13.86
11.08
11.95
3.79
1
7
1
5
11.02
19.42
13.95
12.12
1
1
1
1
17.79
20.18
15.37
5.84
4
1
19.54
11.98
9
1
1
1
19.32
19.63
7.14
19.56
Freeze-Drying
Number
Storage
of strains
time (years)
1
1
1
1
21.31
6.83
7.16
39.26
1
1
21.01
15.95
8
1
2
1
1
1
1
1
1
17
3
21.74
5.09
31.68
6.25
29.37
23.28
18.86
28.79
32.41
33.63
22.46
7
Soil
Number
of strains
Storage
time (years)
1
24.34
1
20.46
1
25.98
2
11.98
1
5.37
1
5.99
13
3
28.04
24.30
32.18
6
27.75
8
35.15
3
28.44
19
1
3
3
36.92
16.60
39.12
21.90
6
1
2
26.71
18.53
19.26
S.M. Ozerskaya et al.
692
Name of species
Cryopreservation
Number
Storage
of strains
time (years)
701
702
703
704
705
706
707
708
709
710
711
712
713
714
715
716
717
718
719
720
721
722
723
1
1
19.63
0.14
1
3
34.77
18.48
1
1
8.62
9.59
4
19.67
17
1
40.19
16.83
5
1
28.13
16.22
3
26.19
1
2.61
1
1
2
0.09
26.56
20.52
1
2
24.54
15.41
1
1
3
1
1
4
19.63
11.95
19.58
19.56
19.69
20.05
1
2
5
1
1
9
1
14.72
40.30
31.36
19.44
17.14
38.20
34.63
1
4
1
4
1
1
19.54
18.69
19.67
17.79
20.14
11.78
2
5
1
16
1
4
16.18
27.32
28.03
34.81
14.88
24.74
1
14
5.23
28.41
3
28.30
7
19.54
31
3
38.88
32.34
26
2
30.18
28.30
1
19.63
1
1
17.67
19.87
1
18.75
1
1
2
2
1
15.92
35.92
25.58
22.88
20.64
1
2
1
8.11
23.61
8.38
1
19.63
1
6.83
1
19.63
43
(continued)
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
699
700
M. guilliermondii Nadson et Philippow 1925
M. hiemalis Wehmer 1903 var. corticolus (Hagem 1910)
Schipper 1973
M. hiemalis Wehmer 1903 var. hiemalis
M. hiemalis Wehmer 1903 var. luteus (Linnemann 1936)
Schipper 1973
M. hiemalis Wehmer 1903 var. silvaticus (Hagem 1908)
Schipper 1973
M. inaequisporus Dade 1937
M. indicus Lendner 1930
M. laxorrhizus Y.Ling 1930
M. microsporus Namyslowski 1910
M. mousanensis Baijal et B.S.Mehrotra 1966
M. mucedo Linnaeus 1753
M. oblongiellipticus H.Naganishi, Hirahara et Seshita ex
Pidoplichko et Milko 1971
M. odoratus Treschew 1940
M. piriformis A.Fischer 1892
M. plasmaticus van Tieghem 1875
M. plumbeus Bonorden 1864
M. psychrophilus Milko 1971
M. racemosus Fresenius 1850 var. chibinensis
(Neophytova 1955) Schipper 1976
M. racemosus Fresenius 1850 var. racemosus
M. racemosus Fresenius 1850 var. sphaerosporus
(Hagem 1908) Schipper 1970
M. ramosissimus Samoutsevitch 1927
M. recurvus E.E.Butler 1952 var. indicus Baijal et
B.S.Mehrotra 1965
M. recurvus E.E.Butler 1952 var. recurvus
M. saturninus Hagem 1910
M. sinensis Milko et Beliakova 1971
M. strictus Hagem 1908
M. tuberculisporus Schipper 1978
3
697
698
44
Table 3.1 (continued)
Sr. No.
748
749
750
751
M. ucrainicus Milko 1971
M. variabilis A.K.Sarbhoy 1965
M. zonatus Milko 1967
M. zychae Baijal et B.S.Mehrotra 1965 var. zychae
Mutinus caninus (Hudson 1762) Fries 1849
M. ravenelii (Berkeley et Curtis 1855) E.Fischer 1888
Myceliophthora fergusii (Klopotek 1974) Oorschot 1977
M. lutea Costantin 1892
M. thermophila (Apinis 1962) van Oorschot 1977
Mycena pura (Persoon 1794) P.Kummer 1871
M. viscosa Maire 1910
Mycocladus corymbifer (Cohn 1884) Vanova 1991
Mycogone cervina Ditmar 1817
M. nigra (Morgan 1895) C.N.Jensen 1912
M. rosea Link 1809
Mycosticta cytosporicola Frolov 1968
Mycotypha africana R.O.Novak et Backus 1963
M. indica P.M.Kirk et Benny 1985
Myrothecium cinctum (Corda 1842) Saccardo 1886
M. roridum Tode 1790
M. verrucaria (Albertini et Schweinitz 1805) Ditmar 1813
Myxotrichum setosum (Eidam 1882) G.F.Orr et Plunkett 1963
M. stipitatum (Eidam 1882) G.F.Orr et Kuehn 1963
Nadsoniella nigra Issatschenko 1914 var. hesuelica Lyakh et
Ruban 1970
Nakataea sigmoidea (Cavara 1889) Hara 1939
Nectria cosmariospora Cesati et de Notaris 1863
N. inventa Pethybridge 1919
Nematogonum mycophilum (Saccardo 1886) Rogerson et
W. Gams 1981
1
1
2
1
1
19.58
19.56
6.02
12.30
1.81
2
1
1
6
1
1
1
1
20.49
8.65
12.19
19.54
19.85
19.53
19.37
17.66
2
1
3
19.28
19.47
19.82
1
1
4.06
18.85
1
2
19.29
18.93
1
0.54
Freeze-Drying
Number
Storage
of strains
time (years)
1
1
2
2
17.38
14.04
27.40
39.08
1
1
3
Soil
Number
of strains
Storage
time (years)
1
1
19.25
5.69
1.98
29.47
20.37
1
19.25
18
29.84
17
28.66
4
4
2
1
1
2
4
4
1
1
1
27.93
23.30
22.13
21.41
13.81
16.86
22.02
31.60
12.11
20.40
6.16
2
1
23.42
27.90
2
3.45
S.M. Ozerskaya et al.
724
725
726
727
728
729
730
731
732
733
734
735
736
737
738
739
740
741
742
743
744
745
746
747
Name of species
Cryopreservation
Number
Storage
of strains
time (years)
2
19.32
1
1
18.95
19.39
1
5
1
1
1
1
1
2
1
1
1
1
1
1
15.20
18.94
19.42
18.95
4.39
18.95
17.46
19.45
19.71
9.84
12.57
15.55
0.18
19.57
1
1
1
1
1
2
1
1
1
20.45
15.64
19.75
19.94
16.10
19.54
19.94
15.64
19.85
1
1.88
2
20.63
1
15.69
8
4
1
33.87
25.64
17.57
1
1
4
7.35
27.64
18.56
1
1
1
4
1
1
3
1
1
2
1
1
2
1
12.70
27.80
28.99
27.05
7.85
1.95
8.70
19.95
8.14
33.00
1.05
25.73
20.49
18.62
1
2
14.69
10.07
4
1
9.02
9.02
1
4.37
1
26.03
1
1
7.91
15.98
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
753
754
755
756
757
758
759
760
761
762
763
764
765
766
767
768
769
770
771
772
773
774
775
776
777
778
779
780
781
782
Neocosmospora vasinfecta E.F.Smith 1899 var. africana
(von Arx 1955) Cannon et D.Hawksworth 1984
Neonectria galligena (Bresadola 1901) Rossman et Samuels 1999
Neoscytalidium dimidiatum (Penzig 1887) Crous et Slippers 2006
Neottiospora caricina (Desmazieres 1836) Hoehnel 1924
Neovossia setariae (Ling 1945) Yu et Lou 1962
Neurospora crassa Shear et B.O.Dodge 1927
N. sitophila Shear et B.O.Dodge 1927
N. toroi F.L.Tai 1935
Newbya pascuicola M.C.Vick et M.W.Dick 2002
Niesslia exilis (Albertini et Schweinitz 1805) G.Winter 1887
Nigrospora gossypii Jaczewski 1929
Nigrospora oryzae (Berkeley et Broome 1873) Petch 1924
Nodulisporium verrucosum (J.F.H.Beyma 1929) G. Smith 1954
Nomuraea rileyi (Farlow 1883) Samson 1974
Ochrocladosporium elatum (Harz 1871) Crous et U. Braun 2007
Oedocephalum sp. (Berkeley et Broome 1873) Petch 1924
Oidiodendron cereale (Thuemen 1880) G.L.Barron 1962
O. echinulatum G.L.Barron 1962
O. tenuissimum (Peck 1894) S. Hughes 1958
O. truncatum G.L.Barron 1962
Olpitrichum sp.
Oospora minor Delitsch 1943
O. nicotianae Pezzolato 1899
O. oryzae Ferraris 1902
O. sajanica Ogarkov 1979
O. sulphurea (Preuss 1852) Saccardo et Voglino 1886
O. tenuis (P.Maze 1910) Berkhout 1923
O. uvarum Karamboloff 1931
O. variabilis (Lindner 1898) J.Lindau 1907
Ophiostoma piceae (Münch 1907) Syd. et P. Syd. 1919
Ostracoderma sp.
3
752
(continued)
45
46
Table 3.1 (continued)
Sr. No.
783
784
785
786
787
788
789
790
791
792
Ovadendron sulphureo-ochraceum (J.F.H.Beyma 1933)
Sigler et J.W.Carmichael 1976
Paecilomyces borysthenicus B.A. Borisov et Tarasov 1997
P. inflatus (Burnside 1927) J.W.Carmichael 1962
P. lilacinus (Thom 1910) Samson 1974
P. marquandii (Massee 1898) S.Hughes 1951
P. puntonii (Vuill 1930) Nann. 1934
P. variotii Bainier 1907
Papulaspora biformospora Kirilenko 1971
Papulaspora sp.
Paraconiothyrium sporulosum (W. Gams et Domsch 1969)
Verkley 2004
Parasitella parasitica (Bainier 1884) Sydow 1903
Passalora fulva (Cooke 1883) U. Braun et Crous 2003
Paxillus panuoides (Fries 1818) Fries 1838
Penicillium adametzii K.M.Zalessky 1927
P. albicans Bainier 1907
P. alicantinum C.Ramirez et A.T.Martinez 1980
P. anatolicum Stolk 1968
P. aragonense C.Ramirez et A.T.Martinez 1981
P. arenicola Chalabuda 1950
P. atramentosum Thom 1910
P. aurantioflammiferum C.Ramirez et al. 1980
P. aurantiogriseum Dierckx 1901
P. bilaiae Chalabuda 1950
P. brevicompactum Dierckx 1901
P. brunneum Udagawa 1959
P. camemberti Thom 1906
P. canescens Sopp 1912
P. capsulatum Raper et Fennell 1948
1
Freeze-Drying
Number
Storage
of strains
time (years)
19.77
1
29.32
4
3
1
12
1
1
1
19.45
19.54
16.39
19.40
19.63
19.30
19.39
1
3
12
4
1
19
1
1
2
6.32
2.00
31.69
31.29
11.91
32.02
20.69
23.48
16.28
1
1
2
7.18
18.99
20.01
2
1
22.32
22.51
2
1
1
1
1
1
35.16
22.03
10.67
14.02
10.66
14.54
1
31
1
12
1
10
12
3
10.67
34.27
15.01
28.04
10.59
35.42
36.88
17.24
2
20.50
2
1
20.48
2
2
20.48
19.49
Soil
Number
of strains
Storage
time (years)
5
2
19.37
18.38
13
33.89
1
3.20
3
1
1
1
1
1
1
1
28
1
9
1
9
10
3
33.89
7.15
12.20
15.32
12.20
17.92
2.57
12.20
34.84
18.52
18.10
12.07
24.37
39.72
18.17
S.M. Ozerskaya et al.
793
794
795
796
797
798
799
800
801
802
803
804
805
806
807
808
809
810
Name of species
Cryopreservation
Number
Storage
of strains
time (years)
4
19.34
1
2
1
18.94
18.30
2.51
1
3
2
2.26
18.07
18.30
1
20.48
1
1
20.44
20.50
1
3
29
1
10
16
6
1
1
1
17.69
35.23
35.75
23.35
25.87
37.11
20.26
10.65
28.97
21.73
1
2
22
1
10
12
6
1
1
1
1
9
6
3
1
1
6
5
1
8
9
2
1
1
5
6
3
1
1
1
1
2
5.73
36.80
20.27
17.59
22.03
27.75
19.86
30.96
10.68
30.78
34.77
35.58
15.50
10.68
26.68
19.61
37.41
2.77
10.68
19.85
26.66
29.58
1
7
6
2
1
1
6
5
1
7
9
2
1
1
5
5
3
1
1
1
1
2
12.20
24.93
43.04
14.95
33.89
20.45
18.03
12.20
15.77
9.56
47
18.10
29.21
20.45
21.28
18.04
12.58
20.33
38.34
12.07
25.05
24.90
12.07
25.04
12.20
12.33
24.88
22.37
5.41
12.07
24.92
12.20
14.21
(continued)
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
821
822
823
824
825
826
827
828
829
830
831
832
833
834
835
836
837
838
839
840
841
842
P. castellonense C.Ramirez et A.T.Martinez 1981
P. chermesinum Biourge 1923
P. chrysogenum Thom 1910
P. cinerascens Biourge 1923
P. citreonigrum Dierckx 1901
P. citrinum Thom 1910
P. commune Thom 1910
P. cordubense C.Ramirez et A.T.Martinez 1981
P. corylophilum Dierckx 1901
P. cyaneum (Bainier et Sartory 1913) Biourge 1923 ex
Thom 1930
P. daleae K.M.Zalessky 1927
P. decumbens Thom 1910
P. dierckxii Biourge 1923
P. digitatum (Persoon 1801) Saccardo 1881
P. diversum Raper et Fennell 1948
P. dodgei Pitt 1980
P. duclauxii Delacroix 1892
P. expansum Link 1809
P. fagi A.T.Martinez et C.Ramirez 1978
P. funiculosum Thom 1910
P. glabrum (Wehmer 1893) Westling 1911
P. gladioli Machacek 1928
P. glaucum Link 1805
P. grancanariae C.Ramirez et al. 1978
P. granulatum Bainier 1905
P. griseofulvum Dierckx 1901
P. herqueri Bainier et Sartory 1912
P. hirsutum Dierckx 1901 var. hirsutum
P. hispanicum C.Ramirez et al. 1978
P. humuli J.F.H.Beyma 1937
P. ilerdanum C.Ramirez et al. 1980
P. indonesiae Pitt 1980
3
811
812
813
814
815
816
817
818
819
820
Sr. No.
P. inflatum Stolk et Malla 1971
P. insectivorum (Sopp 1912) Biourge 1923
P. islandicum Sopp 1912
P. italicum Wehmer 1894
P. janczewskii K.M.Zalessky 1927
P. jensenii K.M.Zalessky 1927
P. kirovogradum Beliakova et al.
P. lagena (Delitsch 1943) Stolk et Samson 1983
P. lanosum Westling 1911
P. lapidosum Raper et Fennell 1948
P. lehmanii Pitt 1980
P. lineatum Pitt 1980
P. lividum Westling 1911
P. malacaense C.Ramirez et A.T.Martinez 1980
P. martensii Biourge 1923 var. moldavicum Solovei 1975
P. megasporum Orpurt et Fennell 1955
P. melinii Thom 1930
P. miczynskii K.M.Zalessky 1927
P. minioluteum Dierckx 1901
P. mirabile Beliakova et Milko 1972
P. mongoliae Beliakova et al.
P. multicolor Grigorieva-Manoilova et Poradielova 1915
P. multicolor Novobranova 1972
P. murcianum C.Ramirez et A.T.Martinez 1981
P. novae-zeelandiae J.F.H.Beyma 1940
P. ochrochloron Biourge 1923
P. onobense C.Ramirez et A.T.Martinez 1981
P. ovetense C.Ramirez et A.T.Martinez 1981
P. oxalicum Currie et Thom 1915
P. palmense C.Ramirez et al. 1978
Cryopreservation
Number
Storage
of strains
time (years)
1
1
2.23
14.09
1
2
18.21
19.44
2.26
1
3.99
1
1
1
9.07
4.01
17.98
1
19.47
Freeze-Drying
Number
Storage
of strains
time (years)
1
1
3
3
8
8
1
2
3
3
2
1
2
1
1
2
4
7
6
1
1
1
1
1
5
6
1
1
4
1
2.95
15.05
20.87
20.32
27.39
37.45
5.52
24.97
33.03
23.28
30.40
10.51
15.45
10.65
10.48
15.78
27.39
30.90
10.72
16.95
23.82
19.35
12.94
10.65
22.02
36.27
10.69
10.67
31.32
10.68
Soil
Number
of strains
Storage
time (years)
1
3
3
8
8
1
25.04
12.07
20.35
27.37
26.06
6.07
3
4
2
1
1
1
1
3
4
6
2
1
1
15.57
24.86
24.94
11.55
15.33
12.20
10.71
20.33
20.45
24.92
12.20
9.56
6.07
1
1
5
4
1
1
5
1
15.09
12.20
9.70
17.51
12.20
12.20
24.94
12.20
S.M. Ozerskaya et al.
843
844
845
846
847
848
849
850
851
852
853
854
855
856
857
858
859
860
861
862
863
864
865
866
867
868
869
870
871
872
Name of species
48
Table 3.1 (continued)
1
20.53
1
1
1
18.21
9.25
4.06
2
2
18.30
3.99
4
1
20.48
18.07
2
2
18.30
3.99
2
1
2
17.98
17.98
4.01
1
2.26
7
3
2
2
1
9
1
3
1
9
9
4
12
14
6
1
1
18
7
27.21
20.27
34.66
37.80
5.51
36.78
20.52
36.68
26.72
37.65
21.05
22.90
31.53
36.65
25.87
20.73
11.10
27.37
33.56
8
1
16
1
8
1
1
1
1
1
5
1
9
34.70
7.99
24.81
10.65
21.38
17.63
10.70
32.95
10.66
27.09
34.08
10.63
26.62
5
3
3
2
1
7
1
3
1
7
9
4
12
14
5
8
1
14
1
8
1
1
24.92
6.07
24.93
12.79
24.90
20.12
12.20
1
12.20
3
1
9
15.15
12.20
20.16
(continued)
49
1
17
6
18.52
15.02
24.87
26.35
6.07
24.87
20.45
20.13
17.75
24.92
24.87
15.41
24.99
25.04
18.09
15.32
5.72
24.92
24.90
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
892
893
894
895
896
897
898
899
900
901
902
903
904
P. paxilli Bainier 1907
P. phoeniceum J.F.H.Beyma 1933
P. piceum Raper et Fennell 1948
P. pinophilum Thom 1910
P. poltaviae Beliakova et al.
P. purpurogenum Stoll 1904
P. quercetorum Baghdadi 1968
P. raistrickii G.Smith 1933
P. resticulosum Birkinshaw et al. 1942
P. restrictum J.C.Gilman et E.V.Abbott 1927
P. roqueforti Thom 1906
P. roseopurpureum Dierckx 1901
P. rubrum Stoll 1904
P. rugulosum Thom 1910
P. sclerotiorum J.F.H.Beyma 1937
P. senticosum D.B.Scott 1968
P. severskii Schechovtsov 1981
P. simplicissimum (Oudemans 1903) Thom 1930
P. solitum Westling 1911 var. crustosum (Thom 1930)
Bridge et al. 1989
P. solitum Westling 1911 var. solitum
P. solocongelatus Beliakova et al.
P. spinulosum Thom 1910
P. terraconense C.Ramirez et A.T.Martinez 1980
P. thomii Maire 1917
P. turbatum Westling 1911
P. turolense C.Ramirez et A.T.Martinez 1981
P. umbonatum Sopp 1912
P. valentinum C.Ramirez et A.T.Martinez 1980
P. vanbeymae Pitt 1980
P. variabile Sopp 1912
P. vasconiae C.Ramirez et A.T.Martinez 1980
P. velutinum J.F.H.Beyma 1935
3
873
874
875
876
877
878
879
880
881
882
883
884
885
886
887
888
889
890
891
50
Table 3.1 (continued)
Sr. No.
905
906
907
908
909
910
911
912
913
P. verrucosum Dierckx 1901
P. verruculosum Peyronel 1913
P. vinaceum J.C.Gilman et E.V.Abbott 1927
P. viridicatum Westling 1911
P. vulpinum (Cooke et Massee 1888) Seifert et Samson 1985
P. waksmanii K.M.Zalessky 1927
P. westlingii K.M.Zalessky 1927
P. zacinthae C. Ramírez et A.T. Martínez 1981
Penidiella strumelloidea (Milko et Dunaev 1986) Crous et
U. Braun 2007
Perenniporia medulla-panis (Jacquin 1778) Donk 1967
Periconia macrospinosa Lefebvre et Aar.G. Johnson 1949
Pestalotia macrotricha Klebahn 1914
P. pezizoides de Notaris 1841
Petriella sordida (Zukal 1890) G.L. Barron et J.C. Gilman 1961
Phaeococcomyces nigricans (Rich et Stern 1958) de Hoog 1979
Phaeoisaria hippocrepiformis Milko et Dunaev
Phallus hadriani Ventenat 1798
P. impudicus Linnaeus 1753
Phanerochaete sanguinea (Fries 1828) Pouzar 1973
Phellinus igniarius (Linnaeus 1753) Quelet 1886
P. lundellii Niemelae 1972
P. populicola Niemelae 1975
Phialophora atrovirens (J.F.H.Beyma 1935) Schol-Schwarz 1970
P. bubakii (Laxa 1930) Schol-Schwarz 1970
P. cyclaminis J.F.H. Beyma 1942
P. lagerbergii (Melin et Nannfeldt 1934) Conant 1937
P. melinii (Nannf. 1934) Conant 1937
P. verrucosa Medlar 1915
2
18.07
1
2.26
1
2
1
1
18.54
19.47
19.27
19.85
1
1
1
1
1
3
3
1
1
19.38
15.06
1.76
3.24
20.05
20.01
20.01
20.38
20.40
1
19.37
1
19.74
Freeze-Drying
Number
Storage
of strains
time (years)
Soil
Number
of strains
Storage
time (years)
5
4
4
7
8
5
1
1
1
35.63
31.53
34.62
21.31
37.02
36.82
15.51
26.64
14.15
3
3
4
5
9
5
1
1
19.24
18.52
25.05
24.93
24.80
34.08
9.56
12.20
1
16.13
2
1
1
1
20.58
10.23
23.58
5.26
1
3
1
25.35
15.42
6.36
3
1
5.86
19.52
S.M. Ozerskaya et al.
914
915
916
917
918
919
920
921
922
923
924
925
926
927
928
929
930
931
932
Name of species
Cryopreservation
Number
Storage
of strains
time (years)
952
953
954
955
956
957
958
959
960
961
962
963
1
2
1
1
1
1
1
2.80
20.03
20.09
0.20
8.08
12.19
17.46
1
19.30
1
19.54
1
20.52
1
1
19.45
12.55
1
1
1
2
1
1
2
4
1
3
1
1
18.85
8.01
19.56
19.71
11.44
17.04
15.93
13.13
0.18
4.16
0.19
19.44
2
1
1
6
1
4
24.02
1.73
5.21
25.47
4.61
20.04
1
4
1
1.53
8.45
2.14
1
2
7.21
20.14
1
1
8
2
1
26.61
5.68
31.79
31.87
23.98
1
15.30
1
1.51
51
(continued)
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
945
946
947
948
949
950
951
Phlebia rufa (Persoon 1801) M.P.Christiansen 1960
Phlebiopsis gigantea (Fries 1815) Juelich 1978
Pholiota adiposa (Batsch 1789) P.Kummer 1871
P. lenta (Persoon 1801) Singer 1951
P. nameko (T.Ito 1929) S.Ito et S.Imai apud S.Imai 1933
P. squarrosa (Weigel 1771) P.Kummer 1871
Phoma betae A.B.Frank 1892
P. destructiva Plowright 1881
P. eupyrena Saccardo 1879
P. glomerata (Corda 1840) Wollenweber et Hochapfel 1936
P. hedericola (Durieu et Mont. 1856) Boerema 1976
P. jolyana Pirozynski et Morgan-Jones 1968 var. circinata
(Kuznetzova 1971) Boerema et al. 1977
P. leveillei Boerema et G.J.Bollen 1975
P. lingam (Tode 1791) Desmazieres 1849
P. lycopersici Cooke 1885
P. pinodella (L.K. Jones 1927) Morgan-Jones et K.B. Burch 1987
P. pomorum Thuemen 1879
P. sorghina (Saccardo 1878) Boerema et al. 1973
P. tracheiphila (Petri 1929) L.A. Kantschaweli et Gikaschvili
1948
Phomatospora sp.
Phomopsis helianthi Muntanola-Cvetcovic et al. 1981
Phycomyces blakesleeanus Burgeff 1925
P. nitens (C.Agardh 1823) Kunze 1823
Phyllosticta pucciniospila C. Massalongo 1900
Phytophthora cactorum (Lebert et Cohn 1870) J.Schroeter 1886
P. capsici Leonian 1922
P. cinnamomi Rands 1922
P. cryptogea Pethybridge et Lafferty 1919
P. drechsleri Tucker 1931
P. megasperma Drechsler 1931 var. megasperma
Pidoplitchkoviella terricola Kirilenko 1975
3
933
934
935
936
937
938
939
940
941
942
943
944
Sr. No.
964
965
966
967
968
969
970
971
972
973
974
975
976
977
978
979
980
981
982
983
987
988
Piedraia hortae Fonseca et Leao 1928
P. hortae Fonseca et Leao 1928 var. paraguayensis
Fonseca et Leao 1928
P. sarmentoi M.J.Pereira 1930
Pilaira anomala (Cesati 1851) J.Schroeter 1886
P. caucasica Milko 1970
P. moreaui Y.Ling 1926
Pilobolus crystallinus (F.H.Wiggers 1780) Tode 1784
P. longipes van Tieghem 1878
P. umbonatus Buller 1934
Piptoporus betulinus (Bulliard 1786) P.Karsten 1881
Pirella circinans Bainier 1882
P. circinans Bainier 1882 var. volgogradensis (Milko 1974)
Benny et Schipper 1988
P. naumovii (Milko 1970) Benny et Schipper 1992
Pleurodesmospora coccorum (Petch 1924) Samson,
W. Gams et H.C. Evans 1980
Pleurophoma cava (Schulzer 1871) Boerema 1996
Pleurotus cornucopiae (Paulet 1793) Rolland 1910
P. eryngii (De Candolle 1805) Quelet 1872
P. ostreatus (Jacquin 1775) P.Kummer 1871
P. pulmonarius (Fries 1821) Quelet 1872
Pochonia bulbillosa (W. Gams et Malla 1971) Zare et
W. Gams 2001
P. chlamydosporia (Goddard 1913) Zare et W. Gams 2001
Polycephalomyces tomentosus (Schrad. 1799) Seifert 1985
Polyscytalum pustulans (M.N.Owen et Wakefield 1919)
M.B. Ellis 1976
Preussia fleischhakii (Auerswald 1866) Cain 1961
Protomyces macrosporus Unger 1834
Cryopreservation
Number
Storage
of strains
time (years)
Freeze-Drying
Number
Storage
of strains
time (years)
Soil
Number
of strains
Storage
time (years)
1
1
19.34
19.34
1
1
15.70
7.90
1
19.34
1
0.02
1
3.68
1
1
1
1
1
1
1
17.54
26.00
16.20
16.74
15.64
15.64
15.65
3
1
12.39
19.32
1
1
27.13
14.71
1
14.40
1
19.30
1
1
15.35
28.25
1
25.59
1
1
1
16
2
1
19.80
20.01
20.14
20.14
8.15
19.98
3
23.72
3
28.36
1
1
1
19.45
19.53
15.56
1
1
1
31.72
24.82
30.58
1
18.99
1
19.14
S.M. Ozerskaya et al.
984
985
986
Name of species
52
Table 3.1 (continued)
19.42
20.43
18.86
20.43
19.32
20.37
19.32
19.32
18.50
18.50
12.22
2
3
1
1
1
1
1
1
1
1
12.34
15.82
18.95
5.90
6.50
17.04
15.98
8.99
0.15
0.03
1
19.69
1
17.68
1
1
5
19.45
19.30
19.90
1
3
19.67
19.69
2
1
1
2
1
9
1
2
1
1
21.30
21.45
14.48
20.65
22.82
36.92
16.38
20.63
5.32
1.84
1
6.69
1
1
13.76
17.28
2
1
1
1
1
1
17.40
20.91
5.98
12.15
28.38
21.52
1
1
1
4
5.53
14.94
21.41
28.96
1
3.42
1
3.81
2
1
24.86
24.89
1
3
5.60
10.96
1
4
17.16
22.48
(continued)
53
2
1
1
2
1
9
1
2
1
1
1
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
Pseudallescheria boydii (Shear 1922) McGinnis et al. 1982
Pseudeurotium bakeri C.Booth 1961
P. desertorum Mouchacca 1971
P. ovale Stolk 1955 var. milkoi Belyakova 1969
P. ovale Stolk 1955 var. ovale
P. zonatum J.F.H.Beyma 1937
Pseudogymnoascus caucasicus Cejp et Milko 1966
P. roseus Raillo 1929
Puccinia albescens (Greville 1824) Plowright 1888
P. bupleuri F.Rudolphi 1829
P. suaveolens (Persoon 1801) Rostrup 1869
Pycnidiella resinae (Ehrenberg 1818) Hoehnel 1915
Pycnoporus cinnabarinus (Jacquin 1776) Fries 1881
Pyricularia grisea Saccardo 1880
Pyronema omphalodes (Bulliard 1791) Fuckel 1870
Pythium heterothallicum W.A.Campbell et F.F.Hendrix 1968
P. intermedium de Bary 1881
P. mamillatum Meurs 1928
P. oedichilum Drechsler 1930
P. paroecandrum Drechsler 1930
P. spinosum Sawada 1926
P. sylvaticum W.A.Campbell et F.F.Hendrix 1967
Radiomyces embreei R.K.Benjamin 1960
R. spectabilis Embree 1959
Ramichloridium biverticillatum Arzanlou et Crous 2007
Rhinocladiella atrovirens Nannfeldt 1934
Rhinotrichum aureum Cooke et Massee 1889
R. lanosum Cooke 1871
Rhizoctonia crocorum (Persoon 1801) De Candolle 1815
R. solani J.G.Kuehn 1858
R. tuliparum (Klebahn 1905) Whetzel et J.M. Arthur 1924
Rhizomucor miehei (Cooney et R.Emerson 1964) Schipper 1978
R. pusillus (Lindt 1886) Schipper 1978
3
989
990
991
992
993
994
995
996
997
998
999
1000
1001
1002
1003
1004
1005
1006
1007
1008
1009
1010
1011
1012
1013
1014
1015
1016
1017
1018
1019
1020
1021
54
Table 3.1 (continued)
Name of species
1022
1023
R. tauricus (Milko et Schkurenko 1970) Schipper 1978
Rhizopus microsporus van Tieghem 1875 var. chinensis
(Saito 1904) Schipper et Stalpers 1984
R. microsporus van Tieghem 1875 var. microsporus
R. microsporus van Tieghem 1875 var. oligosporus van Tieghem
1875
R. microsporus van Tieghem 1875 var. rhizopodiformis
(Cohn 1884) Schipper et Stalpers 1984
R. oryzae Went et Prinsen Geerligs 1895
R. stolonifer (Ehrenberg 1818) Vuillemin 1902 var. stolonifer
Robillarda sessilis (Saccardo 1878) Saccardo 1880
Rosellinia mammiformis (Persoon 1801) Cesati et
de Notaris 1863
Rozites caperata (Persoon 1796) P.Karsten 1879
Russula decolorans (Fries 1821) Fries 1838
R. grisea Fries 1838
R. velutipes Velenovsky 1920
R. vesca Fries 1838
Rutola graminis (Desmazieres 1834) J.L. Crane et
Schoknecht 1977
Saksenaea vasiformis S.B.Saksena 1953
Saprochaete gigas (Smit et L.Meyer 1928) de Hoog et
M.T.Smith 2004
Saprolegnia asterophora de Bary 1860
S. blelhamensis (M.W.Dick 1969) Milko 1979
S. ferax (Gruithuisen 1821) Nees 1843
S. litoralis Coker 1923
S. mixta de Bary 1883
S. terrestris Cookson 1937 ex R.L.Seymour 1970
S. unispora (Coker et Couch 1923) R.L.Seymour 1970
1024
1025
1026
1027
1028
1029
1030
1031
1032
1033
1034
1035
1036
1037
1038
1039
1040
1041
1042
1043
1044
1045
Freeze-Drying
Number
Storage
of strains
time (years)
Soil
Number
of strains
Storage
time (years)
1
3
19.63
11.63
2
5
20.82
34.24
2
5
24.69
27.09
3
19.71
10
2
36.72
22.07
10
2
28.45
28.09
4
8.70
38.13
33.61
6.61
10.72
24
14
30.35
28.66
2
3
19.71
19.71
1
19.27
26
23
1
1
1
1
1
1
1
1
12.09
12.02
20.01
12.30
8.81
19.39
1
6.87
1
1
19.05
19.86
1
16.65
1
2
1
1
1
1
1
15.19
13.34
13.80
13.09
0.17
0.17
0.17
S.M. Ozerskaya et al.
Sr. No.
Cryopreservation
Number
Storage
of strains
time (years)
1063
1064
1065
1066
1067
1068
1069
1070
1071
1072
1073
1074
1075
3
1
2
1
1
9
1
20.09
18.95
19.31
19.31
19.25
20.47
15.24
1
1
1
19.28
15.89
19.54
1
1
2
1
7.75
19.96
18.87
19.56
1
1
1
1
1
1
1
1
1
1
1
1
1
1
5.10
1
1
14
1
1
1
23.60
26.22
32.48
21.54
31.33
23.56
1
1
6.85
27.21
3
24.26
19.36
2
20.55
19.41
19.41
19.41
19.41
19.41
19.41
19.76
7.49
12.66
19.98
19.89
15.89
1
1
1
1
1
1
1
22.15
11.45
19.24
13.52
13.52
22.24
13.52
1
5.76
1
1
15.95
2.41
1
1
7
4.86
9.56
18.53
1
20.68
1
9.50
1
7.15
1
1
10.80
5.01
(continued)
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
1062
Schizophyllum commune Fries 1815
Sclerotinia ricini G.H.Godfrey 1919
S. sclerotiorum (Libert 1837) de Bary 1884
Scopulariopsis acremonium (Saccardo 1882) Bainier 1907
S. asperula (Saccaardo 1882) Hughes 1958
S. brevicaulis (Saccardo 1882) Bainier 1907
S. brumptii Salvanet-Duval 1935
S. coprophila (Cooke et Massee 1887) W. Gams 1971
S. flava (Sopp 1912) F.J.Morton et G.Smith 1963
S. halophilica Tubaki 1973
S. koningii (Oudemans 1902) Vuuillemin 1911
Sepedonium ampullosporum Damon 1952
S. macrosporum Saccardo et Cavara 1900
Septoria lycopersici Spegazzini 1881
Serpula lacrymans (von Wulfen 1781) J.Schroeter 1888
Simplicillium lamellicola (F.E.W. Smith 1924) Zare et
W. Gams 2001
Sordaria fimicola (Roberge ex Desmazières 1849)
Cesati et de Notaris 1863
Spadicesporium acrosporum V.N.Borisova et Dvoinos 1982
S.acrosporum-majus V.N.Borisova et Dvoinos 1982
S.bifurcatum V.N.Borisova et Dvoinos 1982
S.bifurcatum-majus V.N.Borisova et Dvoinos 1982
S.copiosum V.N.Borisova et Dvoinos 1982
S.persistens V.N.Borisova et Dvoinos 1982
S.ramosum V.N.Borisova et Dvoinos 1982
Sparassis crispa (von Wulfen 1781) Fries 1821
Sphaerellopsis filum (Bivona-Bernardi 1813–1816) Sutton 1977
Sphaeropsis malorum Peck 1883
S. sapinea (Fries 1823) Dyko et B. Sutton 1980
Sporodiniopsis dichotoma van Hoehnel 1903
Sporormiella australis (Spegazzini 1887) S.I.Ahmed et Cain 1972
3
1046
1047
1048
1049
1050
1051
1052
1053
1054
1055
1056
1057
1058
1059
1060
1061
55
56
Table 3.1 (continued)
Name of species
1076
S. intermedia (Auerswald 1868) S.I.Ahmed et Cain ex
Kobayasi 1969
Sporothrix fungorum de Hoog et G.A. de Vries 1973
Sporotrichum aeruginosum Schweinitz 1886 var. microsporum
Karsten 1905
S. bombycinum (Corda 1839) Rabenhorst 1844
S. gorlenkoanum Kuritzina et Sizova 1967
S. laxum Nees 1816
S. mycophilum Link 1818
S. pruinosum J.C.Gilman et E.V.Abbott 1927
S. roseolum Oudemans et Beijerinck 1903
Stachybotrys chartarum (Ehrenberg 1818) S.Hughes 1958
S. cylindrospora C.N.Jensen 1912
Stachylidium variabile Schaeffer et Saccardo
Stagonospora paludosa (Saccardo et Spegazzini 1879)
Saccardo 1884
Stemphyliomma sp.
Stemphylium botryosum Wallroth 1833
S. sarciniforme (Cavara 1890) Wiltshire 1938
Stenocarpella maydis (Berkeley 1847) B. Sutton 1980
Stephanoma sp.
Stereum hirsutum (Willdenow 1787) Persoon 1800
S. sanguinolentum (Albertini et Schweinitz 1805) Fries 1838
Stigmina carpophila (Leveille 1843) M.B. Ellis 1959
Stilbella bulbicola Hennings 1905
Stilbotulasnella conidiophora Bandoni et Oberwinkler 1982
Strobilomyces strobilaceus (Scopoli 1770) Berkeley 1851
Stropharia rugosoannulata Farlow ex Murrill 1922
Syncephalastrum racemosum Cohn ex J.Schroeter 1886
Syncephalis cornu van Tieghem et G.Le Monnier 1873
1077
1078
1079
1080
1081
1082
1083
1084
1085
1086
1087
1088
1089
1090
1091
1092
1093
1094
1095
1096
1097
1098
1099
1100
1101
1102
1
19.29
1
19.29
6
1
1
19.81
15.19
19.30
1
19.89
1
1
17.05
17.79
1
2
1
1
1
1
1
1
2
16.23
20.03
20.03
19.91
19.44
12.54
15.10
12.20
19.24
Freeze-Drying
Number
Storage
of strains
time (years)
Soil
Number
of strains
Storage
time (years)
1
11.76
1
1
1.94
32.35
3
1
1
1
8
1
11
1
1
29.59
23.64
26.27
27.68
31.16
26.16
26.67
14.86
19.09
1
1
27.59
20.27
1
5.84
2
9.56
1
1
3
1
1
26.05
23.43
23.43
11.57
16.03
1
25.87
1
29.49
12
1
29.66
20.64
12
1
29.24
24.09
S.M. Ozerskaya et al.
Sr. No.
Cryopreservation
Number
Storage
of strains
time (years)
1129
1130
1131
1132
1133
1
19.31
1
18.07
1
18.07
1
1
1
1
1
1
3
18.99
8.12
7.17
18.99
19.28
17.71
19.69
1
19.83
1
1
3
3
14.44
4.06
18.93
19.32
1
3
1
20.50
19.23
18.86
1
1
2
19.49
17.68
19.19
1
12.54
1
1
1
4
8
1
1
3
2
23.87
9.74
21.01
15.44
37.03
27.27
17.19
22.55
22.58
1
19.47
1
4
7
1
1
3
2
11.55
24.86
18.17
12.07
1.25
21.86
12.07
3
4
27.08
29.92
3
4
12.14
28.16
1
2
1
1
3
3
1
25.82
13.79
12.14
17.91
20.63
18.92
17.81
3
2
33.99
19.98
1
3.14
2
1
5
1
1
18.54
7.31
26.17
29.01
1.84
57
(continued)
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
1119
1120
1121
1122
1123
1124
1125
1126
1127
1128
S. nodosa van Tieghem 1875
Taeniolella aquatilis (Woronichin 1925) Milko 1985
Talaromyces emersonii Stolk 1965
T. flavus (Kloecker 1902) Stolk et Samson 1972
T. luteus (Zukal 1889) C.R. Benjamin 1955
T. stipitatus (Thom 1935) C.R.Benjamin 1955
T. thermophilus Stolk 1965
T. ucrainicus Udagawa 1966
T. wortmannii (Kloecker 1903) C.R.Benjamin 1955
Taphrina bergeniae Döbbeler 1979
T. carnea Johanson 1886
T. deformans (Berkeley 1857) Tulasne 1866
T. pruni (Fuckel 1861) Tulasne 1866
Tetraploa aristata Berkeley et Broome 1850
Thamnidium elegans Link 1809
Thamnostylum piriforme (Bainier 1880) Arx et H.P.
Upadhyay 1970
Thelebolus polysporus (P.Karsten 1871) Otani et Kanzawa 1970
Thermomyces ibadanensis Apinis et Eggins 1966
Thielavia appendiculata Srivastava et al. 1966
T. hyrcaniae Nicot 1961
T. inaequalis Pidoplichko et al. 1973
T. ovispora Pidoplichko et al. 1973
T. pallidospora Pidoplichko et al. 1973
T. terrestris (Apinis 1963) Malloch et Cain 1972
T. terricola (J.C.Gilman et E.V.Abbott 1927) Emmons 1930
T. terricola (J.C.Gilman et E.V.Abbott 1927) Emmons 1930 var.
minor (Rayss et Borut 1958) C.Booth 1961
Thielaviopsis basicola (Berkeley et Broome 1850) Ferraris 1912
Thysanophora canadensis Stolk et Hennebert 1968
T. penicillioides (Roumeguere 1890) W.B.Kendrick 1961
Tilachlidium pinnatum Preuss 1851
Tilletia caries (de Candolle 1815) Tulasne et C.Tulasne 1847
3
1103
1104
1105
1106
1107
1108
1109
1110
1111
1112
1113
1114
1115
1116
1117
1118
58
Table 3.1 (continued)
Name of species
1134
1135
1136
1137
1138
1139
1140
1141
1142
1143
1144
Tilletiopsis albescens Gokhale 1972
T. washingtonensis Nyland 1950
Tolypocladium cylindrosporum W.Gams 1971
T. geodes W.Gams 1971
T. inflatum W.Gams 1971
Trametes hirsuta (von Wulfen 1788) Pilat 1939
T. pubescens (Schumacher 1803) Pilat 1939
T. versicolor (Linnaeus 1753) Lloyd 1921
T. zonatella Ryvarden 1978
Tricellula aquatica J.Webster 1959
Trichaptum abietinum (Persoon ex J.F. Gmelin 1792)
Ryvarden 1972
Trichocladium asperum Harz 1871
T. opacum (Corda 1837) S.Hughes 1952
Trichoderma album Preuss 1851
T. aureoviride Rifai 1969
T. citrinoviride Bissett 1984
T. flavofuscum (J.H.Miller et al. 1957) Bissett 1991
T. hamatum (Bonorden 1851) Bainier 1906
T. harzianum Rifai 1969
T. koningii Oudemans 1902
T. longibrachiatum Rifai 1969
T. parceramosum Bissett 1991
T. polysporum (Link 1816) Rifai 1969
T. pseudokoningii Rifai 1969
T. reesei Simmons 1968
T. saturnisporum Hammill 1970
T. virens (J.H. Miller, Giddens et A.A. Foster 1957) Arx 1987
T. virgatum Rifai
T. viride Persoon 1801
1145
1146
1147
1148
1149
1150
1151
1152
1153
1154
1155
1156
1157
1158
1159
1160
1161
1162
1
3
1
16.39
18.85
19.83
2
3
2
1
2
1
2
19.26
12.58
20.05
11.24
8.25
16.21
20.09
1
1
1
3
19.54
19.41
15.66
19.81
2
7
6
3
19.98
19.54
19.81
19.40
3
2
4
1
2
19.29
19.79
19.81
0.54
18.89
15
20.45
Freeze-Drying
Number
Storage
of strains
time (years)
1
3
1
3
4
9.92
29.65
23.58
6.25
22.72
2
22.18
1
3
3
3
1
1
3
12
6
8
1
6
3
5
2
2
1
22
15.64
13.52
32.91
28.44
1.92
14.25
31.34
32.01
31.37
28.10
14.21
29.00
27.88
28.41
14.25
32.33
14.21
37.88
Soil
Number
of strains
Storage
time (years)
2
11.75
1
4
5
2
10.01
10.27
14.29
11.90
1
4
6.33
5.05
2
18.38
12
20.68
S.M. Ozerskaya et al.
Sr. No.
Cryopreservation
Number
Storage
of strains
time (years)
1
19.58
1
1
1
5
1
3
1
19.75
0.54
19.28
20.41
17.68
19.86
19.45
1
1
1
3
1
1
5
3
3
1
1
1
1
1
1
1
1
20.52
19.59
19.54
19.44
19.47
19.30
19.42
14.34
19.42
6.83
18.50
12.13
9.32
12.31
0.20
18.50
12.25
2
19.31
1
1
28.08
19.22
1
1
1
9
1
5
1
1
1
2
6
9
6
3
2
7
2
11
3
1
1
1
1
27.05
27.82
26.16
39.07
6.37
32.87
20.34
30.21
15.83
8.62
24.79
29.83
24.42
24.07
24.59
35.00
16.85
23.76
29.13
1.84
1.64
1.64
1.81
1
1
1
2
1
1
1.64
1.84
19.39
15.60
33.19
9.19
4
8.98
1
3.38
1
8.47
1
1
8.34
25.87
7
2
11
2
28.33
17.20
28.66
21.06
1
4.79
59
(continued)
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
1165
1166
1167
1168
1169
1170
1171
1172
1173
1174
1175
1176
1177
1178
1179
1180
1181
1182
1183
1184
1185
1186
1187
1188
1189
1190
1191
1192
1193
1194
T. viride Persoon 1801 var. kizhanicum Krapivina 1975
Trichosporiella cerebriformis (G.A. de Vries et Kleine-Natrop
1957) W. Gams 1971
Trichosporon dulcitum (Berkhout 1923) Weijman 1979
Trichosporum herbarum Jaap 1916
Trichothecium plasmoparae Viala 1932
T. roseum (Persoon 1801) Link 1809
Trichurus spiralis Hasselbring 1900
Tritirachium oryzae (Vincens 1923) de Hoog 1972
Truncatella angustata (Persoon 1801) S.Hughes 1958
Tympanosporium parasiticum W.Gams 1974
Ugola praticola (Pidoplichko 1950) Stalpers 1984
Ulocladium alternariae (Cooke 1871) E.G. Simmons 1967
U. atrum Preuss 1852
U. botrytis Preuss 1851
U. chartarum (Preuss 1851) E.G.Simmons 1967
U. consortiale (Thuemen 1876) E.G.Simmons 1967
U. oudemansii E.G.Simmons 1967
Umbelopsis isabellina (Oudemans 1902) W.Gams 2003
U. nana (Linnemann 1941) Arx 1984
U. ramanniana (Moeller 1903) W.Gams 2003
U. vinacea (Dixon-Stewart 1932) Arx 1984
Ustilago cordae Liro 1924
U. cynodontis (Hennings 1892) Hennings 1893
U. filiformis (Schrank 1793) Rostrup 1890
U. hordei (Persoon 1801) Lagerheim 1889
U. maydis (de Candolle 1815) Corda 1842
U. perennans Rostrup 1890
U. vinosa (Berkeley 1847) Tulasne et C.Tulasne 1847
Venturia sp.
Verticillium albo-atrum Reinke et Berthold 1879
V. aspergillus Berkeley et Broome 1873
V. cellulosae Dasz. 1912
3
1163
1164
Name of species
1195
1196
1197
1198
1199
1200
1201
1202
1203
1204
1205
1206
1207
V. cercosporae Petrak et Ciferri 1932
V. dahliae Klebahn 1913
V. epiphytum Hansford 1943
V. fumosum Seman 1968
V. insectorum (Petch 1931) W.Gams 1971
V. lecanii (Zimmermann 1898) Viegas 1939
V. leptobactrum W.Gams 1971
V. nigrescens Pethybridge 1919
V. nubilum Pethybridge 1919
V. sulphurellum Saccardo 1882
V. tricorpus I.Isaac 1953
V. villosum Rudakov 1981
Viennotidia humicola (Samson et W.Gams 1974) P.F.Cannon et
D.Hawksworth 1982
Volutella ciliata (Albertini et Schweinitz 1805) Fries 1832
Wallemia sebi (Fries 1832) Arx 1970
Wallrothiella subiculosa Hoehnel 1912
Wardomyces anomalus Brooks et Hansford 1923
Westerdykella dispersa (Clum 1955) Cejp et Milko 1964
W. multispora (Saito et Minoura ex Cain 1961) Cejp et
Milko 1964
Xeromyces bisporus L.R.Fraser 1953
Xylobolus frustulatus (Persoon 1801) Boidin 1958
Zygorhynchus exponens Burgeff 1924
Z. heterogamus (Vuillemin 1886) Vuillemin 1903
Z.macrocarpus Y.Ling 1930
Z. moelleri Vuillemin 1903
Zygosporium echinosporum Bunting et E.W.Mason 1941
Z. mycophilum (Vuillemin 1910) Saccardo 1911
1208
1209
1210
1211
1212
1213
1214
1215
1216
1217
1218
1219
1220
1221
Cryopreservation
Number
Storage
of strains
time (years)
12
19.75
1
19.25
3
2
19.36
8.84
1
1
1
1
19.31
19.26
19.31
1
2
19.83
19.59
1
1
19.34
3.99
1
1
3
18.85
19.99
20.10
3
1
1
19.69
19.54
19.54
Freeze-Drying
Number
Storage
of strains
time (years)
1
15
1
1
1
7
1
1
1
1
1
1
1
18.56
31.61
21.55
23.56
14.98
26.42
15.14
4.67
23.64
14.76
28.32
27.38
5.05
1
2
1
1
1
1
21.53
13.22
21.53
1.11
19.68
19.94
1
15.48
4
1
1
5
1
1
19.46
27.33
9.93
25.09
20.24
10.79
Soil
Number
of strains
Storage
time (years)
12
7.96
1
4.74
1
4.82
4.97
1
5.01
1
1.66
2
1
14.60
1.00
S.M. Ozerskaya et al.
Sr. No.
60
Table 3.1 (continued)
3
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
Freeze-Drying of Filamentous Fungi
Currently, freeze-drying is used to preserve
approximately 80% of filamentous fungi maintained
in VKM (2,991 strains, belonging to 1,010 species and 303 genera). Fungi from different taxonomical groups (Zygomycetes, Ascomycetes—
both teleo- and anamorph) able to produce
dormant structures (spores, sclerotia, etc.) usually
survive freeze-drying [12]. According to our data,
from 87 to 92% strains of these fungal groups
remain alive in this method. We noticed that 57%
of freeze-dried cultures stored at 5°C for more
than 20 years were in a vital state, and cultures of
more than 190 species have been sustained for
even 30–40 years of storage. Some species did not
survive freeze-drying even when the sporulation
is abundant, those are: Conidiobolus coronatus,
C. obscurus (syn. Entomophthora thaxteriana),
C. thromboides (syn. Entomophthora virulenta),
Erynia conica (syn. Entomophthora conica),
Pandora dipterigena (syn. Entomophthora
dipterigena), Cunninghamella homothallica,
Cunninghamella vesiculosa. Species of genus
Botrytis (B. cinerea, B. fabae, and B. gladiolorum), forming only sclerotia as a dormant structure, remain in vital state in freeze-drying only for
rather a short time—less than 10 years [10].
Nonsporulating
microorganisms
from
Oomycetes and Basidiomycetes are not stored in
VKM by freeze-drying, since sterile mycelium
generally do not remain viable. However, some
ectomycorrhizal fungi (e.g., Laccaria laccata)
could be successfully lyophilized. For the positive result preliminary slow freezing (to −32°C)
of fungal material is required [13].
The equipment used in VKM for freezedrying is centrifugal freeze-dryer system Micromodulyo (Edwards, UK).
Drying in Sterile Soil of Filamentous
Fungi
This simple and popular method for preservation of
fungi was applied in the beginning of the twentieth
century [14]. Species of Aspergillus, Penicillium
61
can be maintained by this way more effectively than
other micromycetes. According to T.P. Suprun [15]
who investigated preservation of 78 Penicillium
species (more than 1,000 strains) in sterile soil for
7–10 years, the best preserved strains were representatives of Assymmetrica section. Less effectively
preserved species were Biverticillata-Symmetrica
and the lowest effectiveness was observed with
strains of the section Monoverticillata.
Species of Zygomycetes could be stored in
soil for periods ranging from 6 months
(Cunninghamella elegans) to 5 years (Rhizopus
stolonifer var. stolonifer [syn. R. nigricans]) [16].
This method is also efficient for preservation
of some human, animal, and plant pathogens with
retaining their virulence [7]. For example,
Alternaria japonica (syn. A. raphani), Fusarium
oxysporum, and the species of Septoria (S. avenae, S. nodorum, S. passerinii, S. tritici) have
retained their ability to infect a plant host after
2–5 years of storage [17–19]. Some degraded
strains of micromycetes partly recuperated their
lost qualities after preservation in soil [15].
Species of Alternaria, Pseudocercosporella,
Septoria are genetically more stabile compared
with Fusarium, therefore they did not show these
kinds of changes and can be effectively preserved
in soil [3].
Protocols
Protocol of Cryopreservation
Preparation of Cryovials (2.0 mL
Externally Threaded Polypropylene,
Nunc, Denmark)
• Labelled (6 for each culture) with an index, a
collection number of a strain and a date of
cryopreservation (month, year).
• Sterilized by autoclaving, at 121°C for 20 min.
Preparation of Cryoprotectant: 10% (v/v)
Glycerol
• Pour 5 mL of glycerol into 12 mL glass tubes.
• Sterilized by autoclaving at 121°C for
20 min.
• Stored at +5°C for no longer than a month.
62
S.M. Ozerskaya et al.
Preparation of Cultures
• Grow sporulating fungal cultures on slant agar
under optimal growth condition and on suitable medium (www.vkm.ru).
• Wash off spores from agar surface with 5 mL
of cryoprotectant.
• Titer of spores’ suspension should be not less
than 106 spores/mL.
• Grow nonsporulating fungal cultures on Petri
dishes under optimal growth condition and on
suitable medium with agar concentration 5%
(w/v). Incubate culture to get a mature colony.
• Cut mycelial plugs (5 mm diameter) from vigorously growing colony part.
The Second Protocol
• Place 18 cryovials with cultures (mycelial
plugs) in a special metal container, thermostatic inside by expanded polystyrene (container made in VKM).
• Place the container in the ultra-low temperature freezer (−70°C). It was empirically shown
in VKM that temperature in this container
decreased by 0.4°C/min.
• When temperature achieves −70°C, transfer
cryovials in a special container (thoroughly
fixate a position of every vial), and place in
the vapour phase of liquid nitrogen or in the
ultra-low temperature freezer.
Filling of Vials
• For sporulating cultures add 0.2-mL aliquots
of suspension to each cryovial using a Pasteur
pipet. This procedure is carried out under sterile aerobic conditions.
• For nonsporulating cultures place 4 mycelial
plugs into each cryovial using transfer needle,
and then add 0.2 mL of cryoprotectant.
Thawing
• Pull out the cryovial of container, in which it
was stored—either in cryogenic tank or in an
ultra-low temperature freezer.
• Warm the cryovial rapidly by immersion in a
shaking (for increase in heat exchange) water
bath (37°C) for 1–2 min.
Fast Cooling Rates Regime of Freezing
(~400 grad/min)
• Place the cryovials with cultures (spore
suspension) in special containers; thoroughly
fixate a position of every vial.
• Immediately place cryovials with cultures in
the vapour phase of liquid nitrogen or in ultralow temperature freezer.
Programmed Regime of Freezing
The First Protocol
• Place 18 cryovials with cultures (mycelial
plugs) in a special container “NALGENE™”
(Cryo 1°C Freezing Container, Cat. No. 51000001).
• Fill the container with 250 mL of isopropanol.
• Place the container in a mechanical freezer
(−80°C). Temperature in this container
decreased by 1°C/min.
• When temperature achieves −70°C, transfer
cryovials in special container (thoroughly
fixate a position of every vial), and place in
the vapour phase of liquid nitrogen or in the
ultra-low temperature freezer.
Control of Viability
• To estimate the viability of fungal culture
before cryopreservation, place either one volume of the suspension (0.2 mL) or 4 plugs
under optimal growth conditions on a suitable
medium. This procedure is carried out under
sterile aerobic conditions.
• To estimate the viability of fungal culture after
cryopreservation, sterilize the thawing cryovials surface by wiping with 70% (v/v) ethanol. Asepticaly transfer the contents (spore
suspension or mycelial plugs) using a Pasteur
pipet or a transfer needle onto a suitable
growth medium. This procedure is carried out
under sterile aerobic conditions.
Protocol of Freeze-Drying
Preparation of Ampoules
Glass tubes (gray glass, diameter 7 mm, length
110 mm):
• Wash successively with detergent, tap water,
and distilled water.
• Dry.
3
Long-Term Preservation of Fungal Cultures in All-Russian Collection of Microorganisms (VKM)…
• Plug loosely with cotton wool to a depth of
1 cm.
• Label with an index, a collection number of a
strain, and a date of freeze-drying (month,
year).
• Sterilize in dry oven at 160°C for 2 h.
Preparation of Lyoprotectant Agent
• Pour 5 mL 10% (v/v) skimmed milk into each
12-mL glass tube.
• Sterilized by autoclaving at 105°C for
30 min.
• Stored at +5°C for no longer than a month.
Preparation of Cultures
• Grow sporulating fungal cultures on slant agar
under optimal growth condition and on suitable medium (www.vkm.ru).
• Wash off spores from agar surface with 5 mL
of skimmed milk.
• Titer of spores’ suspension should be more,
than 106 spores/mL.
Filling of Ampoules
• Add 0.2-mL aliquots of suspension to each
ampoule using a Pasteur pipet. This procedure
is carried out under sterile aerobic condition.
The First Stage (Primary Drying)
• Transfer the ampoules to the spin freezedrier.
• Freeze (temperature in a refrigerator −45°C)
under the reduced pressure of the ambient gas
during centrifugation (30 min).
• Dry via water sublimation (temperature of a
freeze dryer is −45°C) in vacuum (from
4 × 10−2 to 6 × 10−2 mbar) till the moisture level
achieves 5–10%.
• Duration of the first stage is 3 h.
• Switch off the vacuum pump after the first
stage. The system is filled with gas; the
ampoules are removed from the centrifuge.
Preparation of Ampoules for the Second
Stage of Freeze-Drying
• Constrict ampoules to diameter of 2–3 mm
using an air/gas torch with horizontal flame
preventing overheating of cultures just below
63
the cotton wool plug (approximately 50 mm
from the ampoule bottom)
The Second Stage (Secondary Drying)
• Attach the constricted ampoules via rubber
tubes to the manifold connecting with the vacuum pump.
• Drying (vacuum 100 mm) till the moisture
level reach 2%.
• Duration of the second stage is 2.5 h.
• Seal the ampoules across the constriction
using an air/gas torch.
Vacuum Control
• Immediately after sealing, test vacuum in
ampoules using a high-voltage spark tester.
Control of Culture Viability
• To estimate the viability of fungal strains
prior to freeze-drying one volume of the spore
suspension (0.2 mL) place under optimal
growth condition on suitable medium. This
procedure is carried out under sterile aerobic
conditions.
• To estimate the viability of fungal culture after
freeze-drying, test ampoules after 24 h storage.
• This procedure is carried out under sterile aerobic conditions.
• Sterilize a control ampoule’s surface with 70%
ethanol and open ampoules using a cutter.
• Reconstitute the dried suspension with sterile
tap water (0.2–0.3 mL) using a Pasteur
pipet.
• After 30 min (when rehydration is complete)
transfer suspension under optimal growth condition on suitable medium.
Storage
• Store the ampoules d at +5–8°C in the dark.
Protocol of Drying in Sterile Soil
Preparation of Sterile Soil
• Place 5 g of finely cultivated (garden) soil into
12-mL glass tube.
• Sterilized by autoclaving, at 121°C for 30 min
for three consecutive days.
64
Preparation of Cultures
• Grow sporulating fungal cultures on slant agar
under optimal growth condition and on suitable medium (www.vkm.ru).
• Wash off spores from agar surface with 5 mL
of sterile tap water.
• Titer of spores’ suspension should be not less
than 106 spores/mL.
Soil Inoculation
• Add 1 mL spore suspension to glass tubes with
sterile air dry soil (moisture is under 20%).
• Incubate at room temperature till soil dry up
(near 1 month).
• Store in the refrigerator at 4–7°C.
Control of Viability
• Transfer a few grains of soil onto fresh agar
medium, add a little water and incubate under
optimal conditions.
Result
The real storage time estimates obtained in VKM
are given in Table 3.1. They are not final data: the
cultures are still being stored, and we expect to
get longer storage times later on. Some cells of
the table are empty; this is the case if the culture
is not stored this method.
Conclusion
The conservation techniques used in VKM presents effective preservation of the stock of
filamentous fungi from different taxonomic
groups. The possibility and practical time estimates of secure long-term storage of fungal cultures belonging to 1,221 species and 424 genera
was shown. The represented information could
be used as a reference for researchers intending
to maintain pure cultures of microorganisms for a
long time. The data produced are also accessible
online on the VKM Web site.
Acknowledgements We thank Ludmila Evtushenko for
constructive comments on the manuscript.
S.M. Ozerskaya et al.
This work was supported by grants of the program
MCB of the Russian Academy of Science and the Ministry
of Education and Science of the Russian Federation
(N. 16.518.11.7035).
References
1. OECD best practice guidelines for biological resource
centers. OECD; 2007. 115p.
2. Glyn NS, Day JGD (2007) Long-term ex situ conservation of biological resources and the role of biological resource centers. In: Day JG (ed) Methods in
molecular biology, vol 368. Humana Press, Totowa,
pp 1–14
3. Smith D (1998) Culture and preservation. In:
Hawksworth DL, Kirsop BE (eds) Living resources
for biotechnology. Filamentous fungi. Cambridge
Univ. Press, UK, pp 75–99
4. Guidelines for the establishment and operation of
collections of cultures of microorganisms. 3rd ed.
WFCC; 2010. 19p. http://www.wfcc.info/pdf/
Guidelines_e.pdf.
5. Smith D, Onions AHS (1994) The preservation and
maintenance of living fungi, 2nd edn. AB International,
Wallingford, UK, p 132
6. Nakasone KK, Peterson SW, Jong S-C (2004)
Preservation and distribution of fungal cultures. In:
Mueller GM et al (eds) Biodiversity of fungi. Inventory
and monitoring methods. Elsevier/Academic Press,
Amsterdam, pp 37–47
7. Ryan MJ, Smith D (2007) Cryopreservation and
freeze-drying of fungi employing centrifugal and
shelf freeze-drying. In: Day JG (ed) Methods in
molecular biology, vol 368. Humana Press, Totowa,
pp 127–140
8. Smith D, Thomas VE (1998) Cryogenic light microscopy and the development of cooling protocols for the
cryopreservation of filamentous fungi. World J Microb
Biotech 14:49–57
9. Sidyakina TM (1988) Methods of preservation of
microorganisms. In: Veprintzev BN (ed) Konservatziya
geneticheskikh resursov. ONTI NTzBI AN USSR,
Pushchino, p 59 (In Russian)
10. Ivanushkina NE, Kochkina GA, Eremina SS,
Ozerskaya SM (2010) Experience in using modern
methods of long-term preservation of VKM fungi.
Mikol Phytopathol 44:19–30 (In Russian)
11. Ryan MJ, Smith D, Jeffries P (2000) A decision-based
key to determine the most appropriate protocol for the
preservation of fungi. World J Microb Biotech
16:183–186
12. Milosevic MB, Medic-Pap SS, Ignatov MV, Petrovic
DN (2007) Lyophilization as a method for pathogens
long term preservation. Proc Nat Sci Matica Srska
Novi Sad 113:203–210
13. Sundari SK, Adholeya A (1999) Freeze-drying vegetative mycelium of Laccaria fraterna and its subsequent regeneration. Biotechnol Tech 13:491–495
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14. Raper KB, Fennell DI (1965) The genus Aspergillus.
The Williams and Wilkins Company, Baltimore,
p 686
15. Suprun TP (1965) Preservation of microscopic fungi in
sterile soil. Mikrobiologiya 34:539–545 (In Russian)
16. Belyakova LA, Lavrova LN, Kudryavtzev VI (1967)
Preservation of fungal cultures. Metody khranenija
kollekcionnykh kul’tur mikroorganizmov. M.: Nauka,
p 7–55 (In Russian)
65
17. Atkinson RG (1953) Survival and pathogenicity of
Alternaria raphani after five years in dried soil
cultures. Can J Bot 31:542–546
18. Hine RB (1962) Saprophytic growth of Fusarium
oxysporum f. niveum in soil. Phytopathology
52:840–845
19. Shearer BL, Zeyen RJ, Ooka JJ (1974) Storage and
behavior in soil of Septoria species isolated from
cereals. Phytopathology 64:163–167
4
Fungal Specimen Collection
and Processing
Anthonia O’Donovan, Vijai Kumar Gupta,
and Maria G. Tuohy
Abstract
The study of fungi relies, in part, on the axenic culture of isolates. Because
so many fungi are found in nature in close proximity to each other, and
many other organisms, study of their structure and function relies on the
ability to grow and maintain a pure culture of the fungi. This chapter
describes the methodology used to isolate a fungus that is present in abundance in a soil sample and covers many techniques that are widely used in
microbiology laboratories.
Keywords
Fungi • Isolation • Culture • Soil • Spread plate • Direct culture • Storage
Introduction
A. O’Donovan (*)
School of Natural Sciences, National University
of Ireland Galway, University Road, Galway, Ireland
e-mail: anthonia.odonovan@nuigalway.ie
V.K. Gupta
Molecular Glycobiotechnology Group,
Department of Biochemistry, School of Natural Sciences,
National University of Ireland Galway,
University Road, Galway, Ireland
Assistant Professor of Biotechnology, Department
of Science, Faculty of Arts, Science & Commerce,
MITS University, Rajasthan, India
M. G. Tuohy
Molecular Glycobiotechnology Group, Department
of Biochemistry, School of Natural Sciences,
National University of Ireland Galway,
University Road, Galway, Ireland
Fungi, including yeasts and molds, are ubiquitous
in nature and have been recovered from
diverse, remote, and extreme environments.
Approximately 100,000 fungal species have
been described to date. Over the last decade,
approximately 1,200 new species of fungi have
been described in each year. It is now estimated
that there may be from 1.5 to 5 million extant
fungal species [1]. Fungi are eukaryotic microbes
with a filamentous growth form. A wide variety
of fungi can be isolated from soil and cultivated
on media in the laboratory. Different media
will encourage the growth of different types of
microbes through the use of inhibitors and
specialized growth substrates [2].
Fungi are often isolated using nonselective
agar medium. This allows the isolation of the
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_4, © Springer Science+Business Media, LLC 2013
67
68
maximum number of fungal taxa from a sample
(it should be noted that all media used in microbiological laboratories are selective to some
degree or another; truly nonselective media do
not exist [3]). So-called nonselective media are
only media and incubation conditions designed
to isolate as large a part of the microbiota in soil
as is possible. The least selective media today
isolate maybe 5–15% of the fungal population of
soils. The media used for fungi are different from
the media recommended for bacteria [4].
Saprotrophic fungi can be subcultured on
media containing nutrients appropriate to their
growth and development. Several different types
of media have been used successfully. The organic
and mineral fractions of media are designed to
supply nutrients similar to or commonly found in
the environment of the fungus. Some commonly
used media consist of fruit or vegetable extracts
mixed with sugars and agar, and set in Petri
dishes. Other commonly used materials include
soil (soil agar), potato (PDA), tomato plus other
vegetables (V8 JUICE Agar), Malt extract (MA),
and dung (DUNG AGAR). These can be more
highly defined by replacing the organic component with known organic materials including
nutrient dextrose (NDY) and sabouraud dextrose
(Sabouraud Agar; [3, 4]).
Specifically targeted fungi can be subcultured
using selective media. Routine methods of providing selective media include the addition of
specific compounds required by an organism as a
nutrient source or the omission of compounds
required by most other organisms [5, 6]. Changing
some physical properties of the media can also
favor growth of specific organisms (e.g., lowering the pH allows fungal growth to prevail over
bacterial growth). Incubation temperature is
another common method to encourage growth of
target organisms or to deter growth of contaminating microbes (e.g., spore-forming fungi can be
isolated by the use of their ability to withstand
high temperature). Altering conditions such as
water content and the presence or absence of light
along with providing aerobic or anaerobic conditions may also help in promoting the growth of a
particular organism [3, 4, 7].
Many media are available from commercial
sources (e.g., from Difco or Oxoid), which have a
A. O’Donovan et al.
more consistent quality than those prepared from
the raw materials each time.
The inclusion of antibiotics in media is a commonly used method to suppress the growth of bacteria. As most antibiotics are denatured by heat, the
antibacterial agents are usually added to sterilized
molten agar, that is just above setting temperature
(hand warm, but not hot to touch). Antibiotics such
as penicillin, streptomycin, and tetracycline are
commonly used, either alone or more commonly,
in combination. Chloramphenicol can be autoclaved and is added during preparation [3, 7].
It is possible to design very selective media
using combinations of the aforementioned techniques. Almost any physiological group of organisms can, in theory, be cultured selectively.
A single stage isolation from a soil sample can be
changed to a multistage isolation process by replica plating. An example of this includes the
transfer of individual colonies in their original
orientation on the plate by pressing a strip of sterile tape onto the surface of the plate, removing a
small sample of each colony and pressing it onto
the surface of a fresh plate. This can be a different
growth medium so that only a part of the original
population is able to grow on the new medium.
Using this technique, progressive selective media
can be used to isolate microorganisms with combinations of properties [3, 4, 8].
Some general observations:
• Fungi grow better on media with a high C:N
ratio (bacteria grow better on low C:N ratio
media).
• More fungi are isolated on low pH media
(bacteria are isolated using neutral pH
media).
• Spores from common fungi such as Aspergillus,
Mucor, and Penicillium in soil grow rapidly
on many fungal isolation media and prevent
growth of the slower growing fungi.
• Many fungi produce antibiotic compounds on
plates and prevent growth of other
microorganisms.
• Some microbes are very difficult to culture at
all on media in the laboratory. They may be
very slow growing or very sensitive to components of normal media [9].
Our knowledge of soil fungi is derived primarily from dilution and plating techniques (which
4
Fungal Specimen Collection and Processing
are described in the following sections). These
methods have limitations in that they are biased
in favor of rapidly growing and sporulating organisms [10, 11] and consequently most of the fungi
identified by these techniques are Fungi Imperfecti
(Pencillium and Aspergillus). It is extremely
difficult to grow soil basidiomycetes on solid
media in the lab [11], but their abundance in soil
can be revealed by microscopy. Nevertheless,
dilution plate techniques are still a popular and
useful tool for studying the relative abundance of
culturable populations.
Sample Collection
Fungi are ubiquitous in their occurrence but are
inconspicuous because of the small size of their
structures, and their cryptic lifestyles in soil, on
dead matter, and as symbionts of plants, animals,
or other fungi. Common sources for their isolation include lakes, river muds, and soils.
Soils are discontinuous heterogenous environments that contain large numbers of diverse
organisms. The main sampling problems in soil
microbiology are usually a result of the complexity of the medium being sampled. If a method
requires a “generalized” sample of the soil, the
problem is to determine what soil horizons to
sample. How many samples are required (to estimate variability), exactly where to take the samples (to determine spatial variation), and how
frequently samples should be taken (to determine
temporal variation) also have to be determined
and can greatly influence findings. These problems are often interrelated, for example, a larger
sample subdivided into smaller ones after mixing is different to many small independent samples. The first will show experimental or
procedural error, while the second will show differences due to natural field variation as well as
procedural errors. Therefore, many factors are to
be considered when sampling soil [9]. Microbial
populations also vary considerably with soil
type, soil depth, moisture levels, use of pesticides, climate, etc.
Once a sampling site is selected, collect the
desired soil samples, ensuring no contamination
of the samples. Store the samples at 4°C until
69
ready to be cultured. Samples should be cultured
within 72 h of collection if possible to eliminate
the storage effects on the microbial population.
Air dry the samples to remove moisture and to
allow for sieving of the soil, which removes large
particulate, litter and debris.
Materials
1. Soil.
2. Balance.
3. The media: Aureomycin-rose Bengalglucose-peptone agar: glucose, peptone,
potassium dihydrogen phosphate (KH2PO4),
magnesium sulfate (MgSO4.7H2O), rose
Bengal, agar, Aureomycin hydrochloride,
and reagent grade water.
4. Hot water bath (45 °C) to keep molten agar.
5. Sterile saline blanks for dilution (9 and
90 mL) in dilution bottles and test tubes.
6. Sterile pipettes (P1000 and P200) and sterile
tips.
7. Vortexer.
8. Spreaders for spread-plating (made from
glass pipette).
9. 70% ethanol (in squirt bottles and dishes).
10. Bunsen burner.
11. Inoculating loops (for transferring colonies).
Methods
Sample Preparation
Sample Dilution
1. To prepare the soil sample for culturing, appropriate dilutions of a suspension must be prepared: Add 10 g of soil sample (air dried and
sieved) to 90 mL of sterile saline. This will create a 1/10 dilution of the soil sample, and the
container should be labeled appropriately.
2. Vortex the solution for at least 1 min. The
intent of the stirring is to suspend the organisms in the solution.
3. Allow the soil particulate matter to settle for
up to 15 min. Decant the liquid portion into
test tubes labeled with the appropriate sample
and the dilution (1/10).
70
4. Transfer 1 mL of the 1/10 dilution into a test
tube containing 9 mL of sterile saline using a
sterile pipette. This will create a 1/100 dilution of the sample. Vortex the test tube until
the sample is thoroughly mixed. Ensure the
sample is labeled with the appropriate sample
and dilution.
5. Transfer 1 mL of the 1/100 dilution into a test
tube containing 9 mL of sterile saline using a
sterile pipette. This will create a 1/1,000 dilution of the sample. Vortex the test tube until
the sample is thoroughly mixed. Ensure the
sample is labeled with the appropriate sample
and dilution.
6. Repeat the above steps (4–5) to create additional dilutions if required. Dilutions for fungal
cultures normally need not exceed 1/10,000.
A. O’Donovan et al.
2.
3.
4.
Preparing Plates
5.
1. An example of one type of media is Aureomycinrose Bengal-glucose-peptone agar. To make
this agar, dissolve 10 g glucose, 5 g peptone,
1 g potassium dihydrogen phosphate (KH2PO4),
0.5 g magnesium sulfate (MgSO4.7H2O),
0.035 g rose Bengal, and 20 g agar in 800 mL
reagent grade water. The pH should be 5.4.
2. Sterilize by autoclaving at 105 °C for 30 min.
Then keep molten in a 45 °C waterbath.
3. Dissolve 70 mg Aureomycin hydrochloride in
200 mL reagent grade water, sterilize by
filtration.
4. Add the sterilized Aureomycin hydrochloride
solution to the cooled (42–45 °C) agar base.
5. Pour 25 mL molten agar into sterile petri dishes
(100 × 15 mm) and allow agar to solidify. Poured
plates can be stored at 4 °C for up to 4 weeks.
Note: Many media are available from commercial sources. Commercial sources have a more
consistent quality than those prepared from the
raw materials each time.
Culturing
Spread Plate
1. Using a sterile pipette transfer 100 mL of the
sample or appropriate sample dilution onto the
6.
agar surface (if you do not know which dilution
is suitable, then several need to be plated out).
(Vortex sample before taking the aliquot.)
Use the spreader (bent glass rod), which has
been flame-sterilized, to spread the dilution
over the surface as evenly as possible to ensure
proper distribution (Note: cool the spreader
before spreading).
Flame-sterilize the rod between different
dilution factors and media or if it comes into
contact with any nonsterile surface. (If dilution factors are spread in order of decreasing
dilution—that is, 1/10,000, 1/1,000, 1/100,
etc.—spreaders need not be resterilized
except with changes of media.) Allow the
moisture to be absorbed into the agar before
the incubation.
Label plates with date, organism, dilution factor, type of agar, and incubation temperature.
Prepare triplicate plates for each dilution.
Invert plates and incubate at the selected temperatures. Observe plates after day 2 and until
day 5/7. Slow growing fungi may not produce
noticeable colonies until day 6 or 7.
After the microbial colonies are readily visible
(2–7 days), count the number of colonies on
each plate and calculate the average number of
fungi in the soil sample. (Plates can be stored
at 4 °C for not longer than 24 h before counting if required).
Results
Choose a plate with a dilution that gives the most
reliable counts for your calculations. The optimal
maximal number of colonies per plate is 100. If
there are more than 150 colonies per plate the
result is recorded as TNTC (too numerous to
count) and a plate with a lower number of colonies (a lower sample dilution) is selected. If there
are no colonies on any of the plates, the results
are recorded as <1 for the highest dilution.
Note: Interpretation of cultured sample data is
complicated by two factors:
1. Certain species only grow well on certain agar
media, and, therefore, may be present but not
culturable because an inappropriate agar was
used. Other species are not culturable at all.
4
Fungal Specimen Collection and Processing
2. Fungi grow at different rates, so slow-growing
fungi may be over-run by fast-growing fungi,
and, therefore, not be apparent in the culture.
Therefore, to perform culturable sampling
correctly, three or more types of agar may need to
be used for each sample.
It is important to note that quantification of
fungi is different to bacteria because a fungal
colony can form from a spore (single cell), a cluster of spores, or from a mycelial fragment (containing more than one viable cell). It is assumed
that each fungal colony developed in a laboratory
culture originates from a single colony forming
unit, which may or may not be a single cell.
71
sample containing what appears to be one kind of
fungus onto the agar: A small fragment of soil is
placed on an agar plate and incubated. Then the
emergent hyphal tips are subcultured. Transfer
these samples to fresh agar plates and incubate to
develop a pure culture.
Storage of Fungi
It is expensive to continually subculture isolated
fungal samples. Furthermore, the fungi may
mutate
during
continuous
subculturing.
Preservation of fungi is essentially reducing the
metabolic rate to the slowest possible. This can
be achieved in several different ways.
Identification
Fungal colonies are identified to genus or species, if possible, using a combination of colony
macroscopic characteristics (e.g., color, morphology, growth rate) and microscopic characteristics (e.g., spore shape, size, color, hypha
morphology [12–14]). Referring to definitive
texts and Web sites regarding fungal identification
is recommended.
Isolate a Pure Culture
Many cultures may consist of multiple types of
fungi. To isolate a single fungus, a method to
specifically select the target fungus for further
fungal growth can be chosen. An example of such
a method is to select one colony if possible or
sweep the end of a hyphal tip and aseptically
plate the sample out onto a fresh agar plate.
Selective plates may be used to culture specific
fungi. Once the target fungus has been isolated in
this manner, further culturing onto nonselective
agar without antibiotic can be conducted to ensure
a pure culture has been achieved.
Direct Culture Methods
Some fungi may not be detected using the soil
dilution/spread plate method previously given. In
this case it is possible to directly inoculate the
fungal sample by directly positioning the soil
Storage in Water
Plugs of agar containing the fungal culture are
placed in vials containing only sterile water. The
vials are lidded and stored at 10 °C for up to
5 years. This is a remarkably useful and cheap
technique. (This is not recommended for many
Basidiomycota).
Cold Temperature
Place the isolate on a slope or plate and store at
4 °C. Some fungi stored at cold temperatures
remain viable for up to 4 years.
Under Oil
The fungus is subcultured onto low sugar media
(e.g., one-sixth strength NDY) in a lidded tube.
The culture is incubated to allow the isolate to
cover the surface of the agar. The fungus is then
covered with sterile mineral oil. The tubes are
kept at room temperature. Fungi may be stored
for up to a year. The oil slows access of oxygen to
the culture. However, it is crucial the culture is
covered by at least 1 cm of oil at all times and that
the lid is kept clean. Contaminating fungi can
grow on the top of the tube if it has nutrients on it.
Also, moderate changes in air pressure lead to the
movement of air in and out of the tube.
Contaminants present in the air can lodge on the
surfaces of the lidded tube. Therefore, topping the
oil to just below the lid is recommended. Flaming
the surfaces before subculturing is essential [15].
72
Freeze Drying or Lyophilization
Cultures are placed in a lyophilization tube,
cooled, and then freeze dried. Fungi preserved in
this way can be stored for up to 20 years. The
process is commonly used for spore-forming
Deuteromycetes. It is less successful for nonspore-forming fungi.
Low-Temperature Storage
Some fungi can be stored for many years when
placed in 10–25% sterile glycerol in water in
sealed vials that are then placed in liquid nitrogen
or stored at – 70 °C. The fungi may need to be
taken in stages through the cooling process. It is
important to experiment with the correct glycerol
dilution as some fungi will dessicate in 10%
glycerol [15].
References
1. Hibbett DS, Ohman A, Glotzer D, Nuhn M, Kirk P,
Nilsson RH (2011) Progress in molecular and morphological taxon discovery in Fungi and options for
formal classification of environmental sequences.
Fungal Biol Rev 25:38–47
2. Hurst CJ, Crawford RL, Garland JL, Lipson DA,
Mills AL, Stetzenbach LD (2007) Manual of environmental microbiology, 3rd edn. ASM Press, Washington,
DC. 20036–2904, USA ISBN 978-1-55581-379-6
3. Maier RM, Pepper IL, Gerb CP (2000) Environmental
microbiology. Academic Press, San Diego, USA
A. O’Donovan et al.
4. http://www.scribd.com/doc/16572739/PROTOZOAAND-FUNGI-CULTURING-IN-THE-LAB
5. Hunter-Cevera JC, Fonda ME, Belt A (1986) Isolation
of cultures. In: Demain AL, Solomon NA (eds)
Manual of industrial microbiology and biotechnology.
American Society for Microbiology, Washington, DC,
pp 3–23
6. Seifert KA (1990) Isolation of filamentous fungi. In:
Labeda DP (ed) Isolation of biotechnological organisms from nature. McGraw-Hill Publishing Co., New
York, pp 21–51
7. Collins CH, Grange JM, Lyne PM, Falkinham JO
(2004) Microbiological methods. Hodder Arnold,
London, UK. ISBN 9780340808962
8. Nautiyal CS, Dasgupta S, Varma A, Oelmüller R
(2007) Screening of plant growth-promoting rhizobacteria advanced techniques in soil microbiology.
Springer, Berlin, Heidelberg
9. http://wvlc.uwaterloo.ca/biology447/modules/module8/soil/chapter3Soil446.htm
10. Bonito G, Isikhuemhen OS, Vilgalys R (2010)
Identification of fungi associated with municipal compost using DNA-based techniques. Bioresour Technol
101:1021–1027
11. Thorn RG, Reddy CA, Harris D, Paul EA (1996)
Isolation of saprophytic basidiomycetes from soil.
Appl Environ Microbiol 62:4288–4292
12. de Hoog GS, Guarro J, Gene J, Figueras MJ (2004)
Atlas of clinical fungi. Centraalbureau voor
Schimmelcultures, Utrecht, The Netherlands
13. Pitt JI, Hocking AD (1997) Fungi and food spoilage.
Academic press, Sydney
14. Samson RA, Hoekstra ES, Frisvad JC (2004)
Introduction to food and airborne fungi. Centraalbureau
voor Schimmelcultures, Utrecht, The Netherlands
15. http://bugs.bio.usyd.edu.au/learning/resources/
Mycology/Growth_Dev/axenicCulture.shtml
5
Chemical and Molecular Methods
for Detection of Toxigenic Fungi
and Their Mycotoxins from Major
Food Crops
S. Chandra Nayaka, M. Venkata Ramana,
A.C. Udayashankar, S.R. Niranjana,
C.N. Mortensen, and H.S. Prakash
Abstract
Mycotoxins are the secondary metabolites produced by certain molds on a
wide range of agricultural commodities and are closely related to human
and animal food chains. Mycotoxins are capable of causing disease in
humans and other animals, and their detection is largely dependent on the
sample matrix and the type of fungus causing their contamination. The
strict regulations on trade of contaminated grains and seeds and other produce in industrial countries lead to economic burdens on farmers. In developing countries, the situation is aggravated where regulations may be
nonexistent or not enforced and where consumption of home-grown cereals leads to a wide exposure to toxins. Important mycotoxins that occur
quite often in food are deoxynivalenol/nivalenol, trichothecenes, zearalenone, ochratoxin A fumonisins, and aflatoxins. High concentrations of
mycotoxins such as aflatoxins are consumed by humans in areas of the
world with higher-than-average levels of liver cancer, childhood malnutrition, and disease. This chapter introduces rapid, robust, and user-friendly
protocols currently applied in the identification of toxigenic fungi and
important mycotoxins.
S.C. Nayaka • A.C. Udayashankar
S.R. Niranjana • H.S. Prakash (*)
Department of Studies in Biotechnology, Asian Seed
Health Centre, University of Mysore Manasagangotri,
Mysore, Karnataka 570 006, India
e-mail: hsp@appbot.uni-mysore.ac.in
M.V. Ramana
Microbiology Division, Defence Food Research
Laboratory, Siddarthanagar, Mysore,
Karnataka 570 011, India
C.N. Mortensen
Department of Agriculture and Ecology, University
of Copenhagen, Thorvaldsensvej 40, DK-1871
Frederiksberg C Copenhagan, 2630 Taastrup, Denmark
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_5, © Springer Science+Business Media, LLC 2013
73
74
S.C. Nayaka et al.
Keywords
Mycotoxins • Fungi • Polymerase chain reaction (PCR) • Thin-layer chromatography (TLC) • High-performance liquid chromatography (HPLC)
Introduction
The term “mycotoxin” combines the Greek word
for fungus “mykes” and the Latin word “toxicum” meaning poison. Mycotoxins have received
considerable attention, especially over the last
few decades. The problem related to mold damage and the hazard of consuming damaged grains
have been recognized since historical times. The
term “mycotoxin” is usually reserved for the
toxic chemical products formed by a few fungal
species that readily colonize crops in the field or
after harvest and thus pose a potential threat to
human and animal health through the ingestion
of food products prepared from these commodities [1].
The possibility of human diseases occurring
as a result of the consumption of mold-damaged
rice and wheat was raised in Japan and other
Asian countries during the first half of this
century. Awareness of risks from eating overwintered millet was reported in the USSR [2].
However, the serious worldwide concern about
mycotoxins began in the early 1960s after it was
discovered in the United Kingdom that Turkey
“X” disease is caused by aflatoxins. More than
300 mycotoxins have been identified, although
only around thirty with toxic properties that are
genuinely of concern for human beings or animals were reported by the French Agency of
Food, Environmental and Occupational Health
and Safety [3]. Some mycotoxins are rather rare
in occurrence; others—such as aflatoxin, ochratoxin, fumonisins, and trichothecenes—are quite
common in some years. The molds primarily
responsible for producing mycotoxins are
Aspergillus, Fusarium, and Penicillium spp.
Occurrence of mycotoxins in food and animal
feed often exhibits a geographical pattern; for
example, Aspergillus species meet optimal
conditions in tropical and subtropical regions,
whereas Fusarium and Penicillium species are
adapted to the moderate climate of North America
and Europe [4]. The toxins can be produced in
major food crops like, maize, wheat, sorghum,
rice, soybeans, peanuts, and other food and feed
crops in the field, during transportation, or
improper storage. Moreover, in animals consuming contaminated feed, mycotoxins can deposit in
different organs and also subsequently affect
food of animal origin (e.g., meat, eggs, milk, and
milk products). Worldwide trade with food and
feed commodities has resulted in a wide distribution of contaminated material [5]. One of the
characteristics of mycotoxins is that they can
exude toxic properties in minute quantities; thus,
sensitive and reliable methods are required for
their detection and quantification, which generally involves sophisticated sampling, sample processing, extraction, and assay techniques.
Different methods have been applied in the
detection and quantification of mycotoxins from
food and feed samples, including ELISA (enzymelinked immunosorbent assay), immunoaffinity
cartridge, solid-phase ELISA, and selective adsorbent mini-column procedures [6]. TLC (thin-layer
chromatography) and HPLC (high-performance
liquid chromatography) [7] are more accurate for
quantification of mycotoxins in food and feedstuffs’ produce. Under practical storage conditions monitoring for the occurrence of fungi is
often conducted; however, in practice, it is difficult
to distinguish several toxigenic fungal species
from their close relatives, and accurate
identification based on traditional methods is very
difficult owing to their genetic variation and high
morphological similarity.
The conventional scheme of isolation and
identification of toxigenic fungi from food
samples is cumbersome and requires skilled
personnel to achieve proper identification. Even
5
Chemical and Molecular Methods for Detection of Toxigenic Fungi…
with taxonomical expertise, identification is
commonly difficult regarding some fungi genus
that contains a large number of closely related
species [8]. Robust DNA-based tools often offer
accurate, rapid, and sensitive identification and
characterization of species (e.g., Fusarium) that
belong to a complex genus [9]. The application of
molecular biology techniques is an alternative to
cumbersome and time-consuming conventional
culture methods for precise identification of toxigenic fungal species before they can enter the
food chain. The polymerase chain reaction (PCR)
assay has allowed rapid, specific, and sensitive
detection of toxigenic species without the need
for prior growth of the organisms. The traditional
molecular markers are mainly based on ribosomal
DNA, b-tubulin, and calmodulin genes or have
been based on anonymous DNA sequences.
These DNA sequences are obtained from an
unbiased sampling of genomic DNA, and these
may or may not contain functional genes involved
in toxin production [10].
Developing markers from anonymous
sequences requires comparative analyses among
related species of DNA profiles generated from
randomly amplified fragments by using RAPD
(random amplified polymorphic DNA) or AFLP
(amplified fragment length polymorphism). In
the last decade, numerous PCR assays have been
developed for rapid detection and differentiation
of toxigenic and nontoxigenic fungi from major
commodities by using specific genes associated
with mycotoxin biosynthesis [11].
75
Fig. 5.1 Structure of aflatoxin B1
Fig. 5.2 Structure of ochratoxin A
Reye’s syndrome, kwashiorkor, and hepatitis [12].
Aflatoxins can also affect the immune system.
A. flavus infects many of our food crops, such as
nuts, grains, and culinary herbs. Primary economic concerns are infestations that occur in corn
and peanuts. The major aflatoxins consist of
aflatoxins B1 (Fig. 5.1), B2, G1, and G2.
Ochratoxin
Some Important Mycotoxins
Aflatoxins (Aspergillus spp.)
Aflatoxins are chemical derivatives of difurancoumarin, mainly produced by Aspergillus
flavus/A. parasiticus. Aflatoxins have been implicated in subacute and chronic effects in humans.
These effects include primary liver cancer,
chronic hepatitis, jaundice, hepatomegaly, and
cirrhosis through repeated ingestion of low levels
of aflatoxin. It is also considered that aflatoxins
may play a role in a number of diseases, including
Ochratoxin A is the most important and most
commonly occurring structurally related group of
compounds; it is often abbreviated to OTA or OA
(Fig. 5.2). Ochratoxin A is the major mycotoxin
of this group, and it is an innately fluorescent
compound produced primarily by Aspergillus
ochraceus and Penicillium verrucosum [13].
Ochratoxin A is a potent toxin that affects mainly
the kidneys, in which it can cause both acute and
chronic lesions. Ochratoxin A is a potent teratogen in mice, rats, hamsters, and chickens, and a
nephrotoxic effect has been demonstrated in all
mammalian species.
76
S.C. Nayaka et al.
Fig. 5.3 Structure of
fumonisin B1
Fumonisins
The fumonisins are a group of nonfluorescent
mycotoxins—FB1 (Fig. 5.3), FB2, and FB3 being
the major entities—produced primarily by
Fusarium verticillioides and F. proliferatum [13],
Fusarium nygamai, as well as Alternaria alternata f. sp. lycopersici. Fumonisins are thought to
be synthesized by condensation of the amino
acid alanine into an acetate-derived precursor.
Numerous species-specific diseases have been
attributed to fumonisin-contaminated feed, including leukoencephalomalacia in horses and pulmonary edema and hydrothorax in swine [14]. These
compounds have been shown to have carcinogenic potential in animal models and are the only
known inhibitors of ceramide kinase, a key
enzyme involved in inflammatory cascades.
Deoxynivalenol
Deoxynivalenol, also known as DON or vomitoxin, is one of about 150 related compounds
known as the trichothecenes that are mainly produced by Fusarium graminearum and, in some
geographical areas, by F. culmorum (Fig. 5.4)
[15]. These two species are important plant
pathogens and cause Fusarium head blight in
wheat and Gibberella ear rot in maize. Toxicity of
deoxynivalenol is characterized by vomiting,
particularly in pigs, feed refusal, weight loss, and
diarrhea. A study reporting human food poisoning by infected wheat containing deoxynivalenol
in India showed a range of symptoms, including
abdominal pains, dizziness, headache, throat
irritation, nausea, vomiting, diarrhea, and blood
in the stool [16].
Fig. 5.4 Structure of deoxynivalenol
The potential presence of toxins in the food
supply means that expensive testing and remedial
actions are necessary to assure that they do not
reach dangerous levels in our food. This testing
and losses in crop quality and yield associated
with these fungal diseases are estimated to cost
agriculture billions of dollars annually, and the
presence of fungal toxins in our crops places the
competitiveness of our agricultural exports at
risk. The presence of mycotoxins is unavoidable;
therefore, testing of raw materials and products is
required to keep our food and feed safe. The presence of mycotoxins in food crops is a serious and
common quality problem that has become more
obvious as a result of the research of recent years.
Several chemical and biological detection systems exist for the determination of mycotoxins.
Biological assays were used when analytical and
methods were not available for routine analysis
because biological assays are qualitative and
often are nonspecific and time-consuming. Various
analytical methods for mycotoxin analysis have
been developed, such as TLC, HPLC, and HPTLC
[7]. Many research laboratories have also adopted
molecular detection methods for rapid and accurate
5
Chemical and Molecular Methods for Detection of Toxigenic Fungi…
detection of toxigenic molds from major food
crops.
This chapter presents information on the general protocols adopted for detection of important
toxigenic Aspergillus, Penicillium, and Fusarium
species by PCR and their mycotoxins mainly by
TLC and HPLC.
Materials
(See Note 1)
77
18. UV transilluminator and camera suitable for
photographing agarose gels (e.g., Syngene
Gene Genius Bioimaging System).
Basic Equipment Required
for Polymerase Chain Reaction
1.
2.
3.
4.
5.
6.
Thermal cycler.
Micropipettes.
Agarose gel electrophoresis unit.
Centrifuge.
UV-gel documentation chamber.
Eppendorf tubes.
DNA Extraction
1. Sterile distilled water.
2. 1× lysis buffer (made freshly approximately
every 2 weeks): 100 mM Tris–HCl pH 7.4
(37°C), 100 mM EDTA, 6% SDS, 2%
b-mercaptoethanol.
3. Vortex mixer.
4. Water bath.
5. Phenol: chloroform (1:1).
6. Microcentrifuge.
7. Disposable polypropylene microcentrifuge
tubes: 1.5 mL conical; 2 mL screw-capped.
8. 3 M ammonium acetate.
9. Isopropanol.
10. Tris EDTA: 10 mM Tris–HCl pH 7.5 (25°C),
0.1 mM EDTA.
11. Phenol solution equilibrated with 10 mM
Tris–HCl pH 8, 1 mM EDTA (Sigma-Aldrich,
Gillingham, UK).
12. Ethanol (70%).
13. Agarose (e.g., molecular biology grade agarose from Sigma-Aldrich, USA).
14. TBE buffer: 50 mM Tris, 50 mM boric acid,
1 mM EDTA. Dilute when needed from a
10× stock.
15. Ethidium bromide: 0.5 mg/mL stock.
16. Gel loading mixture—40% (w/v) sucrose,
0.1 M EDTA, 0.15 mg/mL bromophenol blue.
17. Horizontal electrophoresis equipment (e.g.,
Biorad Wide Mini Sub Cell).
PCR Reagents
1.
2.
3.
4.
5.
6.
7.
8.
9.
Agarose.
dNTP mix.
PCR buffer.
MgCl2.
Template DNA.
Taq DNA polymerase.
Oligonucleotide primers.
Ethidium bromide.
TBE buffer (Tris–EDTA–boric acid buffer).
PCR Assay Set-up
Prepare a master mix (for 50 mL reaction) containing the following:
1. 5 mL 10× PCR buffer.
2. mL MgCl2 (25 mM) × (number of reactions + 1).
3. mL dNTP mix (1.25 mM).
4. 30 mL sterile distilled H2O.
5. Dispense the master mix at 43 mL per PCR
tube.
6. Dispense primers in pairs at 2.5 mL per tube.
7. Dispense template at 1 mL per tube.
8. Prepare the Taq solution by diluting the appropriate amount of stock Taq DNA polymerase
to 1 unit/mL with sterile distilled H2O.
9. Dispense diluted Taq polymerase at 1 mL per
tube.
78
General PCR Conditions
After setting up the reaction, the following
conditions can be used for proper amplification
of target genes:
1. Initial denaturation: 94°C/5 min.
2. Denaturation: 94°C/1 min.
3. Annealing temperature: 52–60°C/1 to
1.30 min.
4. Extension temperature: 72°C/1 to 2 min.
5. Repeated for 30 to 35 cycles.
6. Final extension temperature: 72°C/5 to 10 min.
After successful amplification load 5 mL of
amplified PCR products to agarose gel (1%) in
containing 1% ethidium bromide and visualize
under UV.
Materials Required for
Chromatography-Based Method
1. Silica gel-coated plates (Merck chemicals).
2. Solvents, analytical grade and HPLC grade
(Sigma-Aldrich, USA).
3. Purified mycotoxin standards (SigmaAldrich, USA).
4. Micropipettes (Eppendorf 1–1,000 mL).
5. Chromatography chamber (CMAG).
6. Spray reagents.
7. UV scanner (CMAG).
8. Mechanical cup shaker (CMAG).
9. Conical flasks (250 mL).
10. Filter papers.
11. Separating funnels.
12. HPLC system with UV and fluorescent
detectors (Hitachi F-4500).
13. C18/C8 cartridges.
14. Immunoaffinity/Solid phase extraction columns for clean up.
Methods
DNA Fingerprinting Methods
The methods detailed as follows describe the general protocol for obtaining pure DNA from fungal
culture, food, and grain samples [17]. The volumes
S.C. Nayaka et al.
and number of tubes used per sample may need to
be varied, depending on the type of sample and
the quantity of mycelia being processed.
Extraction of DNA from Pure Cultures
of Fungi [18]
1. Take small pinch of mycelium into microcentrifuge tubes.
2. Add 500 mL of lysis buffer, and macerate the
mycelium with the help of a sterile glass
rod.
3. Add 50 mL of 10% SDS, vortex, and incubate
at 65°C for 10 min.
4. Add 500 mL phenol: chloroform (1:1) to each
tube and vortex briefly.
5. Centrifuge 5 min at 10,000 rpm in the
microfuge, then carefully transfer as much as
possible of the top aqueous layer to a clean
tube. Do not disturb the debris at the
interface.
6. Add 40 mL 6 M ammonium acetate, 700 mL
isopropanol, invert gently to mix, and spin
for 2 min, and incubate the mixture at −20°C
for 1–2 h.
7. Centrifuge at 10,000 rpm for 10 min and discard the supernatant.
8. Add 300 mL cold 70% ethanol, centrifuge for
2 min, and discard the supernatant.
9. Centrifuge for 10 s and remove the remaining liquid with a micropipette.
10. Allow the pellet to dry for 20 min in a fume
hood and then resuspend it in 50 mL TE or
sterile distilled water.
Extraction of DNA from Contaminated
Food Samples (e.g., Maize)
This method has been used to prepare DNA from
contaminated food grains at several laboratories.
The steps are as follows:
1. Ground the contaminated food grains.
2. Add 400 mL of lysis buffer, vortex, and incubate at 65°C for 10 min.
3. Add 500 mL phenol:chloroform (1:1) to each
tube and vortex briefly.
5
Chemical and Molecular Methods for Detection of Toxigenic Fungi…
4. Centrifuge 15 min at 12,000 rpm in the
microfuge, then carefully transfer as much as
possible of the top aqueous layer to a clean
tube. Do not disturb the debris at the interface.
5. 40 mL 6 M ammonium acetate, 600 mL isopropanol, invert gently to mix and spin for
2 min. Incubation of the mixture at −20°C
for 10–60 min before centrifugation may
improve recovery of DNA but can result in
reduced purity of the sample. Remove the
supernatant.
6. Add 50 mL RNase in TE to the pellet and
incubate at 37°C for 15 min.
7. Pool into 200 mL samples or add 150 mL TE,
and then add 200 mL phenol, vortex, and centrifuge at 10,000 rpm for 6 min.
8. Transfer carefully the top aqueous layer to a
fresh tube.
9. Add 10 mL 6 M ammonium acetate, 600 mL
isopropanol, and incubate at −20°C for 10 min.
10. Centrifuge at 12,000 rpm for 10 min and discard the supernatant.
11. Add 800 mL cold 70% ethanol, centrifuge at
10,000 rpm for 2 min, and discard the
supernatant.
12. Centrifuge at 10,000 rpm for 15 s and decant
the remaining liquid with a micropipette.
13. Allow the pellet to dry for 20 min in a fume
hood and then resuspend it in 50–200 mL TE
buffer.
Polymerase Chain Reaction
During the last few decades great advances have
been made in molecular diagnostic technology,
especially in the development of rapid and sensitive methods for the detection of plant pathogenic
fungi [19]. A number of DNA-based techniques
that have been developed include restriction fragment length polymorphism, pulse field gel electrophoresis, and PCR. PCR has been gaining
popularity mainly because of its ease of application compared to other DNA-based techniques.
There are already many examples of PCRbased assays developed for the detection of fungi
in plant pathology, but the reports on their use in
specific detection of toxigenic fungi are limited.
79
Many mycotoxin biosynthetic pathway genes are
present within gene clusters, and some of these
appear to have undergone horizontal transfer
from one species to another and are now present
in several species [7]. Regions of homology
within mycotoxin biosynthetic gene from the different species can be used to develop primers to
detect the presence of the relevant mycotoxigenic
species. This strategy was successfully applied
for aflatoxin producers [20], trichothecene-producing fungi [18], fumonisin-producing Fusarium
species [18], and also for producers of ochratoxin
[21]. PCR-based detection has been applied as an
alternative assay, replacing cumbersome and
time-consuming microbiological and chemical
methods for detection and identification of the
most serious pathogenic and mycotoxin producers in the fungal genera Fusarium, Aspergillus,
and Penicillium spp. (Table 5.1).
Polymerase Chain Reaction-Based
Detection of Aflatoxigenic Fungal Species
Target
gene
and
primers:
Nor1—
5¢CGCTACGCCGGCACTCTCGGCA3¢ (forward) and 5¢TGGCCGCCAGCTTCGACACTC3¢
(reverse). Amplicon size 400 bp.
Reaction conditions:
1. Initial denaturation: 94°C/5 min.
2. Denaturation: 94°C/1 min.
3. Annealing temperature: 58°C/1 min.
4. Extension temperature: 72°C/1 min. Repeated
for 30 cycles.
5. Final extension temperature: 72°C/5 min.
6. After successful amplification, load 5 mL of
amplified PCR products to agarose gel (1%)
containing 1% ethidium bromide and visualize under UV (Fig. 5.5).
Polymerase Chain Reaction-Based
Detection of Ochratoxigenic Fungi
Target gene and primers: pks1—5¢AGT
CTTCGCTGGGTGCTTCC3¢ (forward) and 5¢
AGCACTTTTCCCTCCATCTATCC3¢ (reverse).
Amplicon size 630 bp.
80
S.C. Nayaka et al.
Table 5.1 Primer sequences developed for metabolic pathway genes for the detection of toxigenic fungi
Target
S. No. Toxin
gene
1
Trichothecenes Tri5
2
3
4
Fumonisins
Aflatoxins
Ochratoxin
Primer sequence 5¢–3¢
GAGAACTTTCCCACCGAATAT
GATAAGGTTCAATGAGCAGAG
GATCTAAACGACTATGAATCACC
Tri6
GCCTATAGTGATCTCGCATGT
AGA GCC CTG CGA AAG(C/T) ACT GGT GC
Fum5 GTC GAG TTG TTG ACC ACT GCG
CGT ATC GTC AGC ATG ATG TAG C
Fum13 AGTCGGGGTCAAGAGCTTGT
TGCTGAGCCGACATCATAATC
CGC GCT CCC AGT CCC CTT CAT T
Aflr1
CTT GTT CCC CGA GAT GAC CA
GCC GCA GGC CGC GGA GAA AGT GGT
Ver1
GGG GAT ATA CTC CCG CGA CAC AGCC
Nor1 ACCGCTACGCCGGCACTCTCGGCAC
GTTGGCCGCCAGCTTCGACACTCCG
Omt1 GTG GAC GAA CCT AGT CCG ACA TCAC
GTC GGC GCC ACG CAC TGG GTT GGGG
AGTCTTCGCTGGGTGCTTCC
Pks1
AGCACTTTTCCCTCCATCTATCC
Amplicon
Tm (°C) size (bp) Reference
58
450
[18]
58
541
62
845
58
998
65
1,032
65
537
65
400
65
797
56
550
[22]
[20]
[21]
Fig. 5.5 Detection of aflatoxin-producing Aspergillus species targeting aflR gene (400 bp). Lane M,1-kb DNA ladder;
lane 2, negative control; lanes 3–8 aflatoxigenic Aspergillus spp
Reaction conditions:
1. Initial denaturation: 94°C/5 min.
2. Denaturation: 94°C/1 min.
3. Annealing temperature: 56°C/1 min.
4. Extension temperature: 72°C/1 min. Repeated
for 30 cycles.
5. Final extension temperature: 72°C/5 min.
6. After successful amplification, load 5 mL of
amplified PCR products to agarose gel (1%)
containing 1% ethidium bromide and visualize under UV (Fig. 5.6).
Polymerase Chain Reaction-Based
Detection of Tricothecene-Producing
Fusarium Species
Target
gene
and
primers:
tri6—
5¢GATCTAAACGACTATGAATCACC3¢ (forward) and 5¢GCCTATAGTGATCTCGCATGT3¢
(reverse). Amplicon size 446 bp.
Reaction conditions:
1. Initial denaturation: 94°C/5 min.
2. Denaturation: 94°C/1 min.
5
Chemical and Molecular Methods for Detection of Toxigenic Fungi…
81
Fig. 5.6 Detection of ochratoxin-producing fungi targeting PKS gene (630 bp). Lane 1, 1-kb DNA ladder; lane 2,
negative control; lanes 3–5, OTA-producing Aspergillus; lanes 6 and 7, OTA-positive strains of Penicillin spp
Fig. 5.7 Detection of trichothecene-producing Fusarium spp. by targeting tri6 gene (440 bp). Lane1, 1-kb DNA ladder; lanes 2–9, positive strains of Fusarium spp.; lane 10, negative control
3. Annealing temperature: 56°C/1 min.
4. Extension temperature: 72°C/1 min. Repeated
for 30–35 cycles.
5. Final extension temperature: 72°C/8 min.
6. After successful amplification, load the PCR
amplicons into ethidium bromide- containing
agarose gel and visualize bands under UV
(Fig. 5.7).
Reaction conditions:
1. Initial denaturation: 94°C/5 min.
2. Denaturation: 94°C/1 min.
3. Annealing temperature: 58°C/1 min.
4. Extension temperature: 72°C/1.30 min.
Repeated for 30 to 35 cycles.
5. Final extension temperature: 72°C/8 min.
6. After successful amplification, load 5 mL of
amplified PCR products to agarose gel (1%)
containing 1% ethidium bromide and visualize under UV (Fig. 5.8).
Polymerase Chain Reaction-Based
Detection of Fumonisins Producing
Fusarium Species
Chromatography Methods
Target
gene
and
primers:
Fum13—
5¢AGTCGGGGTCAAGAGCTTGT3¢ (forward)
and
5¢TGCTGAGCCGACATCATAATC3¢
(reverse). Amplicon size 998 bp.
Sample Extraction and Clean-up
for Mycotoxins Analysis
During the chromatographic methods the determination step is usually preceded by a number of
82
S.C. Nayaka et al.
Fig. 5.8 PCR amplification fum 13 gene (998 bp) of
toxigenic F. verticillioides and F. proliferatum. Lane M,
1-kb DNA marker; lanes 1–5, F. verticillioides standard
strains; lanes 5–10, F. proliferatum isolates; lane 11, nontoxigenic F. verticillioides isolate
operations such as sampling, sample preparation,
extraction, and clean-up. The reliability of the
results obtained by these procedures is highly
dependent on the efficiency of these steps. A
large number of components that are originally
present in the sample must be reduced, and interfering compounds that show the same behavior in
the chromatographic column must be removed as
much as possible [7].
Conventional techniques such as column chromatography and liquid–liquid extraction usually
require high amounts of solvent, are timeconsuming, tedious to apply, and expertise is
needed. Therefore, new approaches have been
investigated to simplify the extraction and cleanup procedures. A number of clean-up columns
have been developed that are used after the conventional extraction step. These procedures make
use of different principles (immunoaffinity columns, solid phase extraction, ion exchangers, and
others), but all have in common that they are
commercially available and are easy to use. They
have the additional advantages that less solvent is
required and sample preparation can be speeded
up considerably. The immunoaffinity columns
(IACs) reveal high selectivity, as only the analyte
is retained on the column and can then be eluted
easily after a rinsing step in order to remove
interfering components. Clean-up procedures are
used for the removal of interfering compounds
such as lipids, carbohydrates, and proteins [23].
IACs for clean-up purposes have become
increasingly popular in recent years because they
offer high selectivity. IACs are easy to use, and
their application for purification of samples that
are contaminated with several mycotoxins has
already been well investigated. Because mycotoxins are low weight molecules, they are only
immunogenic if they are bound to a protein carrier. If this problem is overcome, specific antibodies can be produced and bound to an agarose,
sepharose, or dextran carrier. The mycotoxin
molecules bind selectively to the antibodies
after a preconditioning step, and subsequent to a
washing step the toxin can be eluted with a solvent, causing antibody denaturation. Interfering
substances do not interact and the column is
therefore washed to remove the matrix [24].
Thin-Layer Chromatography
TLC was a very popular technique to separate
and detect mycotoxins. TLC is the most commonly utilized test because more than one mycotoxin can be detected from each test sample. TLC
5
Chemical and Molecular Methods for Detection of Toxigenic Fungi…
is based on the separation of compounds by their
migration on a specific matrix with a specific solvent. The distance that a compound will travel is
a unique identifier for specific compounds and a
retention factor (Rf) has been determined for most
mycotoxins. As with any detection system, a positive control containing purified mycotoxins must
be run in parallel to ensure accuracy. For mycotoxin assays, silica gel TLC, with both precoated
and self-coated plates, can be used. Detection
and identification procedures have been
specifically developed for each single mycotoxin,
making use of molecular properties or reactions
with spray reagents [25].
High-Performance Liquid
Chromatography
HPLC or high-pressure liquid chromatography
became available for the analysis of foodstuffs in
the early seventies and gained importance in the
determination of mycotoxins, particularly when
several types of column packings and detectors
became available. HPLC is the method of choice
because it offers the advantages of good resolution, high degree of precision, reproducibility,
and sensitivity. HPLC methods are mainly used
for the final separation of matrix components and
detection of the analyte of interest. Nowadays,
HPLC methods are widespread, because of their
superior performance and reliability compared
with TLC. HPLC methods have been developed
for almost all major mycotoxins in cereals and
other agricultural commodities. Reversed phase
(RP) chromatography is most commonly used
for the determination of mycotoxins in agricultural samples—for example, a C8 or C18 hydrocarbon phase with mixtures of polar solvents
(e.g., water: methanol or water:acetonitrile combinations). Detection is mainly performed using
diode array detection; alternatively, fluorescence
detection (FLD), which utilizes the emission of
light from molecules that have been excited to
higher energy levels by absorption of electromagnetic radiation, is employed. FLD features
superior sensitivity, although frequently derivatization of the analyte has to be performed in order
83
to make the detection possible at all or enhance
the sensitivity even further [26]. A short summary for the determination of the common toxins
of Aspergillus and Fusarium spp. by chromatographic method is presented in the following
section.
Detection of Aflatoxin by Thin-Layer
Chromatography
A number of methods have been developed for
the determination of aflatoxins by TLC. Silica
plates are most commonly in use, with a number
of solvent systems based on chloroform and small
amounts of methanol or acetone. However, a shift
can be observed to less toxic and more environmentally friendly mixtures (e.g., toluene/ethylacetate or acetone/isopropanol).
Extraction and Clean-up
Place 50 g of finely ground sample in a wide-mouth
polypropylene screw-cap bottle with 100 mL of
chloroform and water mixture (1:1) and place on
a wrist-action shaker for 30 min. Allow the contents to settle, and a 10- to 25-mL aliquot of the
solvent extract is filtered through four layers of
Whatman filter paper. Dry the organic layer by
rotary evaporator and reconstitute the compound
in 3–4 mL of methanol. Load onto a preconditioned C18 clean-up column. Wash the column
with 5 mL of phosphate buffered saline and then
elute with chloroform–methanol (97:3) mixture.
Eluants are allowed to dry for a few minutes and
re-dissolved in suitable solvents for TLC analysis, as recommended by the supplier.
Thin-Layer Chromatography
Spot the sample (0.5–2 mL) by using a capillary
tube on TLC plate. The spot should be as small
and compact as possible, with a distance of
1–2 cm from the edges of the plate and between
the spots.
1. Place TLC plate in chromatography chamber
and run until solvent front is 2–3 cm from top
of the plate (approximately 30–45 min).
2. Developing solvent: 80% benzene or toluene,
15% methanol, 5% acetic acid.
84
S.C. Nayaka et al.
Table 5.2 Rf values and visible color of aflatoxin
Rf value
Mycotoxin
Aflatoxin B1
Aflatoxin B2
Aflatoxin G1
Solvent system(B:M:A)
0.14
0.20
0.23
Color
Visible light
Yellow
Color after spray treatment
UV light
Long wave
Green
Blue
Blue
Short wave
Faint green
Faint blue
Faint blue
Long-wave ultraviolet light
Blue
Pink
Pink
Table 5.3 Rf value and visible color of Ochratoxin A
Rf value
Mycotoxin
Ochratoxin A
Solvent system (TEF)
0.55
Color
Visible light
Yellow
3. Observe the TLC plate under UV scanner at
256 nm compared with standard toxin. Toxins
were visualized in visible or ultraviolet light,
before and after the plate was sprayed with
freshly prepared mixture of 0.5 mL of p-anisaldehyde in 85 mL of methanol containing
10 mL of glacial acetic acid and 5 mL of concentrated sulfuric acid and then heated at
130°C for 10 min. The 5- to 10-min heating
time was better for fluorescence development
(Table 5.2).
Color after spray treatment
UV light
Long wave
Green
Short wave
Green
Long-wave ultraviolet light
Faint blue
5. Observation: Toxins were visualized in visible
or ultraviolet light, before and after the plate
was sprayed with a freshly prepared mixture
of 0.5 mL of p-anisaldehyde/silver chloride in
85 mL of methanol containing 10 mL of glacial acetic acid and 5 mL of concentrated sulfuric acid and then heated at 130°C for 10 min.
The 5- to 10-min heating time was better for
fluorescence development (Table 5.3).
Detection of DON by Thin-Layer
Chromatography
Detection of Ochratoxin by Thin-Layer
Chromatography
OTA detection by TLC can be performed by spotting samples and spikes onto a SG-60 plate and
development with a mixture of toluene/methanol/
acetic acid or toluene/ethyl acetate/formic acid.
Under long-wavelength UV light OTA will
appear blue–green at a retention value of 0.55.
1. Extraction and clean-up: As mentioned
above.
2. Spot the sample (0.5–2 mL) by using a capillary tube on TLC plate. The spot should be as
small and compact as possible with a distance
of 1–2 cm from the edges of the plate and
between the spots.
3. Developing solvent: 90% toluene, ethyl acetate, formic acid (5:4:1, v/v/v)
4. Observe the TLC plate under UV scanner at
256 nm compared with standard toxin.
TLC is still common, and with the introduction
of high-performance TLC (HPTLC) and scanning instruments, separation efficiency and precision have increased. Reagents (e.g., sulfuric acid
or para-anisaldehyde) are necessary to visualize
the only short wavelength absorbing DON.
Other spray reagents include 4-para-nitrobenzylpyridine or nicotinamide in combination with
2-acetyl-pyridine) or AlCl3, which is the most
useful visualization reagent for DON. Typical
detection limits by TLC are in the range of
20–300 ng/g.
1. Extraction and clean-up: 50 g of finely ground
sample were placed in a wide-mouth polypropylene screw-cap bottle with 100 mL of a
methanol–water mixture (1:1) and placed on a
wrist-action shaker for 30 min.
2. The contents were allowed to settle, and a 10- to
25-mL aliquot of the solvent extract was
5
Chemical and Molecular Methods for Detection of Toxigenic Fungi…
85
Table 5.4 Rf value and visible color of DON
Rf value
Mycotoxin
Deoxynivalenol
3.
4.
5.
6.
7.
8.
9.
Color
Solvent system (CMW) Visible light
0.45
Yellow
filtered through Whatman 4 filter paper and
extracted with 25 mL of ethyl acetate.
Ethyl acetate was completely evaporated by
rotary evaporation and the pellet resuspended
in 3 to 4 mL of acetone–water (1:1).
Extracts were passed through preconditioned
C18 column, and elutes were dried and reconstituted with 1 mL of a methanol–water mixture (1:1).
These extracts were used for TLC analysis.
Thin-layer chromatography: Spot the sample
(0.5–2 mL) by using a capillary tube on TLC
plate. The spot should be as small and compact
as possible, with a distance of 1–2 cm from
the edges of the plate and between the spots.
Place TLC plate in chromatography chamber
and run until solvent front is 2–3 cm from top
of plate (approximately 30–45 min).
Developing solvent: chloroform: methanol:
water (9:1:0.2).
Observe the TLC plate under UV scanner at
256 nm compare with standard toxin. Toxins
were visualized in ultraviolet, before and after
the plate was sprayed with a freshly prepared
mixture of 0.5 mL of p-anisaldehyde/silver
chloride in 85 mL of methanol and then heated
at 130°C for 20 min (Table 5.4).
Detection of Fumonisins by Thin-Layer
Chromatography [27]
TLC is the simplest and most frequent screening
method used for detection of fumonisins, but like
all other methods, extraction and clean-up make
a major contribution to accuracy and precision of
obtained data. Derivatization is necessary before
fluorescent detection can be performed, because
fumonisins do not contain a chromophore to
exhibit radiation. Reversed phase TLC (on C18
Color after spray treatment
UV light
Long-wave
Long wave Short wave Visible light ultraviolet light
Brown
Brown
Brown
Brick red
modified silica plates) has also been employed
with acidic vanillin or fluorescamine/sodium
borate buffer as a spray reagent.
1. Extraction and clean-up: 50 g of finely ground
sample were placed in a wide-mouth polypropylene screw-cap bottle with 100 mL
acetonitrile: water (1:1) and placed on a wristaction shaker for 30 min.
2. The contents were allowed to settle, and a 10to 25-mL aliquot of the solvent extract was
decanted and filtered using Whatman 4 paper.
3. A C18 clean-up column was preconditioned
with 5 mL of methanol followed by 5 mL of 1%
aqueous potassium chloride (KCl). Two milliliters of the filtrate was combined with 5 mL
1% aqueous KCl and applied to the column.
4. The column was washed with 5 mL 1% aqueous KCl followed by 2 mL acetonitrile: 1%
aqueous KCl (1:9), and the eluants were
discarded.
5. The fumonisins were eluted with 4 mL acetonitrile: water (7:3), and the column eluant
was evaporated to dryness under a stream of
air on a heating module for TLC analysis.
6. Thin-layer chromatography: The sample residue was dissolved in 100 mL acetonitrile:
water (1:1).
7. 10 mL was spotted on a C18 TLC plate along
with 10-mL fumonisins standards (5, 10, and
100 ppm) dissolved in acetonitrile: water (1:1).
8. Observation: The TLC plate was developed in
methanol: 1% aqueous KCl (3:2), air dried,
and sprayed with 0.1 M sodium borate buffer
(pH 8–9) followed by fluorescamine (0.4 mg/
mL in acetonitrile). After 1 min, the plate was
sprayed with 0.01 M boric acid: acetonitrile
(40:60). The plate was then air dried at room
temperature and examined under long-wave
UV light. Fumonisin levels were estimated by
visual comparison with standards (Table 5.5).
86
S.C. Nayaka et al.
Table 5.5 Rf value and visible color of fumonisins
Mycotoxin
Fumonisins
Rf value
Solvent system
(M:Kcl)
0.5 (FB1)
0.1 (FB2)
Color after spray
treatment
Long-wave ultraviolet
light
Bright yellowish-green
fluorescent bands
Detection of Aflatoxins
by High-Performance Liquid
Chromatography
1. Instrument: Liquid chromatography methods
for the determination of aflatoxins in foods
by reversed-phase HPLC (Hitachi F-4500).
The emitted light is detected at 435 nm after
excitation at 365 nm. Stationary phases for
HPLC usually include C18 material, with mobile
phases being mixtures of water, methanol, or
acetonitrile. A fluorescence detector and a suitable data system are required to provide sensitive and specific detection and quantification
of aflatoxins.
2. Solvents: All solvents shall be of HPLC grade,
and all reagents should be analytical grade.
3. Extraction: A ground sample (20 g) is extracted
with a methanol–water (7:3) mixture (80 mL).
Corn and wheat samples will be kept in a
vibrating shaker for 15–30 min. Extracts are
filtered immediately after extraction through
filter paper. After filtration the sample is evaporated to dryness at 40°C in a rotary
evaporator.
4. Clean-up by IAC: The use of IACs is now well
established in aflatoxin determination.
MycoSep® (Romer Labs, Union, MO) columns, which remove matrix components
efficiently and can produce a purified extract
within a very short time, are also available.
Conventional clean-up with silica columns
has also been reported [16].
5. Standard preparation: Aflatoxin B1, B2, G1,
and G2 standard can be purchased from private companies (Sigma-Aldrich, USA). Each
of aflatoxins was diluted in methanol to 1 mg/
mL solution of G2, 1 mg/mL of B2 10 mg/mL
of G1, and 10 mg/mL of B1. 100 mL aliquot of
each aflatoxin solution was then combined in
a 2-mL glass vial and mixed well. This mixture
was further diluted in series to 100,000 folds
in water: methanol (7:3 v/v) and used as the
standard solution.
6. Chromatography conditions: Column: Hypersil
GOLD®, 3 mm, 100 × 2.1 mm; Flow Rate:
800 mL/min lex: 365 nm lem: 455 nm; Mobile
Phase: Water: Methanol (7:3 v/v) (isocratic
elution); Column Temperature: 40°C; Injection
Volume: 10 mL of the prepared standard solution; Analytes: aflatoxin B1 and aflatoxin B2.
The instruments will be controlled and the data
analyzed using the suitable data system. No
step changes of the excitation and emission
wavelengths will be used during the run.
7. Observation: Aflatoxins fluoresce strongly on
illumination with 365-nm ultraviolet light.
Figure shows the fluorescence chromatogram
of the two common aflatoxins with an excitation wavelength of 365 nm and an emission
wavelength of 435 nm.
Detection of Ochratoxins by HighPerformance Liquid Chromatography
1. Instruments: The liquid chromatograph
equipped with quaternary pump, autoinjector
with a stainless steel reverse phase
150 × 4.6 mm, 3-mm particle size C18 Supelco
HPLC column (Supelco, USA). A fluorescence
detector and a suitable data system are required
to provide sensitive and specific detection and
quantification of ochratoxins derivatized with
OPA/mercaptoethanol.
2. Solvents: All solvents shall be of HPLC grade,
and all reagents should be analytical grade.
3. Extraction: Sample extraction is generally
performed with a mixture of water and organic
solvents depending on the type of matrix. An
IUPAC/AOAC method for the determination
of OTA in barley uses a mixture of CHCl3 and
H3PO4 [28]; for green coffee, CHCl3 is only
employed [29]. For determination in wheat, a
number of extraction solvents are used, including mixtures of toluene/HCl/MgCl2, CHCl3/
ethanol/acetic acid, and dichloromethane/
H3PO4.
5
Chemical and Molecular Methods for Detection of Toxigenic Fungi…
4. Clean-up by IAC: The use of IACs is now
well established in ochratoxin determination.
The extract is forced through the column, and
ochratoxins are bound to the antibody. Five
milliliters of the final extract, corresponding
to 5% (v/v) of the original material, was placed
into the IAC. The sample was allowed to pass
though the column at a flow rate of 2–3 mL/
min. Slowly elute the bound ochratoxin from
the column using 1.5 mL of desorption solution; allow this to pass through the column by
gravity and collect in a sample vial.
5. Standard preparation: Ochratoxin will be purchased from private companies (SigmaAldrich, USA). Ochratoxin was diluted 50 mg/
mL in benzene: acetic acid (99:1). 50 mL aliquot of each solution was then combined in a
2-mL glass vial and mixed well. This mixture
was further diluted in series to 100,000 folds
in acetonitrile: water (7:3 v/v) used as the
standard solution.
6. Chromatography conditions: Reversed phase
HPLC approach with a C18 column [21].
Flow Rate: 800 mL/min lex: 365 nm lem:
455 nm; and an acidic buffer (acetic acid) in
an acetonitrile/water mixture as a mobile
phase. Column Temperature: 40°C; Injection
Volume: 10 mL of the prepared standard
solution; Analytes: Ochratoxin A.
7. The instruments will be controlled and the
data analyzed using the suitable data system.
No step changes of the excitation and emission wavelengths will be used during the run.
8. Observation: Quantify the ochratoxin A concentration by comparing the sample peak area
to that of a standard.
Detection of Deoxynivalenol by HighPerformance Liquid Chromatography
1. Instruments: The liquid chromatograph
equipped with quaternary pump, autoinjector,
and UV detector was used with a stainless
steel reverse phase 150 × 4.6 mm, 3 mm particle size C18 Supelco HPLC column.
2. Solvents: All solvents shall be of HPLC grade,
and all reagents should be analytical grade.
87
3. Extraction: Place 10 g of the ground sample
into the ultraturax and then add 40 mL of distilled water and 2 g of polyethylene glycol.
The mixture is stirred for 1 min. The extract is
filtered through a fluted filter and then through
a microfiber filter.
4. Clean-up by immunoaffinity chromatography:
Place 1 mL of the final extract into the IAC.
Use 10 mL of redistilled water for column
washing. The elution of DON is conducted
with 1 mL of methanol. The elution solvent is
removed by a gentle stream of nitrogen and
re-dissolved in 300 mL mobile phase.
5. Standard preparation: DON purchased from private companies (Sigma-Aldrich) is diluted to
200 mg/mL in ethyl acetate: methanol (95:5).
50 mL. Aliquot of the solution is then combined
in a 2 mL glass vial and mixed well. Serially
dilute this mixture to 1,000 folds in methanol:
water (7:3 v/v) used as the standard for HPLC.
6. Chromatography condition: Samples of 50 mL
are injected into the HPLC column and heated
to 30°C. The used mobile phase consisted of a
methanol: water solution (8:2 v/v). The flow
rate is of 0.6 mL/min. Deoxynivalenol is
determined at a wavelength of 218 nm by
using UV detector.
7. Observation: Quantify the deoxynivalenol
concentration by comparing the sample peak
area to that of a standard.
Detection of Fumonisins by HighPerformance Liquid Chromatography
1. Instrument: HPLC system consisting of an
isocratic pump capable of a flow rate of 1 mL/
min and a suitable injector capable of 10 mL
injections. Columns containing C18- or C8modified silica packing material of 3- to 5-mm
particle size. A fluorescence detector and a
suitable data system are required to provide
sensitive and specific detection and
quantification of fumonisins derivatized with
OPA/mercaptoethanol.
2. Solvents: All solvents will be used of HPLC
grade and all regents should be analytical
grade.
88
3. Extraction: Place finely ground sample (25 g)
into a container suitable for centrifuging (250mL polypropylene centrifuge bottle). Add
100 mL extraction solvent (methanol–water,
3:1) and homogenize the contents for 3 min.
Centrifuge the container at 10,000 rpm for
10 min at 4°C. Filter the supernatant through a
Whatman 4 filter paper.
4. Solid phase extraction (SPE) cartridges:
Sample extracts are generally cleaned up on
SPE columns containing strong anion
exchange material. For optimal simultaneous
handling of cartridges, the use of a commercial SPE manifold is recommended.
5. Standard preparation: Fumonisin standards
are prepared in acetonitrile: water (1:1) and
stored at 4°C. Stock solutions of individual
fumonisins standards of concentration 250 mg/
mL are used, from which a working standard
is prepared containing 50 mg/mL of each analog. Derivatize standards by mixing 25 mL
working standard with 225 mL OPA reagent at
the base of a small test tube. Inject 10 mL into
the HPLC using a standardized time of
1–2 min between the addition of OPA reagent
and injection.
6. Chromatography conditions: The HPLC
mobile phase is a mixture of methanol and
0.1 M sodium dihydrogen phosphate in water.
For most reversed-phase columns, a solvent
composition of 75% to 80% methanol will be
required. The pH of the mixture is adjusted to
3.35 with o-phosphoric acid and filtered
through a 0.45-mm membrane filter.
7. OPA reagent: OPA reagent for derivatizing the
fumonisins is prepared by dissolving 40 mg
OPA in 1 mL of methanol and diluting with
5 mL of 0.1 M disodium tetraborate.
8. Observation: Quantify the fumonisins’ concentration by comparing the sample peak area
to that of a standard.
Summary
Many agricultural commodities are vulnerable to
attack by fungi that produce mycotoxins.
Detection of mycotoxins and toxin-producing
S.C. Nayaka et al.
fungi from food and feeds are very essential. In
the present chapter we discussed available techniques for detection and quantification of major
mycotoxigenic fungi and their toxins from agricultural produce. The standard methods varied
from lab to lab and toxin to toxin and also from
commodity to commodity. International agencies
such as International Union of Pure and Applied
Chemistry (IUPAC), Association of Official
Analytical Chemists (AOAC), and The European
Mycotoxin Awareness Network have developed
their own methodologies for detection of mycotoxins from different food matrices. In conclusion, a broad range of techniques for practical
analysis and detection of a wide spectrum of
mycotoxins are available. This chapter presented
some recent developments in scientific and technological basis analytical methods that offer
flexible and broad-based methods for analysis of
toxins and toxigenic fungi.
Notes
DNA Extraction and Polymerase Chain
Reaction Conditions
1. Make use of suitable microbiological aseptic
technique when working with DNA. Wear
gloves to prevent nuclease contamination
from the surface of the skin. Use sterile, disposable plasticware and automatic, aerosolresistant pipettes reserved for DNA work.
2. Wipe pipettes with Dnase-removal solutions
when transitioning between handling crude
extracts to handling more purified material.
3. Equilibrated phenol can typically be purchased
from
commercial
sources.
Alternatively, you can equilibrate it yourself.
There are also commercial sources of phenol
and chloroform mixed together and equilibrated. The pH is important because chromosomal DNA will end up in the phenol phase
if the pH is acid (around pH 5).
4. Phenol and chloroform should be used in a
hood. Phenol is a dangerous substance that
will burn you if it gets on your skin. Always
wear gloves and be careful. A solution of
5
Chemical and Molecular Methods for Detection of Toxigenic Fungi…
PEG 400 is recommended for first aid. Phenol
is both a systemic and local toxic agent.
5. DNA should be kept frozen in a non-frostfree freezer. DNA should not be allowed to
defrost between uses, as this will break long
molecules.
6. Make a PCR master mix for 50 mL reaction
containing DNA and PCR ingredients. After
setting up the reaction, specific reaction conditions can be used for proper amplification
of target genes. After successful amplification,
the PCR amplicons can be stained with
ethidium bromide-containing agarose gel
and bands can be visualized under UV.
Mycotoxin Analysis
7. A laboratory or part of a laboratory should be
reserved for mycotoxin analysis only and the
work confined to that area. The bench top
should be of a nonabsorbent material, such
as formica, for example (Whatman Benchkote
can also be used, but it must be removed and
destroyed after use), and should be screened
from direct sunlight.
8. Analyses should be performed in a well-ventilated laboratory, preferably under an
efficient extraction hood, and fume cupboard
facilities should be available.
9. Many of the solvents used are highly
flammable and have low flash points. Bunsen
burners, electric fires, and sparking apparatus such as centrifuges should not be used in
the same laboratory. The amount of
flammable solvents in the laboratory should
be kept to a minimum and stored in a fireresistant cupboard or bin.
10. Swab accidental spills of toxin with 1%
NaOCl bleach, leave 10 min, and then add
5% aqueous acetone. Rinse all glassware
exposed to aflatoxins with methanol, add 1%
NaOCl solution, and after 2 h add acetone to
5% of the total volume. Allow a 30-min reaction and wash thoroughly.
11. Weighing and transferring mycotoxins in dry
form should be avoided; they should be dissolved in a solvent. The electrostatic nature
89
of a number of the mycotoxins in dry form
results in a tendency for them to be easily
dispersed in the working area and to be
attracted to exposed skin and clothes.
12. Containers of mycotoxin standard solutions
should be tightly capped, and their weights
may be recorded for future reference before
wrapping them in foil and storing them in a
freezer.
13. During the grinding and weighing of samples,
there is a risk of absorbing toxin either through
the skin or by the inhalation of dust. There is
also the risk of developing allergic reactions
due to spores and organic material. These
risks should be minimized by working under
an extraction hood, by good hygiene, and by
wearing protective clothing and masks.
14. Glassware and TLC plates should be decontaminated by soaking for 2 h in a 1% sodium
hypochlorite solution. After this time an
amount of acetone equal to 5% of the total
volume of the bleach bath should be added,
and the glassware soaked for an additional
30 min. Spraying of TLC plates must be carried out in an efficient fume cupboard or
spray cabinet. Always ensure that this equipment is working before commencing use.
When viewing chromatograms under UV
light the eyes should be protected by UV
filter or by wearing protective spectacles.
References
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of foods and feeds: overview, occurrence and economic impact on food availability, trade, exposure of
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SR, Ednar GW, Mortensen CN, Prakash HS (2010)
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SR, Mortensen NM, Prakash HS (2011) Prospects of
Molecular markers for Fusarium diversity. Appl
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of ochratoxin A producing fungi. In: Rai M (ed)
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Delhi, India, p 320
12. http://www.mycotoxins.org/. Accessed 20 Sep 2011
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human systems. Task Force Report No. 139. Ames,
IA: Council for Agricultural Science and Technology,
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18. Ramana MV, Balakrishna K, Murali HS, Batra HV
(2011) Multiplex PCR-based strategy to detect contamination with mycotoxigenic Fusarium species in
rice and fingermillet collected from southern India. J Sci
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(2003) Molecular diagnostics for fungal plant pathogens. Pest Manag Sci 59(2):129–142
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Salomon R (1996) Detection of aflatoxigenic molds
in grains by PCR. Appl Environ Microbiol 62:
3270–3273
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(2002) Multiplex polymerase chain reaction assay for
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6
Identification Key for the Major
Growth Forms of Lichenized Fungi
Jeyabalan Sangeetha and Devarajan Thangadurai
Abstract
Lichens are classified as cup fungi, mainly under the phylum of Ascomycota
and rarely as Basidiomycota. The body of the lichen is called a thallus, in
which the mycobionts and photobionts are stratified in separate layers.
Based on their characteristics and thalli, lichens are classified into four
main groups. This chapter consists of a pair of parallel and opposing statements that can be compared and will help to identify several genera of
foliose, fruticose, crustose, and squamulose lichens.
Keywords
Lichenized fungi • Foliose • Fruticose • Crustose • Squamulose •
Identification keys
Introduction
Kingdom Fungi comprises seven major phyla,
such as Ascomycota, Basidiomycota, Blastocladiomycota, Chytridiomycota, Glomeromycota,
Microsporidia, and Neocallimastigomycota
[1–3]. The total number of fungi is estimated at
700,000 to 1.5 million species. Members of
Ascomycota are commonly known as sac fungi.
J. Sangeetha (*)
Department of Zoology, Karnataka University,
580003, Dharwad, Karnataka, India
e-mail: drsangeethajayabalan@gmail.com
D. Thangadurai
Department of Botany, Karnataka University,
Dharwad, 580003 Karnataka, India
They are the largest phylum of fungi, containing
about 32,000 named species. The fungal symbionts in the majority of lichens belong to the
Ascomycota phylum, and a few belong to
Basidiomycota. Among the identified fungal species, about 17,500 lichens are recorded [4–7].
The number of undescribed species of lichenized
fungi has been estimated at roughly 10,000.
These fungi form meiotic spores called
ascospores, which are enclosed in a special saclike structure called an ascus. However, some
members of Ascomycota do not reproduce sexually and do not form asci or ascospores. These
members are assigned to Ascomycota based upon
morphological or physiological similarities to
ascus-bearing taxa and, in particular, by phylogenetic comparisons of DNA sequences [8–11]. In
addition to lichens, this phylum includes morels
(e.g., Morchella deliciosa, M. elata), a few
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_6, © Springer Science+Business Media, LLC 2013
91
92
mushrooms and truffles, single-celled yeasts, and
many filamentous fungi living as saprotrophs,
parasites, and mutualistic symbionts [12–15].
Structurally, lichens are the strangest of all
forms of fungi and are dual organisms formed by
a symbiotic relationship of a fungus (mycobiont)
with an algae or cyanobacterium (photobiont)
[16,17]. Some factors, like substratum chemistry,
stability, and longevity, and light and moisture
availability, are affecting the presence and abundance of lichens. Some substrata that support
lichen growth include rock surface, woody plant
bark, wood, soil, microhabitats, and broad evergreen leaves in the humid tropics [3,18]. Lichens
appear in a variety of colors, like white, grey,
brown, orange, brilliant red, or yellow [19–21].
Lichen morphology is quite different from that of
other fungi [22,23]. Each lichen thallus has a
complete microscopic feature with unique characteristics [17]. From an ecological perspective,
lichen may be even more complex than free-living
bacteria and nonsymbiotic fungi. Longevity and
considerable environmental stress tolerance seem
to be the major features shared by all ascomycetes
[24]. The basic lichen body is called a thallus, and
it consists of fungal hyphae that are organized
into various tissues and thread-like fungal cells.
Lichens consist of four layers: the top surface,
which is a layer of tightly packed thallus called
upper cortex; an algal layer, where the photobionts lives; the medulla, which is bound with loose
hyphae; and another protective covering, the
lower cortex [25,26]. In lichen, the algae and the
fungi can reproduce individually but to form recognizable lichen, viable fungal spores have to
unite with the appropriate algal spores to form
new lichen. The surface of the lichen has three
important structures: fruiting body, vegetative
powdery, and hard-bodied peg-like stalk [27,28].
The detection and identification of lichens are
based on the conventional method of observation
of physical characteristics of spores, hyphae,
arrangement thallus, life cycle, and relationship
with the host cell. An identification key can be
prepared for each genus by using these observations. Normally, an identification key is a printed
or computer-aided document that provides the
details of the lichen and helps in the identification.
J. Sangeetha and D. Thangadurai
An identification key is a single-access key, and it
will work with fixed sequence of identification
steps, each with multiple alternatives, the choice
of which determines the next step. Some lichens
can be identified with a few notes, and others
may require more notes for identification. This
key will assist the investigator in identifying an
unknown specimen to its genus. The amount of
information presented in the key limits the effectiveness of the key. Hence, the key should be as
comprehensive as possible; inclusion of too much
information in a single point makes the key
difficult to follow. The keys are organized in
groups of options in the middle column, and these
are numbered in ascending order in the left-hand
column (1a, 1b, 2a, 2b, etc.). Having first decided
which key to use, start at the top and choose the
most appropriate option to describe the specimen
in question (e.g., 1a or 1b). Description of the
option will lead to either a possible solution in
the form of a name or the number of a further
option, which will be in the right-hand column.
Moving down with the key to the number indicated earlier once again offers either a possible
solution or a further pair of options to choose and
so on until the search ends with no further options
offered [26,29]. The identification obtained from
a key should be viewed as only a suggestion of
the specimen’s real identity. Full identification
requires comparison of the specimen with some
authoritative source and further well-developed
molecular techniques [30].
This chapter has two primary objectives. The
first is to provide protocol for the collection and
isolation of lichenized fungi. The second is to
provide comprehensive identification keys to the
main growth forms of lichens.
Materials
1.
2.
3.
4.
5.
6.
7.
Autoclave
Incubator
Laminar air flow chamber
Centrifuge
Dissecting microscope
Compound microscope
Inverted microscope
6
Identification Key for the Major Growth Forms of Lichenized Fungi
8.
9.
10.
11.
12.
Microscopic slide
Distilled water
Petroleum jelly
Ethanol
4 % distilled water agar medium: agar, 4 g;
distilled water, 100 mL
13. Malt/yeast extract medium (MY medium):
malt extract, 20 g; yeast extract, 2 g; agar,
20 g; distilled water, 1,000 mL
14. Lilly and Barnett’s medium (LB medium):
glucose, 10 g; asparagine, 2.0 g; KH2PO4,
1.0 g; MgSO4⋅7H2O, 0.5 g; Fe(NO3)3⋅9H2O,
0.2 mg; ZnSO4⋅7H2O, 0.2 mg; MnSO4⋅4H2O,
0.1 mg; thiamine, 0.1 mg; biotin, 5 mg; distilled water, 1,000 mL
15. Lactophenol cotton blue
Methods
Lichens can be isolated from ascospores, conidia,
isidia, soredia, and thallus fragments. Lichens
show maximum growth at 15–20 °C under the
pH of 5–6. Lichens can be cryopreserved so that
they remain viable for extended periods of time
[16,17,31]. Spot tests (see Note 1) can be done
with sodium hypochlorite (C), potassium hydroxide (K), potassium iodide (I), or paraphenylenediamine (P) to identify the color reaction of lichen
substances (see Note 2). For laboratory cultures
the most useful methods are isolating lichens
from discharged spores and thallus fragments.
When mycobionts cannot be isolated from the
spores, other parts like conidia, isidia, and soredia may be applicable for isolation [10,26,32].
Spore Discharge Method
1. Collect thalli from the field and clean it (see
Note 3).
2. Remove the apothecia or perithecia from the
thallus.
3. Place it into the dishes containing distilled
water and wash it.
4. Blot dry to remove excess water.
5. Fix these structures to the bottom of a Petri
dish with petroleum jelly.
93
6. Pour 4 % water agar medium on the lid of the
Petri dish and allow it to solidify (see Note 4).
7. Replace the top cover of the Petri dish with
new covers containing freshwater agar
medium.
8. Observe spores discharged onto the water
agar medium under inverted microscope.
9. Keep discharged spores on a sterile glass
slide containing medium.
10. Keep the slides in a Petri dish in humid
conditions.
11. Observe continuously for the germination of
spores and mycelia growth, using lactophenol cotton blue stain under microscope (see
Note 5).
12. After germination, excise small blocks of the
agar containing spores and transfer to the
culture tubes or Petri dishes containing MY
medium and LB medium.
13. Note all measurements and match with the
lichen identification key for the identification
of the isolated lichen.
Isolation of Mycobionts from Thallus
1. Using sterilized cutter or blade, cut pieces
from a fresh thallus.
2. Store in small test tubes containing water or
on wet filter paper at 15 °C. After 2 weeks a
new medullary hyphae will be elongated.
3. Excise a portion of the newly elongated
hyphae and transfer to a test tube containing
fresh culture medium (MY medium or LB
medium).
4. Prepare sufficient number of replicates for
confirmation.
5. Identify the isolated lichen with the list of
lichen identification keys.
Key for the Identification of Lichenized
Fungi
Based on the growth forms, lichens are grouped
into four main categories: foliose, fruticose, crustose, and squamulose. Basic growth forms used
for identification of lichen specimens are thallus
94
adenation or attachment to the substrate, lobe
width, color, soredia, isidia, cilia, ahizines, tomentum, veins, and pores. Pairs of keys are constructed
based on the contrast characters. Each key is followed until one reaches a match for the description
of the specimen. Keys to the major growth forms
are as follows [20,25,26,28,33–36]:
1. Foliose Lichens. Thallus foliose, leaf-like with
branching lobes, adnate to suberect or umbilicate with a central umbilicus below; thalli are
flattened, having upper and lower cortex; the
lobes can be narrow or broad; rhizines present.
2. Fruticose Lichens. Thallus fruticose or
shrubby, richly divided, rounded or flattened
in cross section, decumbent or tufted to pendulous, and attached at the base or free growing; thallus filamentous forming small tufts or
mats; no lower cortex and rhizines.
3. Crustose Lichens. Thallus crustose, closely
attached to the substrate and lacking a lower
cortex and rhines but sometimes with a lobed
margin; or thallus consisting of small crowded
squamulose.
4. Squamulose Lichens. Thallus horizontally
spreading, blister-like, squamulose; lacking
lower cortex and rhizines; the lower layer is
white; some lichens produce fruiting structure
called podetium, an erect, hollow stalk.
J. Sangeetha and D. Thangadurai
substratum at one point, and the remaining major
portion is either growing erect or hanging. Thalli
may be pendulous strands or hollow, upright
stalks. Presence of soredia and isidia is also
important for identification. The key for the
identification of fruticose lichens is listed in
Table 6.2 [16,25,35,38–40,43,45,46].
Identification Key
to the Crustose Lichens
The thallus in crustose lichen is closely attached
to the substrate and may not be removed from it
without destruction of the substrate. The thallus
usually lacks lower cortex and rhizines. About
75 % of the lichens are crustose and found in icefree areas of the highest mountains. Compared to
other lichens, crustose lichens can grow faster on
culture medium. Crustose lichens can tolerate
extreme conditions, such as exposure to a rock
surface, and also demonstrate a clearly defined
growth type. Crustose lichens have two types of
fruiting body: open disc-shaped apothecia and
flask-shaped perithecia. These characters play
important roles in identification of crustose. The
identification key for the crustose lichens is given
in Table 6.3 [22,25,38,39,43,44,46–53].
Identification Key to the Foliose Lichens
Certain lichens are leaf-like and composed of lobes
usually strap-shaped and live on leaves, sometimes
as parasites. These special leaf-living lichens are
known as foliose lichens. Foliose are loosely
attached to the substrate and have an upper and
lower cortex. Foliose lichens grow slowly on culture medium. The identification keys for the foliose
lichens are listed in Table 6.1 [16,22,35,37–45].
Identification Key
to the Fruticose Lichens
Fruticose lichens are mostly three-dimensional.
Most are branched, the thallus is attached to the
Identification Key
to the Squamulose Lichens
Sometimes, crustose lichens develop blister-like
“squamules” where the areolae are enlarged in
their upper part and become partially free from
the substrate. Such lichens are known as squamulose lichens. Here the lichen thallus is in the form
of minute lobes. Their form is similar to that of
crustose lichens in that they possess an upper cortex but no lower cortex. The squamulose are often
developed in rock surfaces in hot and arid regions
of the world. A key for the identification of
squamulose lichens is listed in Table 6.4
[23,28,35,38,39,43,46,54–59].
6
Identification Key for the Major Growth Forms of Lichenized Fungi
95
Table 6.1 Identification keys for foliose lichens
1a
b
2a
b
3a
b
4a
b
5a
b
6a
b
7a
b
8a
b
9a
b
10a
b
11a
b
12a
b
13a
b
14a
b
15a
b
16a
b
17a
b
18a
b
19a
b
20a
b
21a
b
22a
b
23a
b
Thallus squamulose; photobiont cyanobacterium .............................................................................................2
Thallus gelatinous, homomerous, without arachnoid; photobiont Nostoc.........................................................3
Photobiont cells arranged in chains; cortex more than one layer thick; hypothallus present ..........................11
Photobiont cells arranged in small groups; lacking a distinct hypothallus;
brown and smooth upper cortex.........................................................................................................................5
Thallus with pseudoparenchymatic cortex of one cell layer ..............................................................................4
Thallus with perithecia.......................................................................................................................................5
Thallus > 0.5 mm thick; spores simple .................................................................................................... Physma
No thallospores and anthraquinones ..................................................................................................................7
Thallus attached by umbilicus ................................................................................................................. Thyrea
Thallus isidiate or papillate ................................................................................................................................6
Thallus attached by umbilicus ...........................................................................................................................7
Thallus blackish, attached on rock .....................................................................................................................9
Thallus surface grey to brown or dark brown ....................................................................................................8
Thallus green; apothecia lecanorine...................................................................................................Rhizoplaca
Perithecia embedded in thallus, immersed in the thallus, producing simple,
hyaline ascospores ..................................................................................................................... Dermatocarpon
Apothecia lecideoid; ascocarps absent; thallus dark color .............................................................. Umbilicaria
Thallus surface yellow or orange .....................................................................................................................10
Thallus surface dark brown or greenish grey or yellowish grey ......................................................................15
Thallus surface yellow; K- negative; ascospores simple; apothecia absent ....................................................11
Thallus surface orange; K + dark violet; ascospores polarilocular ...................................................................13
Thallus loosely appressed, lobes rounded with concentric ridges, not gelatinous, photobiont
bluegreen algae; apothecia often with thalline margin ....................................................................................59
Thallus lobes elongate, without concentric ridges; photobiont green algae ....................................................12
Thallus soridiate, greyish yellow or bright yellow to greenish; K-; pigment mainly in cortex;
apothecia sessile; narrow lobes hardly > 1 mm wide ........................................................................ Candelaria
Thallus lobes > 1 mm wide ...............................................................................................................................50
Ascospores bilocular ........................................................................................................................................14
Ascospores tetralocular .................................................................................................................... Teloschistes
Upper and lower cortex prosoplectenchymatic; with elongate lumina not wider than walls .......... Josefpoeltia
Upper cortex paraplectenchymatic; lower cortex prosoplectenchymatic .................................. Xanthomendoza
Lower side smooth; with or without rhizines ..................................................................................................16
Lower side tomentose; with or without cortex, with or without rhizines ........................................................53
Upper surface of lobes with white; rounded or elongate pseudocyphellae .....................................................17
Upper surface of lobes without pseudocyphellae ............................................................................................20
Thallus yellowish to greenish grey; with usnic acid in the cortex .............................................. Flavopunctelia
Thallus whitish grey; with atranorin in the cortex ...........................................................................................18
Pseudocyphellae with reticulate pattern...........................................................................................................43
Pseudocyphellae rounded; without reticulate pattern ......................................................................................19
Lobes ascending; > 1 cm wide; lower side black ................................................................................... Cetrelia
Lobes appressed to substrate; < 1 cm wide; lower side white ............................................................. Punctelia
Cilia present on the lobe margins.....................................................................................................................21
Cilia absent from the margins ..........................................................................................................................34
Cilia with inflated base.....................................................................................................................................22
Cilia without inflated base ...............................................................................................................................23
Upper side of thallus yellowish green; usnic acid present ..................................................................... Relicina
Upper side of thallus whitish grey; atranorin present .........................................................................Bulbothrix
Marginal rhizines or cilia white or grey, with perpendicular side branchlets; ascospores one-septate .......... 45
Marginal rhizines or cilia black, unbranched; ascospores simple and colorless..............................................24
(continued)
96
J. Sangeetha and D. Thangadurai
Table 6.1 (continued)
24a
b
25a
b
26a
b
27a
b
28a
b
29a
b
30a
b
31a
b
32a
b
33a
b
34a
b
35a
b
36a
b
37a
b
38a
b
39a
b
40a
b
41a
b
42a
b
43a
b
44a
b
Rhizines one or more times dichotomously branched .....................................................................................43
Rhizines unbranched ....................................................................................................................................... 25
Thallus lobes linear, with parallel margins, dichotomously branched .............................................................26
Thallus lobes elongate or short and wide; irregularly branched ......................................................................28
Lower side brown, without rhizines; terricolous lichen with upright lobes ....................................................40
Lower side black; with rhizines; predominantly epiphytic with spreading lobes ............................................27
Lobes regularly dichotomously branched; apothecia concave when young .................................. Everniastrum
Lobes irregularly branched; apothecia flat to convex ................................................................... Cetrariastrum
Thallus < 2 mm wide; flattened ........................................................................................................................29
Thallus > 5 mm wide; ascending ......................................................................................................................31
Cilia robust, tapering; upper surface white .................................................................................. Canomaculina
Cilia slender, not tapering; upper surface grey ................................................................................................30
Medulla yellow to orange ................................................................................................................ Myelochroa
Medulla white .............................................................................................................................. Parmelinopsis
Underside near lobe tips without rhizines........................................................................................................41
Underside rhizinate to the margin ....................................................................................................................32
Lobe tips appressed to substratum; rhizines with uniform length;
shorter towards the lobe tips ..................................................................................................... Parmotremopsis
Lobe tips ascending; rhizines with variable length; dimorphic .......................................................................33
Underside near margin brown; with scattered rhizines of variable length ............................................. Rimelia
Underside near margin pale; with dense; short rhizines mixed with scattered longer ones ............... Rimeliella
Rhizines absent ................................................................................................................................................35
Rhizines present .............................................................................................................................................. 40
Thallus lobes hollow; with cavity between medulla and black lower cortex ..................................................36
Thallus lobes compact......................................................................................................................................37
Lobes with large pore; thallus with perforations on upper side; soralia laminal;
atranorin, sticitic and constictic acids ...............................................................................................Menegazzia
Lobes without large pores; thallus without perforations on upper side;
soralia terminal or subterminal ....................................................................................................... Hypogymnia
Thallus lobes erect; attached near to the base; lower side near the tip often white; epiphytic
Thallus attached to the substrate; lower side black; epilithic ..........................................................................38
Upper side yellowish or greenish grey, with usnic acid; ascospores hyaline, simple ................... Psiloparmelia
Upper side pale to dark grey, without usnic acid; ascospores grey, 1-3-septate; widespread......................... 39
Thallus K + yellow; cortex containing atranorin; apothecia with dark hypothecium;
thallus lobes elongate and laterally confluent ..................................................................................................46
Thallus K-; cortex lacking atranorin; apothecia with pale hypothecium;
thallus lobes not laterally confluent .............................................................................................. Hyperphyscia
Thallus pale yellowish on both sides; lower side with pseudocyphellae; rhizines scarce ..................... Cetraria
Thallus not pale yellowish on both sides; lower side without pseudocyphellae............................................. 41
Rhizines absent in marginal zone, scarce and restricted to small patches;
thallus lobes wide, > 10 mm wide ...................................................................................................Parmotrema
Rhizines present up to lobe margins, regularly spread all over the surface;
thallus lobes usually > 2 mm wide, rather narrow and deeply dissected ..........................................................42
Rhizines frequently dichotomously branched............................................................................... Hypotrachyna
Rhizines mostly unbranched ........................................................................................................................... 43
Apothecia completely black, without grey thalline margin ...................................................................... Pyxine
Apothecia with grey thalline margin................................................................................................................44
Ascospores one-septate, grey to brown; thallus closely appressed to the substrate;
lower side whitish or black ..............................................................................................................................45
Ascospores simple, hyaline; thallus mostly loosely appressed to the substrate;
lower side brown to black ................................................................................................................................49
(continued)
6
Identification Key for the Major Growth Forms of Lichenized Fungi
97
Table 6.1 (continued)
45a
b
46a
b
47a
b
48a
b
49a
b
50a
b
51a
b
52a
b
53a
b
54a
b
55a
b
56a
b
57a
b
58a
b
59a
b
Thallus ascendant; cortex prosoplectenchymatic, longitudinally arranged hyphae;
upper surface faintly longitudinally striate ......................................................................................................84
Thallus flattened; cortex paraplectenchymatic or prosoplectenchymatic;
hyphae not longitudinally arranged; upper surface not striate .........................................................................46
Apothecia with dark hypothecium; thallus closely appressed to substrate with adjoined lobes,
with divaricatic acid ............................................................................................................................. Dirinaria
Apothecia with pale hypothecium; thallus less closely appressed to substrate and clearly foliose,
without divaricatic acid ....................................................................................................................................47
Upper surface pale grey, K + yellow; medulla white............................................................................... Physcia
Upper surface dark grey, K-negative; medulla orange to red ....................................................... Phaeophyscia
Thallus upper surface whitish grey, with atranorin; without usnic acid ..........................................................49
Thallus upper surface yellowish grey to greenish grey; without atranorin; with usnic acid ...........................52
Lower surface pale brown to brown; ascospores < 10 mm long .......................................................................50
Lower surface dark brown to black; ascospores > 10 mm long ........................................................................51
Thallus closely appressed to rock ................................................................................................. Paraparmelia
Thallus loosely appressed to bark ............................................................................................. Pseudoparmelia
Pycnoconidia bifusiform, < 10 mm long .......................................................................................Canoparmelia
Pycnoconidia curved, > 10 mm long ...............................................................................................Parmeliopsis
Thallus lobes with rounded tips, > 2 mm wide, flat to concave; epiphytic;
lower surface black ...................................................................................................................... Flavoparmelia
Thallus lobes with slightly incised tips, 0.5-10 mm wide, flat to convex;
lower surface pale brown ...........................................................................................................Xanthoparmelia
Lobes with tomentose or felty lower side; veins, rhizines or cilia present ......................................................54
Lobes with naked, much longer than wide, smooth lower side; lacking long cilia;
underside pigmented; veins and rhizines absent, apothecia present; cilia present ....................... Heterodermia
Thallus linear, lower side with thick tomentose layer ................................................................................Anzia
Thallus irregularly lobed, lower side with thin or interrupted tomentose
layer or with felty lower side ...........................................................................................................................55
Thalli with usually large, 10-30 mm wide lobes, ascending or loosely appressed;
photobiont green algae .....................................................................................................................................56
Thallus lobes < 5 mm wide, closely appressed; upper surface tomentose;
on lower side tomentum often restricted to marginal patches; Photobiont blue-green algae;
apothecia present................................................................................................................................ Erioderma
Lower side pale brown tomentose, with large mottled spots; cyphellae
or pseudocyphellae present ..............................................................................................................................57
Lower side continuously tomentose or with larger delimited patches without tomentum;
cyphellae or pseudocyphellae absent ...............................................................................................................58
Cyphellae present; white spots on lower side with prominent, raised margin............................................Sticta
Pseudocyphellae present, yellow spots on lower side without raised margin ....................... Pseudocyphellaria
Thallus larger not sharply delimited; lobes broad and rounded; veins absent;
pseudocyphellae absent in upper side; epiphytic .................................................................................... Lobaria
Thallus wider or narrower, vein-like raised bands on lower side, thallus bright green;
black cephalodia present on the surface................................................................................................ Peltigera
Apothecia on upper surface, often absent .......................................................................................... Leioderma
Apothecia present on lower side at the tip of lobules ......................................................................... Nephroma
98
J. Sangeetha and D. Thangadurai
Table 6.2 Key for identification of fruticose lichens
1a
b
2a
b
3a
b
4a
b
5a
b
6a
b
7a
b
8a
b
9a
b
10a
b
11a
b
12a
b
13a
b
14a
b
15a
b
16a
b
17a
b
18a
b
19a
b
20a
b
21a
b
22a
b
23a
b
24a
Dimorphic, composed of fruticose and crustose or squamulose part; thallus containing algae or not;
ascocarp or basidiocarp present or absent ....................................................................................................... 2
Uniformly fruticose; thallus containing algae and ascocarps ....................................................................... 11
Thallus without algae; producing basidiocarps, subulate or mushroom-shaped ............................................ 3
Thallus with algae; producing ascocarps, branched ....................................................................................... 6
Fruticose thallus subulate................................................................................................................................ 4
Fruticose thallus mushroom-shaped ............................................................................................... Omphalina
Photobiont containing thallus crustose ........................................................................................................... 5
Photobiont containing thallus squamulose.................................................................................. Lepidostroma
Fruticose thallus < 2 cm long ....................................................................................................... Clavulinopsis
Fruticose thallus < 2 mm long; photobiont Nostoc ........................................................................Massalongia
Thallus crustose; podetia not branched, with single apothecium ................................................................... 7
Thallus squamulose; podetia branched, without apothecium ....................................................................... 10
Apothecia terminal, disc-like to globose ........................................................................................................ 8
Apothecia convex, covering thallus .....................................................................................................Cetraria
Cephalodia present on crustose thallus ............................................................................................Pilophorus
Cephalodia absent ........................................................................................................................................... 9
Apothecia terminal pink; cephalodia and phyllocladia absent ....................................................... Baeomyces
Apothecia whitish to pink; with convex disc and thin margin.............................................................. Dibaeis
Fruticose thallus branches hollow, branched; without ascocarps; cilia present ............................................ 14
Fruticose thallus branches solid, unbranched; with terminal ascocarp; cilia absent...................... Phyllobaeis
Thallus branches hollow; without arachnoid ................................................................................................ 12
Thallus branches solid, with arachnoid ......................................................................................................... 20
Thallus surface glossy, pale brown to brown; frequent perforations or pseudocyphellae ............................ 13
Thallus surface dull, whitish, greenish or greyish; perforations absent
or restricted to axils of ramifications ............................................................................................................ 14
Branches < 2 mm thick; true perforations present .................................................................................. Cladia
Branches < 2 mm thick; pseudocyphellae present ......................................................................................... 16
Thallus branches with squamules ................................................................................................................. 19
Thallus branches without squamules ............................................................................................................ 15
Epiphytic; pseudocyphellae present .............................................................................................................. 16
Pseudocyphellae absent ................................................................................................................................ 17
Thallus branches thin, < 1 mm wide; whitish ................................................................................... Oropogon
Thallus branches thick, > 1 mm wide; greenish grey ................................................................................... 24
Thallus richly branched and ascending .................................................................................. Dendriscocaulon
Thallus unbranched; without anastomoses ................................................................................................... 18
Thallus, whitish grey, subulate with smooth surface ....................................................................... Thamnolia
Thallus greenish grey to brown, with smooth or tomentose surface ............................................................ 19
Thallus surface slightly arachnoid, without cortex .............................................................................. Cladina
Thallus hallow; surface smooth, with thin cortex .............................................................................. Cladonia
Thallus branches with tough axial strand; ascospores simple ................................................................ Usnea
Thallus branches without tough central strand ............................................................................................. 21
Epiphytic ....................................................................................................................................................... 22
Epilithic ......................................................................................................................................................... 34
Thallus branches flattened; thallus greenish grey or white ........................................................................... 23
Thallus branches cylindrical; flattened sections; thallus bluish grey or greenish to whitish yellow ............ 26
Thallus < 0.2 mm long, stiff hairs composed of bundled hyphae;
apothecium disc orange, K + dark violet ......................................................................................... Seirophora
Thallus > 0.2 mm long; not hairy; apothecium disc not orange .................................................................... 24
Thallus greenish grey, > 2 cm long, pendant ................................................................................................ 32
(continued)
6
Identification Key for the Major Growth Forms of Lichenized Fungi
99
Table 6.2 (continued)
b
25a
b
26a
b
27a
b
28a
b
29a
b
30a
b
31a
b
32a
b
33a
b
34a
b
35a
b
36a
b
37a
b
38a
b
39a
b
40a
b
41a
b
42a
b
Thallus whitish, < 2 cm long, erect; surface smooth .................................................................................... 37
Thallus greenish grey to brown; with green algae; branches flattened; with green algae ............................ 26
Thallus bluish grey to black, basal branches whitish; branches cylindrical; with blue-green algae ............ 30
Thallus branches with granular to tomentose surface, grey to blue-grey ..................................................... 27
Thallus branches with smooth surface, greenish grey to brown ................................................................... 28
Thallus branching dendroid with flattened terminal ..................................................................................... 35
Thallus branching more irregular....................................................................................................... Tornabea
Thallus branches flattened, composed of apothecia ........................................................................ Polystroma
Thallus branches cylindrical ......................................................................................................................... 29
Thallus brown; branches < 0.5 mm wide, slender and branched ................................................................... 38
Thallus greenish grey; branches > 0.5 mm .................................................................................................... 40
With scattered tomentum; no apothecia; thallus with multilayered cortex,
not gelatinous ......................................................................................................................... Dendriscocaulon
Without tomentum; apothecia present; thallus with single layered cortex
or without cortex, gelatinous......................................................................................................................... 31
Richly branched; main branches < 0.1 mm wide; with tomentum; cortex
composed of a single cellular layer; algae Scytonema, gelatinous ............................................... Polychidium
Sparingly branched; main branches > 0.1 mm wide, without tomentum; no cellular cortex;
algae Nostoc in chains, not gelatinous ....................................................................................... Lempholemma
On green algae .............................................................................................................................................. 33
On blue-green algae ...................................................................................................................................... 41
Fruticose thallus with granular, coralloid, cylindrical or squamulose, phyllocladia; with green algae ........ 34
Fruticose thallus without phyllocladia; with blue-green algae ..................................................................... 35
Surface of cephalodia and phyllocladia present and smooth; apothecia terminal,
lateral or absent ............................................................................................................................ Stereocaulon
Surface of phyllocladia tomentose ................................................................................................ Leprocaulon
On soil ........................................................................................................................................................... 36
On rock or mossy tree trunks ........................................................................................................................ 38
Thallus whitish yellow, with loose, long, and slender branches ........................................................ Alectoria
Thallus whitish, with dense, short, and blunt branches ................................................................................ 37
Thallus branches flattened, never with ascocarps ................................................................................. Siphula
Thallus branches coralloid; mazaedium present ........................................................................... Acroscyphus
Thallus brown; branches rounded; < 0.5 mm thick .............................................................................. Bryoria
Thallus greenish grey to grey; branches flattened, > 0.5 mm thick .............................................................. 39
Ascocarps black with mazaedium ............................................................................................. Bunodophoron
Ascocarps yellow to grey without mazaedium ............................................................................................. 40
Thallus and ascocarps grey; cortex of palissadic structure; without cartilaginous strands................. Roccella
Thallus greenish grey, apothecial disc yellowish; cortex of strongly conglutinated,
periclinal hyphae; cartilaginous strands present ................................................................................ Ramalina
Thallus not gelatinous, white or grey..................................................................................................... Peltula
Thallus gelatinous, black .............................................................................................................................. 42
Lobes flat, unbranched, > 1 cm long .................................................................................................. Jenmania
Lobes rounded and branched, < 1 cm long ........................................................................................... Ephebe
100
J. Sangeetha and D. Thangadurai
Table 6.3 Identification key for crustose lichens
1a
b
2a
b
3a
b
4a
b
5a
b
6a
b
7a
b
8a
b
9a
b
10a
b
11a
b
12a
b
13a
b
14a
b
15a
b
16a
b
17a
b
18a
b
19a
b
20a
b
21a
b
22a
b
23a
b
Ascocarp present ...............................................................................................................................................2
Ascocarp absent; Apothecia present .................................................................................................................5
Perithecia present ..........................................................................................................................................130
Lirellae present..............................................................................................................................................169
Thallus crustose undifferentiated; perithecia spherical within thalloid warts.................................................13
Thallus crustose poorly developed; perithecia embedded in verrucose around ostiole ..................................13
Conidangia present........................................................................................................................................188
Conidangia absent .........................................................................................................................................193
Apothecia covered by powdery masses of ascospores, ripening above the asci ..............................................6
Apothecia covered by ascospores; thallus with soredia; spores globose ....................................... Srangospora
Apothecia pin-shaped, with up to 2 mm long, thin stalk ..................................................................................7
Apothecia sessile or with a short stalk ..............................................................................................................9
Ascospores simple, pale brown or colorless .....................................................................................................8
Ascospores 1-3 septate; brown color ................................................................................................... Arthonia
Apothecia dark brownish, often covered by colored pruina; ascospores pale brown ....................Chaenotheca
Apothecia reddish brown; hyaline .................................................................................................. Sclerophora
Apothecia without thalline margin..................................................................................................................10
Apothecia with thalline margin .......................................................................................................................11
Apothecia with distinct proper margin, clearly delimited; ascospores ripening
and coloring in the asci ...................................................................................................................... Pyrgidium
Apothecia without proper margin, indistinctly delimited; ascospores
ripening and coloring inside the asci ..............................................................................................................12
Ascospores one-septate ...................................................................................................................................38
Ascospores two-septate ...................................................................................................................................12
Ascospores 7-9 x 5-6 mm; excipulum wide, with corona-like extension........................................ Nadvornikia
Ascospores 9-13 x 4.5-6.5 mm; excipulum without corona-like extension ..................................................188
Thallus white to pale grey to blackish; paraphyses unbranched; hymenium hyaline ...................................210
Thallus yellowish to brownish; paraphyses slender; acicular hyaline ..........................................................210
Hymenium I + blue, throughout or around the asci; ascospores I-,
simple or variously septate, with equal cells; paraphyses straight..................................................................15
Hymenium I-; asci I-; paraphyses unbranched .............................................................................................109
Ascospores grey to brown at old, usually one-septate ....................................................................................16
Ascospores persistently hyaline, variously septate .........................................................................................17
Epiphytic on moss or plant debris...................................................................................................................17
Epiphytic on rock ............................................................................................................................................19
Ascospore wall with septa thin .......................................................................................................................19
Ascospore wall with septa thickened ..............................................................................................................18
Apothecia lecideine; ascospore wall thickening along lateral outer walls .....................................................19
Apothecia lecanorine or biatorine; ascospore wall thickening near apex and septa .......................................20
Apothecia lecanorine with thalline margin .....................................................................................................20
Apothecia lecideine without thalline margin ..................................................................................................21
Thallus margin lobate........................................................................................................................ Dimelaena
Thallus margin not lobed ................................................................................................................................22
Spores with halo, muriform ............................................................................................................................86
Spores without halo, transversely one-, rarely three-septate ..........................................................................22
Ascospore septa thin, lumina edged........................................................................................................ Buellia
Ascospore septa thickened, lumina rounded ...................................................................................................23
Apothecia lecideine; Ascospores thin-walled, pointed poles .................................................................Hafellia
Apothecia biatorine; ascospores thickened polar walls, not pointed ................................................... Rinodina
(continued)
6
Identification Key for the Major Growth Forms of Lichenized Fungi
101
Table 6.3 (continued)
24a
b
25a
b
26a
b
27a
b
28a
b
29a
b
30a
b
31a
b
32a
b
33a
b
34a
b
35a
b
36a
b
37a
b
38a
b
39a
b
40a
b
41a
b
42a
b
43a
b
44a
b
45a
b
46a
b
47a
b
Ascospores simple ..........................................................................................................................................25
Ascospores muriform ......................................................................................................................................81
Apothecia lecanorine, margin as of thallus color............................................................................................26
Apothecia lecideine or biatorine, without distinct margin ..............................................................................39
Ascospores < 20 mm long ................................................................................................................................27
Ascospores > 20 mm long ................................................................................................................................56
Ascospores > 8 in each ascus ...........................................................................................................................28
Ascospores < 8 in each ascus ...........................................................................................................................31
Thallus coarse-areolated, brown; apothecia immersed .................................................................... Acarospora
Thallus continuous or granular-areolated, grey; apothecia sessile .................................................................29
Thallus yellow; epiphytic on rock...................................................................................................................31
Thallus greenish; epiphytic on other substrates ..............................................................................................30
Asci of Lecanora-type, with distinct I + pale axial mass in tholus ...................................................... Maronina
Asci of Fuscidea-type, with continuously I + blue axial mass in tholus............................................... Maronea
Ascocarps immersed in thallus; ascus apex I ....................................................................................... Aspicilia
Ascocarps sessile in thallus; ascus apex I + blue .............................................................................................32
Hypothecium pale brown to brown.................................................................................................................33
Hypothecium colorless....................................................................................................................................34
Hymenium colorless ......................................................................................................................... Vainionora
Hymenium pale purplish .................................................................................................................. Tephromela
Thallus bright yellow ................................................................................................................... Candelariella
Thallus white or grey ......................................................................................................................................35
Conidia pleurogenous; cupular, thick-walled in excipulum ....................................................... Protoparmelia
Conidia acrogenous; cupular thick-walled in excipulum absent ........................................................ Lecanora
Ascus apex I + pale blue; spores > 8/ascus .......................................................................................................37
Ascus apex I + strongly blue; spores 4-8/ascus; ascocarps immersed in thallus warts ...................................37
Ascocarps immersed in thallus warts; ascospores < 4/ascus; epiphytic or on moss;
cephalodia absent ..............................................................................................................................Megaspora
Ascocarps lecanorine; ascospores > 8/ascus; flat, rounded cephalodia present................................... Placopsis
Ascocarps immersed in thallus warts; ascospores thick-walled (5 mm thick), > 60 mm long ........................39
Ascocarps lecanorine; ascospores thin-walled (2 mm thick), < 50 mm long .................................. Ochrolechia
Apothecial disc orange to bright red, K + purplish to dark violet ...................................................................40
Apothecial disc usually brown or black ..........................................................................................................41
Apothecial disc bright red, K + purplish......................................................................................... Pyrrhospora
Apothecial disc orange, K + dark violet; margin black ......................................................................Bahianora
Ascospores > 16 per ascus, small and globose ...................................................................................... Piccolia
Ascospores < 16 per ascus, not globose...........................................................................................................42
Paraphyses anastomosing; exciple absent; apothecia globular .......................................................................43
Paraphyses not anastomosing; exciple present; apothecia often flat ..............................................................45
Hymenial gelatine absent .......................................................................................................................Vezdaea
Hymenial gelatine present...............................................................................................................................44
Thallus yellow to green, completely sorediate; apothecia colorless ................................................. Psilolechia
Thallus grey to brown, partially sorediate; apothecia variously colored ........................................................64
Asci without tholus, I ........................................................................................................................ Schaereria
Asci with distinct, I + blue tholus ....................................................................................................................46
Asci with weakly I + blue tholus .....................................................................................................................47
Asci with strongly I + blue tholus ....................................................................................................................50
Epiphytic or on organic detritus ......................................................................................................................48
Saxicolous; ascus apex variable ......................................................................................................................51
(continued)
102
J. Sangeetha and D. Thangadurai
Table 6.3 (continued)
48a
b
49a
b
50a
b
51a
b
52a
b
53a
b
54a
b
55a
b
56a
b
57a
b
58a
b
59a
b
60a
b
61a
b
62a
b
63a
b
64a
b
65a
b
66a
b
67a
b
68a
b
Epiphytic; central amyloid tube in ascus apex present ................................................................. Malcolmiella
On organic detritus; no distinct amyloid structure in ascus apex ...................................................................49
Thallus grey; apothecia pink ...........................................................................................................................57
Thallus dark brown; apothecia black ............................................................................................ Placynthiella
Apothecia pink, usually < 1 mm wide, with thin proper margin ........................................................... Trapelia
Apothecia dark brown, 1-3.5 mm wide, with thick, prominent proper margin ........................................ Ainoa
Central tube in ascus apex present ....................................................................................................... Porpidia
Small apical cap in ascus apex present .................................................................................................. Lecidea
Asci with I + pale axial mass in tholus ............................................................................................................53
Asci with I + blue-staining tholus ....................................................................................................................54
Axial mass with Lecanora-type asci; apothecia dark brown to black;
thallus whitish, often K + yellow or C + orange .................................................................................... Lecidella
Axial mass conical; apothecia variously colored; thallus reactions different .................................................55
Asci with I + weakly blue-staining tholus; apothecia brownish to blackish discolored;
thallus grey, granular, K-, C + red.................................................................................................... Trapeliopsis
Asci with I + strongly blue-staining apical layer; apothecia dark brown to black;
thallus brown........................................................................................................................................ Fuscidea
Apothecia with distinct margin same as thallus color.....................................................................................56
Apothecia with margin same as disc color or without distinct margin ...........................................................62
Apothecia immersed, lacerate, erect margin; disc grey to pale brown,
often white-pruinose .......................................................................................................................................57
Apothecia sessile with constricted base, with entire or crenulate margin ......................................................58
Paraphyses thick and conspicuously septate ...................................................................................... Phlyctidia
Paraphyses slender, simple .................................................................................................................Phlyctella
Ascospores with thin septa..............................................................................................................................59
Ascospores with swollen septa .......................................................................................................................60
Spores one-septate; disc yellow to brown, K ......................................................................................... Lecania
Spores many-septate; disc red, K + purplish ................................................................................ Haematomma
Spores one-septate, > 15 mm wide; septa thinner than lumina; disc grey, K ........................... Megaloblastenia
Spores many-septate; septa thicker than lumina; disc yellow to brownish,
mostly K + purplish .........................................................................................................................................61
Ascospores one- to three-septate ....................................................................................................................63
Ascospores > three-septate ..............................................................................................................................81
Ascospores one-septate ...................................................................................................................................63
Ascospores many-septate ................................................................................................................................69
Ascospores 2-4/ascus, > 30 mm long; hymenium guttulate ............................................................................34
Ascospores 8/ascus, < 30 mm long; hymenium unclear..................................................................................65
Ascospores 2/ascus, > 40 mm long; hymenium clear......................................................................... Lopezaria
Ascospores 2-8/ascus, > 30 mm long; hymenium inspersed with oil-like droplets ........................................65
Ascospores with thick septa; disc K + dark violet .............................................................................. Caloplaca
Ascospores with thin septa..............................................................................................................................66
Apothecia without distinct margin ..................................................................................................................67
Apothecia with distinct margin .......................................................................................................................67
Asci with ocular chamber surrounded by I + weakly staining, rounded axial mass;
ascospores not halonate; exciple with well-defined cortical and medullary parts ..........................................69
Asci without rounded, I + weakly staining axial mass around ocular chamber;
ascospores not halonate; exciple compact ......................................................................................................68
Ascus tholus containing a conical, I + weakly staining axial mass around ocular chamber;
ascospores halonate ............................................................................................................................. Catinaria
Ascus tholus containing a tubular, I + strongly staining; apothecia yellowish;
ascospores not halonate....................................................................................................................... Catillaria
(continued)
6
Identification Key for the Major Growth Forms of Lichenized Fungi
103
Table 6.3 (continued)
69a
b
70a
b
71a
b
72a
b
73a
b
74a
b
75a
b
76a
b
77a
b
78a
b
79a
b
80a
b
81a
b
82a
b
83a
b
84a
b
85a
b
86a
b
87a
b
88a
b
89a
b
90a
b
Ascospores large, broadly ellipsoid, 75-135 x 30-40 mm, single....................................................................81
Ascospores < 10 mm wide, long fusiform to filiform, 8/ascus.........................................................................70
Ascospores with thick septa, up to 3-septate; disc mostly K + dark violet ........................................ Caloplaca
Ascospores with thin septa, up to 3-septate; disc K- ......................................................................................71
Apothecia with weak inapparent margin; apothecia convex to globose; paraphyses branched ........... Micarea
Apothecia with prominent margin; disc flat; paraphyses unbranched; sessile................................................72
Ascus tholus conical, I + weakly staining axial mass around ocular chamber ................................................73
Ascus tholus containing a tubular, I + strongly staining ..................................................................................77
Ascospores fusiform, < 5 times as long as wide .............................................................................................74
Ascospores acicular, > 5 times as long as wide ..............................................................................................76
Apothecia with brown lid-like appendage on the margin ................................................................. Auriculora
Apothecia without lid-like appendage ............................................................................................................75
Ascus tholus with rounded, I + weakly staining axial mass around ocular chamber ......................... Megalaria
Ascus tholus with pointed, I + weakly staining axial mass around ocular chamber .............................. Biatora
Excipulum paraplectenchymatic; asci with rounded axial mass.......................................................... Bacidina
Excipulum prosoplectenchymatic; asci with conical axial mass ........................................................... Bacidia
Excipulum with byssoid outer layer ................................................................................................. Byssoloma
Excipulum smooth outside..............................................................................................................................78
Apothecia < 0.5 mm side, soon convex; exciple paraplectenchymatic ............................................. Fellhanera
Apothecia usually > 0.5 mm wide, flat; exciple prosoplectenchymatic ..........................................................79
Campylidia absent; tubular structure in ascus apex distinctly
stained over its whole length.........................................................................................................Mycobilimbia
Campylidia present; tubular structure in ascus apex widened and less distinct towards the tip .....................80
Ascospores fusiform; dark brown ellipsoidal; one septate thallus distinct ....................................... Cyphelium
Ascospores needle-shaped to cylindrical; campylidia absent; pale brown with greenish .......... Microcalicium
Apothecia yellow to red, K + dark purple........................................................................................................82
Apothecia not yellow to red, K .......................................................................................................................83
Thallus white or grey .......................................................................................................................Brigantiaea
Thallus greenish yellow ...................................................................................................................... Letrouitia
Hymenium inspersed with oil-like droplets; spores single .............................................................................89
Hymenium clear; spore number per ascus various .........................................................................................84
Ascus tholus containing a tubular I + strongly staining; no campylidia present;
paraphyses unbranched ....................................................................................................................... Bapalmui
Ascus tholus without tubular, I + structure, often with wide ocular chamber;
campylidia present; paraphyses strongly branched.........................................................................................85
Ascomata with tomentose margin ...................................................................................................... Lasioloma
Ascomata with smooth margin .......................................................................................................................86
Campylidia absent...........................................................................................................................................87
Campylidia present .........................................................................................................................................88
Ascospores with gelatinous sheet .................................................................................................. Rhizocarpon
Ascospores without gelatinous sheet ................................................................................................. Schadonia
Campylidia consisting of a thalloid tube and a short, brownish;
ocular chamber of ascus wide; excipulum paraplectenchymatic ..................................................................190
Campylidia consisting of a large, greyish; ocular chamber of ascus wide;
excipulum paraplectenchymatic....................................................................................................................189
Ascocarps compound with several punctiform discs in usually raised areas
which are differently colored; ascospores transversely septate ......................................................................90
Ascocarps simple, with single disc, at age sometimes deformed; ascospores transversely muriform ...........96
Thallus felt-like, greenish ...............................................................................................................................95
Thallus with compact upper layer ...................................................................................................................91
(continued)
104
J. Sangeetha and D. Thangadurai
Table 6.3 (continued)
91a
b
92a
b
93a
b
94a
b
95a
b
96a
b
97a
b
98a
b
99a
b
100a
b
101a
b
102a
b
103a
b
104a
b
105a
b
106a
b
107a
b
108a
b
109a
b
110a
b
111a
b
Discs arranged in lines, often loosely accumulated ........................................................................................92
Discs not arranged in lines, densely accumulated ..........................................................................................94
Spores dark grey-brown; saxicolous .................................................................................. Sclerophytonomyces
Spores hyaline; usually corticolous ................................................................................................................93
Ascospores with enlarged terminal cells.........................................................................................................97
Ascospores with terminal cells not larger than median cells ..........................................................................96
Ascocarp discs wider, pruinose ............................................................................................................. Syncesia
Ascocarp discs punctiform, blackish ..............................................................................................................95
Ascospores bacillar, 3-8-septate, widest above the middle and gradually tapering towards both ends; no red
pigment in medulla .......................................................................................................................... Chiodecton
Ascospores biclavate, 4-7-septate, with a larger and a smaller swollen part;
spotted red pigment in medulla .....................................................................................................................199
Ascocarps immersed in thallus wart; hymenium separated from the margin; spores bacillar.......... Conotrema
Ascocarps sessile on smooth thallus; hymenium connected with margin; spores various .............................97
Ascocarps without margin, adnate beneath their whole hymenium; asci broadly clavate to globose, with
thick apical dome with large ocular chamber; ascospores variously septate, often with one terminal cell....98
Ascocarps with distinct, thalloid margin, often with constricted base; asci elongate, with thin apical
dome with small ocular chamber, often surrounded by small I + blue ring; ascospores transversely
septate, never with terminal cell ...................................................................................................................101
Hymenium gelatinous, not felt-like; ascocarps not clearly distinct ................................................................99
Hymenium not gelatinous, felt-like; ascocarps scarcely distinct from thallus .............................................100
Ascospores transversely septate only............................................................................................................170
Ascospores muriform ....................................................................................................................................171
Ascospores transversely septate only.................................................................................................... Stirtonia
Ascospores muriform ....................................................................................................................................194
Apothecial margin carbonized throughout, black .........................................................................................102
Apothecial margin not carbonized externally, whitish..................................................................................103
Ascospores acicular, 3-45-septate; disc permanently black............................................................ Bactrospora
Ascospores fusiform, 3-19-septate; disc often yellow-pruinose ........................................................Cresponea
Apothecia with thallus-like margin usually containing algae .......................................................................104
Apothecia biatorine to lecideine, without algae in the margin .....................................................................108
Corticolous ....................................................................................................................................................105
Saxicolous .....................................................................................................................................................107
Apothecium margin thin; apothecia not constricted at base ................................................ Krischsteiniothelia
Apothecium margin thick; apothecia constricted at base..............................................................................106
Thallus P + yellow (psoromic acid present), C ...................................................................................... Sigridea
Thallus P- (psoromic acid absent), C + red....................................................................................................107
White medulla layer under the black hypothecium ................................................................................. Dirina
Black hypothecium reaching to the substrate and confluent
with black hypothallus when present .................................................................................................Roccellina
Ascospores 1-septate; apothecia with strongly constricted base, rarely > 2 mm, usually yellowish ............109
Ascospores 5-7-septate; apothecia sessile with constricted base, > 1.5 mm wide ........................... Lecanactis
Ascospore lumina rounded at maturity; endospore usually I + violet ...........................................................110
Ascospore lumina edged, at most with lightly rounded edges, without endospore; ascospores I ................116
Hymenium separated from the surrounding thalline margin by a split; in dry state seemingly with a double
margin ...........................................................................................................................................................112
Hymenium not separated from the margin ................................................................................................... 111
Margin not carbonized; apothecia immersed in the thallus; apothecia without columella ...........................112
Margin partly carbonized and black; apothecia exerted; apothecia often
with carbonized central columella ................................................................................................................113
(continued)
6
Identification Key for the Major Growth Forms of Lichenized Fungi
105
Table 6.3 (continued)
112a
b
113a
b
114a
b
115a
b
116a
b
117a
b
118a
b
119a
b
120a
b
121a
b
122a
b
123a
b
124a
b
125a
b
126a
b
127a
b
128a
b
129a
b
130a
b
131a
b
132a
b
Apothecial margin rounded to flat; discs tiny, rarely > 0.5 mm wide ............................................................200
Apothecial margin lacerate, forming slips which cover the disc in part; discs
often several mm wide ..................................................................................................................................194
Apothecia with raised thalline margin, discs invisible through thallus splits,
pale, often white-pruinose .............................................................................................................................200
Apothecia exerted above thallus, without thalline margin; discs widely exposed, brownish.......................114
Apothecia compound with elongated discs level with the margin; ascospores hyaline,
transversely septate .......................................................................................................................................119
Apothecia simple ..........................................................................................................................................115
Ascospores hyaline, muriform ....................................................................................................... Gyrostomum
Ascospores grey, bacillary ............................................................................................................................116
Paraphyses branched and anastomosing throughout.....................................................................................117
Paraphyses unbranched, with indistinct transverse connections...................................................................119
Thallus without bristles .................................................................................................................. Gyalideopsis
Thallus with white or black bristles or hyphophores ....................................................................................118
Apothecia with constricted base and prominent margin; bristles usually black;
hyphophores absent..............................................................................................................................Tricharia
Apothecia widely adnate and with indistinct margin; bristles usually white,
when present; hyphophores often present .....................................................................................................188
Ascospores acicular, 4 mm wide ...................................................................................................................120
Ascospores ovoid to fusiform, generally over 4 mm wide ............................................................................121
Epiphytic; thallus grey; apothecia very elongated, tubular, forming 2
mm long stalks, brown .................................................................................................................... Gomphillus
Terrestrial; thallus yellow; apothecia discoid, black .................................................................... Arthrorhaphis
Apothecia immersed; ascospores grey, ovoid and muriform ........................................................ Diploschistes
Epiphytic; apothecia sessile; ascospores various ..........................................................................................122
Apothecia persistently immersed, with flat disc, often > 1 mm large, elongate; margin lacerate .................123
Apothecia initially immersed, finally sessile with constricted base, with concave disc,
mostly < 1 mm large, always rounded; margin entire....................................................................................125
Margin inapparent, granular; paraphyses with thin transverse connections;
asci I + pale blue, with thin apex ........................................................................................................ Phlyctidia
Margin lacerate; paraphyses simple; asci I-, with thickened apex................................................................124
Margin rounded to flat; discs rounded to elongate, often several mm long ..................................................170
Margin lacerate; discs rounded to slightly elongate........................................................................Chroodiscus
Apothecia forming coralloid-branched structures, greenish; ascospores 5-7-septate;
hymenium usually absent................................................................................................................. Polystroma
Apothecia not proliferating ...........................................................................................................................126
Apothecia with carbonized excipulum covered by pale pruina; ascospores muriform ....................... Ramonia
Apothecia not with carbonized margin .........................................................................................................127
Hymenium I + (pale) blue ..............................................................................................................................128
Hymenium I ..................................................................................................................................................130
Photobiont blue-green alga ............................................................................................................. Bryophagus
Photobiont Trentepohlia-like ........................................................................................................................129
Spores 8/ascus, muriform..................................................................................................................... Gyalecta
Spores 8-16/ascus, transversely septate ......................................................................................... Cryptolechia
Apothecia pale yellow to orange; ascospores one-septate ..................................................................Dimerella
Apothecia brown; ascospores bacillar-pluriseptate.............................................................................. Gyalidea
Ascospores simple ........................................................................................................................................132
Ascospores septate ........................................................................................................................................133
Ascospores 16-20/ascus ....................................................................................................................... Kalbiana
Ascospores 8/ascus .......................................................................................................................................133
(continued)
106
J. Sangeetha and D. Thangadurai
Table 6.3 (continued)
133a
b
134a
b
135a
b
136a
b
137a
b
138a
b
139a
b
140a
b
141a
b
142a
b
143a
b
144a
b
145a
b
146a
b
147a
b
148a
b
149a
b
150a
b
151a
b
152a
b
153a
b
154a
b
155a
b
156a
b
Ascospores thick-walled, with spines; paraphyses persistent ........................................................ Monoblastia
Ascospores thinwalled, smooth; paraphyses disappearing in an early stage .................................... Verrucaria
Ascospores brown or grey-brown .................................................................................................................135
Ascospores persistently colorless .................................................................................................................144
Ascospore with thin septa and edged lumina ................................................................................................136
Ascospore with thick septa and rounded lumina ..........................................................................................139
Asci ovoid with narrow ocular chamber .......................................................................................................137
Asci subcylindrical with wide ocular chamber .............................................................................................138
Ascospores with 1-3 transverse septa; ascocarps simple .........................................................Mycomicrothelia
Ascospores transversely septate to muriform; ascocarps mostly multi-chambered .....................................144
Ascospores one-septate ................................................................................................................ Clypeopyrenis
Ascospores muriform .............................................................................................................. Anthracothecium
Ascospores one-septate .................................................................................................................................140
Ascospores transversely septate only, with 3 septa ......................................................................................141
Ascospores without pigment granules in endospore ...................................................................... Distopyrenis
Ascospores with pigment granules in endospore .......................................................................Granulopyrenis
Ascospores three-septate, < 50 mm long .......................................................................................................142
Ascospores three-septate, > 70 mm long ................................................................................ Architrypethelium
Paraphyses absent; algae present in hymenium ............................................................................... Staurothele
Paraphyses present; no algae in hymenium ..................................................................................................143
Spores without longitudinal grooves, with prominent median transverse septum .............................. Pyrenula
Spores with longitudinal grooves, with prominent median longitudinal septum ........................ Sulcopyrenula
Ascospores with thickened septa and rounded lumina; ascocarps often compound ....................................145
Ascospores with thin septa and edged lumina; ascocarps usually simple ....................................................153
Ascospores transversely septate ....................................................................................................................146
Ascospores muriform ....................................................................................................................................150
Thallus poorly developed; ascocarps naked at maturity, never aggregated
in pseudostromata ..................................................................................................................... Pseudopyrenula
Thallus well developed; ascocarps immersed in pseudostromata.................................................................147
Ostioles free, apical .......................................................................................................................................148
Ostioles fused to form a compound ascocarp ................................................................................ Astrothelium
Ascospores 1-septate, > 45 mm long, with needle-shaped crystals in the wall .............................Megalotremis
Ascospores more than 2-septate, < 45 mm long, without crystals ................................................................149
Paraphyses branched; ascus apex with narrow ring surrounding a small ocular chamber;
wall thickening of ascospores pronounced at the edges ................................................................Trypethelium
Paraphyses unbranched; ascus apex with wide apical ring and wide ocular chamber;
wall thickening of the ascospore more equal ................................................................................. Lithothelium
Ostiole apical; jigsaw puzzle-like hyphae.....................................................................................................151
Ostiole lateral ................................................................................................................................................152
Ascocarps in brown shiny pseudostromata, K + red; pseudostroma wall composed of brown.......... Bathelium
Ascocarps not in brown pseudostromata, K-; wall not composed of brown ......................................... Laurera
Ostioles free ............................................................................................................................. Campylothelium
Ostioles fused ...............................................................................................................................Cryptothelium
Ascus tip thin, truncate; paraphyses unbranched ..........................................................................................154
Ascus tip thickened with an ocular chamber, rounded; paraphyses often branched.....................................157
Ascocarps with subapical whorl of black bristles ........................................................................ Trichothelium
Ascocarps without bristles ............................................................................................................................155
Ascospores transversely septate; asci with chitinoid apical ring .............................................................Porina
Ascospores muriform; asci without chitinoid apical ring .............................................................................156
Medulla white .............................................................................................................................. Clathroporina
Medulla yellow ................................................................................................................................ Myeloconis
(continued)
6
Identification Key for the Major Growth Forms of Lichenized Fungi
107
Table 6.3 (continued)
157a
b
158a
b
159a
b
160a
b
161a
b
162a
b
163a
b
164a
b
165a
b
166a
b
167a
b
168a
b
169a
b
170a
b
171a
b
172a
b
173a
b
174a
b
175a
b
176a
b
Thallus gelatinous; spores one-septate........................................................................................Pyrenocollema
Thallus subcuticular; grey or green, spores variously septate ......................................................................158
Paraphyses absent; hymenial gelatine I + reddish .........................................................................................159
Paraphyses present, persistent; hymenial gelatine I- ....................................................................................160
Spores transversely septate ................................................................................................................ Thelidium
Spores muriform .............................................................................................................................. Polyblastia
Paraphyses unbranched; macroconidia usually present, cylindrical, septate;
Photobiont Cephaleuros ......................................................................................................................... Strigula
Paraphyses branched; macroconidia more or less lacking ............................................................................161
Ascocarps simple ..........................................................................................................................................162
Ascocarps multilocular .................................................................................................................................168
Ascospores transversely septate ....................................................................................................................163
Ascospores muriform ....................................................................................................................................167
Ascospores ovoid-fusiform, 1-5-septate, > 4 mm wide .................................................................................164
Ascospores filiform, 6-10-septate, 2 mm wide ................................................................................ Celothelium
Ascospores ovoid-fusiform, 1-septate, rarely finally 3-septate ....................................................................165
Ascospores fusiform, 3-11-septate, > 4 mm wide ........................................................................ Polymeridium
Lower ascospore cell shorter; lichenized; microconidia globose to ellipsoid;
macroconidia present; ostiole often lateral ................................................................................ Anisomeridium
Lower ascospore cell longer; nonlichenized; microconidia bacillar;
macroconidia lacking; ostiole apical .............................................................................................................166
Paraphyses slender, without refractive bodies near the septa; asci clavate....................................Arthopyrenia
Paraphyses short-celled, with refractive bodies near the septa; asci obpyriform ..........................Naetrocymbe
Asci with indistinct apical thickening; ascospores 8 in each ascus; algiferous thallus present .......... Helenella
Asci with pronounced apical thickening; ascospores 2 in each ascus; nonlichenized ............................. Julella
Paraphyses indistinct, with many oil droplets................................................................................. Mycoporum
Paraphyses distinct, without oil droplets ......................................................................................................169
Ascospores with 1-3 transverse septa; asci rather cylindrical;
paraphyse cell ends with refractive bodies ....................................................................................... Tomasellia
Ascospores submuriform, with enlarged end cells; paraphyse cells without refractive bodies....... Exiliseptum
Paraphyses branched; hymenium I + red or blue; ascospore lumina not rounded,
with slightly rounded edges, septa I-; asci with rather thin tholus with small
ocular chamber surrounded by a small I + blue ring .....................................................................................171
Paraphyses unbranched; hymenium I-, rarely I + pale blue; ascospore lumina rounded,
lentiform, septa often I + blue or violet; asci with thick tholus, completely I-..............................................175
Ascocarp walls conspicuous and carbonized ................................................................................................172
Ascocarp walls indistinct, not carbonized; Asci ovoid to globose, with strongly thickened tip...................174
Ascocarps immersed in thallus with thick thalline carbonized margin;
hymenium always gelatinous ........................................................................................................................173
Ascocarps exerted with prominent carbonized margin; hymenium not gelatinous ......................................174
Ascospores muriform, hyaline; ascocarps short and exerted with thick thallus margin .........Helminthocarpon
Ascocarps variously septate, often brown at maturity; ascocarps level with thallus,
with thin thalline margin ................................................................................................................ Sclerophyton
Lirellae with closed gaping labiae; asci with I + blue cap in tip, which extends laterally;
spores transversely septate ................................................................................................................Opegrapha
Lirellae with wide open disc; asci with tiny I + blue cap in tip; spores transversely septate ...... Lecanographa
Hymenium gelatinous; ascospores muriform ................................................................................Arthothelium
Hymenium gelatinous; ascospores transversely septate ...................................................................... Arthonia
Ascocarps stellate..........................................................................................................................................177
Ascocarps single ...........................................................................................................................................180
(continued)
108
J. Sangeetha and D. Thangadurai
Table 6.3 (continued)
177a
b
178a
b
179a
b
180a
b
181a
b
182a
b
183a
b
184a
b
185a
b
186a
b
187a
b
188a
b
189a
b
190a
b
191a
b
192a
b
193a
b
194a
b
195a
b
196a
b
197a
b
198a
b
199a
b
200a
b
Ascospores muriform ....................................................................................................................................178
Ascospores transversely septate ....................................................................................................................179
Ascospores colorless; merocarps with rounded ends....................................................................... Medusulina
Ascospores grey; merocarps with pointed ends .......................................................................... Sarcographina
Ascospores brown; merocarps with rounded ends................................................................................. Glyphis
Ascospores grey to brown; merocarps with pointed ends ............................................................. Sarcographa
Mature ascospores on top of hymenium, forming a mazaedium ................................................. Schistophoron
Mature ascospores in the asci, not forming a mazaedium ............................................................................181
Ascospores one-septate, with edged lumina ..................................................................................... Melaspilea
Ascospores many-septate, with lenticular lumina ........................................................................................182
Ascocarps without lateral exciple, immersed with thin margins ........................................................ Fissurina
Ascocarps carbonized lateral exciple, raised with thickened margins ..........................................................183
Paraphyses with smooth tips; lirellae white, C + red ......................................................................... Dyplolabia
Paraphyses with smooth tips; lirellae variously colored, C- .........................................................................184
Ascospores transversely septate, with lenticular lumina ..............................................................................185
Ascospores muriform ....................................................................................................................................186
Ascospores colorless ..............................................................................................................................Graphis
Ascospores grey to brown ............................................................................................................ Phaeographis
Ascospores colorless, without lenticular lumina ..........................................................................................187
Ascospores grey to brown, with lenticular lumina .................................................................... Phaeographina
Paraphyses unbranched; ascospore lumina lenticular ......................................................................... Graphina
Paraphyses branched; ascospore lumina persistently edged ....................................................... Cyclographina
Conidangia campylidia, with ear shaped, grey or brownish ....................................................... Pyrenotrichum
Conidangia superficial black spots................................................................................................................192
Campylidia producing simple, pyriform or short-bacillar conidia ...............................................................190
Campylidia producing septate, filiform conidia............................................................................................191
Ear-shaped part short, on top of short thalloid cylinder................................................................ Sporopodium
Ear-shaped part large, directly on thallus........................................................................................ Musaespora
Campylidia grey, producing conidia without appendages ................................................................. Calopadia
Campylidia brown, producing conidia with appendages ...................................................................... Badimia
Conidia brown, simple or 2-4-celled ............................................................................................... Tylophoron
Conidia brown, obovate and 4-6-celled ..........................................................................................Sporostigma
Thallus felt-like, composed of very loose hyphae ........................................................................................196
Thallus felt-like, usually sorediate ................................................................................................................199
Thallus bright yellow; with diffuse margin......................................................................................Chrysothrix
Thallus shades of pale grey; with lobate margin...........................................................................................195
Thallus shades of grey; with diffuse margin ........................................................................................ Lepraria
Thallus shades of grey; with lobate margin ...................................................................................... Leproloma
Thallus not zoned, usually with an abrupt border, often lobed ............................................................ Crocynia
Thallus zoned, with a differentiated marginal zone of pale color .................................................................197
Thallus blue green, without isidia, with filamentose cyanobacteria ............................................... Dictyonema
Thallus pinkish to greenish, with isidia, with green algae ............................................................................198
Thallus margin whitish; with felty isidia .................................................................................. Dichosporidium
Thallus margin blackish; thallus pinkish, without isidia.............................................................. Sagenidiopsis
Medulla conspicuously colored ....................................................................................................................200
Medulla white ...............................................................................................................................................203
Medulla completely or partly red; isidiate ....................................................................................................201
Medulla orange or yellow; sorediate.............................................................................................................202
(continued)
6
Identification Key for the Major Growth Forms of Lichenized Fungi
109
Table 6.3 (continued)
201a Medulla pink throughout; with glossy, short, clavate isidia .........................................................................203
b Medulla red throughout, exposed along the margins ..................................................................... Cryptothecia
202a Thallus greenish, glossy, with scattered soralia-like, yellow spots.................................................. Myeloconis
b Thallus grey, dull, with raised, dense, yellow soralia ................................................................... Megalospora
203a With soradi fine or corsea .............................................................................................................................204
b With schizidia ...............................................................................................................................................206
204a Thallus C + red....................................................................................................................................Pertusaria
b Thallus C .......................................................................................................................................................205
205a Soredia fine; with stictic acid (P + orange, K + orange) .................................................................. Thallotrema
b Soredia coarse; with hypoprotocetraric acid (P-, K-) ...................................................................................206
206a Schizidia accumulated in groups, shortly stalked .........................................................................................207
b Schizidia arising single, leaving scattered, round scars on thallus ...............................................................207
207a Thallus bluegrey, pruinose ............................................................................................................................208
b Thallus greenish, not pruinose ......................................................................................................................208
208a Schizidia small, < 0.2 mm wide; with protocetraric acid (P + red) ..............................................................209
b Schizidia > 0.5 mm wide; with psoromic acid (P + yellow, K-) ........................................................ Ocellularia
209a Isidia cylindrical; with psoromic acid (P + yellow) ......................................................................... Myriotrema
b Isidia gradually tapering; no lichen substances (P ........................................................................... Thelotrema
210a Medulla I + blue; globuse 8 elongate, multiseptate ................................................................................ Belonia
b Medulla I-; Spores 8 globuse elipsoid ................................................................................................Biatorella
Table 6.4 Identification key for squamulose lichens
1a
b
2a
b
3a
b
4a
b
5a
b
6a
b
7a
b
8a
b
9a
b
10a
b
11a
b
12a
b
Perithecia present ................................................................................................................................................2
Apothecia present................................................................................................................................................5
Ascospores simple or transversely septate; no algae present in hymenium .......................................................3
Ascospores muriform ..........................................................................................................................................4
Ascospores simple; Squamules large; no algae in the hymenium ................................................. Catapyrenium
Ascospores transversely septate ..........................................................................................................................5
Algae present in hymenium; cortical cells of thallus smooth-walled; spores muriform, brown ......................28
Algae absent in hymenium; cortical cells of thallus finely papilose; spores simple, colorless ..........................6
Squamules rounded, whitish with raised margin; perithecia absent ................................................ Normandina
Squamules usually elongated and greenish, without raised margin....................................................................7
Photobiont green algae......................................................................................................................................24
Photobiont bluegreen algae .............................................................................................................. Psoroglaena
Photobiont bluegreen algae .................................................................................................................................8
Photobiont green algae......................................................................................................................................13
Thallus not gelatinous, closely appressed to substrate; heteromerous, lower cortex lacking or weakly
developed; with distinct cortex and medulla ......................................................................................................9
Thallus umbilicate, reddish brown, not sorediate or isidiate; apothecia immersed in warts .............. Phylliscum
Ascospores > 100 per ascus; photobiont cells in tetrads or single................................................... Phyllopeltula
Thallus blue-grey to brown, epiphytic ..............................................................................................................10
Thallus with distinct, often tomentose, prothallus; apothecia with thalline margin .......................... Parmeliella
Thallus without distinct prothallus; not umbilicate; apothecia without thalline margin...................................11
Apothecial with thalline margin; photobiont bluegreen algae,
cells in densely winding chains; ascospores simple, hyaline; isidiae or papillae bluish ..................................12
Apothecia without thalline margin; ascospores simple, hyaline; corex composed of longitudinally arranged
hyphae; thallus with concentric ridges ............................................................................................. Coccocarpia
Asci with I + blue apical plug; thallus brownish; with white spots on lobe margins ................... Fuscopannaria
Asci without I + blue apical plug; thallus usually blue-grey ................................................................. Pannaria
(continued)
110
J. Sangeetha and D. Thangadurai
Table 6.4 (continued)
13a
b
14a
b
15a
b
16a
b
17a
b
18a
b
19a
b
20a
b
21a
b
22a
b
23a
b
24a
b
25a
b
26a
b
27a
b
28a
b
29a
b
30a
b
31a
b
32a
b
33a
b
34a
b
35a
b
Upper surface of squamules byssoid, lacking cortex ........................................................................................14
Upper surface of squamules smooth, with cortex, pruinose .............................................................................15
Epiphytic; thallus squamules connected into rosette-like thalli, algiferous ........................................... Crocynia
Thallus squamules widely scattered, without algae ...................................................................Cyphellostereum
Tomentose prothallus present ...........................................................................................................................16
No tomentose prothallus ...................................................................................................................................19
Squamules with an upper and lower cortex comprised of a thin layer of cubic cells .......................................30
Squamule cortex otherwise, lower cortex usually absent .................................................................................17
Apothecium margin concolorous with thallus; with algae.................................................................... Physcidia
Apothecium margin not concolorous with thallus, without algae ....................................................................18
Ascospores generally > 25 mm, transversely many-septate .............................................................. Squamacidia
Ascospores generally < 25 mm, simple or one-septate ...................................................................... Phyllopsora
Thallus yellow to orange...................................................................................................................................20
Thallus pale grey to greenish grey or brownish ................................................................................................22
Thallus lobes ascending, 2 mm long, convex; medulla yellow; apothecia black,
globose; thallus K + dark violet .................................................................................................... Xanthopsorella
Thallus K- or K + weakly reddish......................................................................................................................21
Apothecia lecanorine, yellow............................................................................................................... Candelina
Apothecia lecideine, black ................................................................................................................................27
On soil ...............................................................................................................................................................23
On rock..............................................................................................................................................................29
Squamules whitish grey on both sides, elongate and erect;
cortical cells of thallus finely papillose; squamules delicate, lacerate ..............................................................28
Squamules more pale on lower side, appressed to substrate; ascocarps present ..............................................25
Cortical cells of thallus finely papillose; squamules delicate, lacerate ................................................. Agonimia
Squamules pale grey-green, orbicular, with concentric wrinkles, usually
with a raised sorediate margin; spores 5 septate .............................................................................. Normandina
Ascospores simple ............................................................................................................................................26
Ascospores transversely septate; apothecia lecideine ...............................................................................Toninia
Apothecia lecanorine ........................................................................................................................................33
Apothecia lecideine ...........................................................................................................................................27
Squamules whitish grey on both sides, erect ....................................................................................................28
Squamules greenish or brownish above, pale below ........................................................................................29
Cortical cells of thallus finely papillose; squamules delicate .............................................................. Agonimia
Thallus of appressed squamules;, lacerate, pale perithecial wall......................................................................35
Squamules elongated .................................................................................................................. Pseudohepatica
Squamules rounded to moderately elongated and incised, mostly < 5 mm long...............................................30
Thallus thin, corticate with a thin layer of cubic cells on both sides ..............................................Eschatogonia
Thallus without cortex ......................................................................................................................................31
Ascospores simple ............................................................................................................................................32
Ascospores transversely septate ........................................................................................................................34
Thallus C + red, with labriform soralia......................................................................................... Hypocenomyce
Thallus C-; apothecia lecanorine ......................................................................................................................33
Ascospores acicular; with capitate soralia ...................................................................................... Bacidiopsora
Ascospores fusiform to bacillary; without capitate soralia ..................................................................... Psorella
Ascospores with many in each ascus, < 10 mm long ......................................................................... Acarospora
Ascospores 8 in each ascus, > 20 mm long ........................................................................................... Placopsis
Thallus appressed squamules, 1-3 mm across, brown, pale perithelial wall ...................................... Staurothele
Thallus erect squamules; perithelial wall brown to black ................................................................. Endocarpon
6
Identification Key for the Major Growth Forms of Lichenized Fungi
Notes
1. Note that K+ or C+ denotes a positive color
reaction and K- or C- indicates that there is no
color change. A very small amount is enough,
and it can be put in eye-dropper bottles and
kept in the fridge when not in use.
2. Sodium hypochlorite (C) is common bleach; it
is best to use bleach without any additives.
Avoid getting it on your clothes.
3. Potassium hydroxide KOH (K) or alternatively sodium hydroxide is used as a 10 %
solution. Care should be taken while preparing this chemical as it is highly caustic in the
concentrated form.
4. Clean and freeze the material for future use.
Before use allow a few hours for equilibration.
5. Placing the media in the upper lid limits contamination. Discharged spores will be attached
to the agar surface either singly or groups. For
single-spore isolation reduce the discharge
time or increase the distance between the
ascocarp and the water agar medium.
Approximate spore discharge time is 24 h.
6. In some lichens, spores germinate within
1 day after dispersal.
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7
Microscopic Methods for Analytical
Studies of Fungi
De-Wei Li
Abstract
Optical microscopy is essential in mycological research and analytical
studies of fungi. This chapter describes techniques and procedures
commonly used for microscopic studies of fungi and analytical studies of
airborne fungal spores with an optical microscope.
Keywords
Fungi • Optical microscopy • Fungal spores • Fungal morphology
• Freehand sectioning • Freezing microtome • Mounts • Stains • Bright
field • Phase contrast • Differential interference contrast
Introduction
In the last decade, the development of molecular
technology has advanced fungal systematics in
an unprecedented and revolutionary way. It has
allowed mycologists to examine the phylogenetic
relationships of fungal taxonomic groups at a
molecular level from a new perspective. More
importantly, mycologists can examine the natural
essence of these organisms, rather than view them
from artificial and superficial perspectives. Such
a development has led to significant changes at
all taxonomic ranks of fungal classification. It has
also led to the discovery of “Cryptomycota” (or
Rozellida), a new phylum arguably in Mycota or
De-Wei Li (*)
Valley Laboratory, The Connecticut Agricultural
Experiment Station, 153 Cook Hill Road,
Windsor, CT 06095, USA
e-mail: dewei.li@ct.gov
even a new Kingdom in biology [1]. The final
placement of “Cryptomycota” is subject to debate
at present. One thing for sure is that this group of
organisms is a new clade on the evolutionary tree
of life and it has not been discovered before.
However, morphology-based fungal taxonomy is
an undisputable fundamental basis of modern
molecular fungal systematics. Traditional morphology-based taxonomy will still be imperative
to complete a global inventory of Fungi [2].
Fungal molecular systematics will not replace
morphology-based fungal systematics in the foreseeable future; rather, the two schools are supplementary to each other. The International Code of
Nomenclature for algae, fungi, and plants
(Melbourne Code) requires morphological Latin
or English diagnosis for valid new fungal taxon
descriptions [3]. Microscopic observation and
analysis of fungi is still a simple, economic, and
efficient way to morphologically study, characterize, and identify fungi. Many mycologists still
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_7, © Springer Science+Business Media, LLC 2013
113
114
enjoy observing the beauty of fungi and are very
excited to see an undescribed species under a
microscope. In the last 10 years, an average of
1,196 new species of fungi were described in
each year [2]. Among these newly described species, a large number of fungi (74.4%, 8,895 species) were described based on only morphological
characters without DNA sequence data from 1999
to 2009 [2]. At this time, fungal morphology and
microscopy still have practical and scientific
value in studying fungi. Microscopic methods are
widely employed in fungal research and commercial laboratories, which serve the indoor mold as
well as the indoor air quality industries.
Fungi are very diverse in their morphology.
Thus, microscopic observation of their reproductive structures, such as spores and their arrangement, conidiophores, conidiogenesis, acervuli,
picnidia, ascomata, basidiomata, and sporangiophores, is very important in the classification of
fungi and in the diagnosis of various infections of
plants, human beings, and animals.
In this chapter we mainly discuss optical
microscopy, which is commonly used in observation and analysis of fungi. Other microscopes
used in fungal research, such as scanning electronic microscope (SEM), transmission electronic
microscope (TEM), and confocal microscope,
and so on, are not covered in this chapter. A number of reference books can be consulted for techniques for using these microscopes [4, 5].
Procedures and Techniques for
Microscopic Studies of Fungi
Specimen Preparation
Pretreatment
When dried herbarium specimens are used for
study, it is necessary to rehydrate these specimens
by placing a drop of water directly on the fruiting
bodies of Ceolomycetes or Ascomycetes for a
few minutes [6]. Alternatively, one may place a
dried specimen on sterile wet filter paper in a
Petri dish or a moisture chamber and rehydrate it
overnight at room temperature. Another simple
pretreatment method is to add a drop of 5–10%
De-Wei Li
potassium hydroxide (KOH) solution to rehydrate,
soften, and clear fungal material prior to staining.
The pretreatment will make superficial fruiting
bodies of fungi much easier to be removed from
substrates or specimens. It will also make fruiting
bodies embedded in plant materials much easier
to section.
When sporulation is not present or fruiting
bodies are not mature in/on fresh plant materials,
the fresh specimen can be pretreated by placing it
on sterile wet filter paper in a Petri dish and incubating it for two to several days at room temperature to assist sporulation or maturation of fruiting
bodies.
For fungi already growing on artificial media
and fresh, living materials with sporulating fungal structures, such pretreatment is not
necessary.
Sectioning
Freehand Sectioning
Freehand sectioning of fungi growing in plant tissues often provides an adequate method for rapid
and inexpensive microscopic observation of their
structures, if done properly. This very simple
technique often leads to high-quality images for
publication.
Equipment and Materials
• A good light source, desk lamp or a reading
light.
• A comfortable position for yourself, e.g., sitting in front of a bench or desk.
• A number of double-edged razor blades, not
single-edged ones.
• A small Petri dish filled with water.
• A beaker filled with water.
• A small paintbrush (#1).
• The material to be sectioned.
Procedure
Hold the fungal material to be sectioned between
the thumb and index finger of one hand; a razor
blade held by the other hand is drawn across the
material with the edge towards the operator.
The razor slides on the index finger of the hand
7
Microscopic Methods for Analytical Studies of Fungi
115
holding the fungal material. Wet the upper
surface of the blade edge with water before sectioning. As the sections are cut, they should slide
into water and float in it. The sections in the water
are removed with a small brush onto a glass slide
for observation under a microscope. Freehand
sectioning needs to be practiced a number of
times before satisfactory sections (smooth, even,
and thin) can be cut.
If the fungal material is small or soft, it should
be placed in a small Styrofoam block or chip
made out of packing material with a slit or pith
cut on the top to hold the fungal material or use
two pieces tied together with an elastic band or a
piece of tape to hold the fungal material. The
holding material should be able to hold the fungal
material firmly, but not too rigid and hard to crush
the fungal material.
A double-edged razor blade is crucial for successful freehand sectioning. The low angle of the
edges of a double-edged razor blade allows one
to cut subject materials into thin sections. The
blade will become dull by slicing the materials
several times. Thus, it is necessary to move to an
unused portion of the edge of the blade after cutting a few sections. When full lengths of both
edges of the blade are used, the blade is no longer
useable.
To practice freehand sectioning, it is better to
start with herbaceous plant materials. The stem
should be < 5 mm in diameter and relatively soft.
Trim the materials to the size and length that are
easy to handle in your hand.
For information about operating a microtome
and preparation of samples, consult the manufacturer’s manual.
Sectioning with a Freezing Microtome
A number of brands/designs of microtomes are
available for fungal research, such as sled microtome, rotary microtome, cryomicrotome (freezing
microtome), ultramicrotome, vibrating microtome, saw microtome, and laser microtome.
The freezing microtome is commonly used.
Nag Raj [7] described a freestanding freezing
microtome (a modified Reichert freezing microtome) in detail in his monumental book
Coelomycetous Anamorphs with AppendagesBearing Conidia. Sections are cut to 10–20 mm
thick and transferred to water from the cutting
edge of the blade with a fine needle or a fine
camel or goat hair brush [8, 9].
Sample Mounting
Tapelift Mounts
To tapelift is to use a small piece of 2 × 2 to
5 × 5 mm clear/transparent tape to gently touch a
colony so that fungal structures are removed from
the colony by the sticky side of the tape with a
pair of tweezers. The tools needed for tapelift are
a roll of clear tape mounted on a tape dispenser, a
pair of tweezers, and a pair of scissors.
Procedure
1. Examine the Petri dish under a stereo microscope to locate the area or colony of interest.
2. Flame tweezers and scissors.
3. Cut off a piece of tape with desired size with
the scissors and tweezers.
4. Place the sticky side of the tape on the colony
with the tweezers and press it against the
colony gently.
5. Move the tape out of the Petri dish and flop
its sticky side up.
6. Place it on a slide.
7. Add a drop of 70% alcohol over it and let it
evaporate for 2–3 min.
8. Add a drop of water, lactic acid, or mounting
agent of choice with staining agent.
9. Place a coverslip on it.
10. Place it under a compound microscope.
11. Take photomicrographs, if necessary.
It is a rather simple, convenient, and efficient
method to prepare wet mounts for microscopic
observation with a reduced disturbance to fungal
structures. Many commercial laboratories are
routinely using this method. However, the disadvantage of this method is that the tape itself
reduces the resolution and clarity of the field and
results in reduced quality of images. By choosing
the proper tape and careful preparation, this
method still can produce quality photomicrographs, which are high enough for peer-reviewed
publications.
The quality of tape is the foremost important
factor for the quality of the wet mount. A number
116
of brands of clear, transparent, ultra-transparent,
or super clear tapes (12.7–19 mm wide) are available on the market. Titan Ultra Clear Tape (59212
95882) is considered the best one (Seifert, personal communication; 2010); unfortunately, this
brand is not available in the United States. The
author was able to purchase Titan Ultra Clear
Packing Tape (5 cm wide), which yielded satisfactory, but not super, results. Among the different
types of tape the author has tested with water, 85%
lactic acid, or 0.1% lacto-fuchsin, the best one is
Moore Crystal Clear Tape (No. CCD-134-C,
Moore Push Pin Co., Wyndmoor, PA). The fresh
mounts provide rather good clarity and resolution.
The observation should be finished in one half to
one hour. This tape will shift the color to the slight
bluish side. White balance can easily correct its
color when color photomicrographs are taken.
This tape is not available in stores, but it can be
ordered online. It is sold for torn book and blueprint repair or restoration. Mainstays Crystal Clear
Tape (imported by Walmart Canada, Mississauga,
Ontario) is rather satisfactory in resolution and
clarity. Scotch Transparent Tapes 600 (12.7 and
19 mm wide) (3 M Co., St. Paul, MN) did not produce photomicrographs suitable for publication
owing to the mosaic background induced by its
uneven glue coating. This tape is not recommended. Try several brands yourself with the
mounting media of your choice and choose one
which gives the most satisfactory results.
Scotch Crystal Clear Tape (Ultra ClearPermanent) has not been tested. Packing clear
tapes (2.5–3.75 cm wide) sold in post offices are
much thicker than other tapes. Their thickness
deteriorates resolution significantly, and they are
not suitable for microscopic fungal studies.
Wet Mounts Using a Needle
or a Cutting Tool
The other method to prepare a wet mount for
microscopic observation is to use a needle (an
inoculating needle, an insect pin no. 4, a sewing
needle, a miniature scalpel, or a custom-made
cutting tool with a small chisel point, etc.) to cut a
small piece of colony including fungal reproductive structures (conidiophores or ascomata) [10].
De-Wei Li
The points of needles or cutting edges of scalpel
should be so sharp so as to cut the colony without
too much disturbance to fungal structures. The
area to be cut on a colony should have mature
conidiophores or ascomata, but not excessive
conidia. It is helpful to use a stereo microscope to
examine the colony and determine which area to
cut first for microscopic observation. It is rather
difficult to wash excessive conidia from a cut
colony sample on a slide. When a piece of colony
sample with excessive conidia is picked, it would
be better to prepare another one. Float the cut
colony sample from the needle tip onto a slide
with a drop of 70% alcohol. When the alcohol
evaporates, apply a drop of lactic acid or a mounting medium of choice, with or without staining
agent, depending on the fungus and the optics to
be used for observation. For example, it is not
necessary to use a staining agent for dematiaceous
hyphomycetes; water or lactic acid would be
adequate.
Squash or Tease Mounts
This technique is useful for observing conidiophores and conidiogenesis in conidiomata (avervuli or picnidia) and arrangements of asci and
ascospores in ascomata as well as the tissue structures of these fruiting bodies. Often a wet mount
is prepared without sectioning for a quick
identification.
Under a stereo microscope, one or two
superficial conidiomata or ascomata can be
removed from substrates with a sharp pin without
difficulty. Dip the tip of the pin in water so that a
droplet of water at the tip of the pin can be used
to retrieve and transfer conidiomata or ascomata
to a drop of water on a slide. Place a coverslip
over the conidiomata or ascomata on the slide.
Press the coverslip gently with a needle or a rubber eraser and move it sideways to obtain a reasonably good squash mount of well-separated
conidiogenous cells or asci.
Excessive plant material attaching to fungal
fruiting bodies prevents making clean and clear
slides for observation. Rehydrating dried specimens often assists in the separation of fungal
fruiting bodies from plant materials.
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Microscopic Methods for Analytical Studies of Fungi
117
Teasing the tissues of the conidiomata or
ascomata with two fine insect pins will produce
similar, but frequently inferior results. When
teasing is used to remove excessive plant tissues
from the fungal fruiting bodies, it may improve
the quality of the mounts.
microscopic studies on certain taxonomic groups.
It is necessary to check their publications or
monographs for the detailed information on how
they prepare their specimens for their studies.
It often saves time by following these methods.
Malloch [15] indicated that some fungi are
hydrophobic and difficult to “wet,” even with a
wetting agent. He recommended adding a drop or
two of 95% ethanol to the fungal material for a
few seconds. Before the alcohol is completely
evaporated, adding a drop of a mounting medium
of your choice will improve the quality of wet
mounts.
Some staining media widely used in the past
contain lacto-phenol, such as lacto-phenol blue
[11]. Lacto-phenol cotton blue solution is used in
some research [16]. These staining agents are no
longer recommended because of health concerns
for phenol, which is readily absorbed through the
skin and known to result in some detrimental
health effects from long-term exposure [13, 17].
Thus, these staining agents are not covered in this
chapter.
Staining and Mounting
Whether to use a staining agent and what kind of
staining agent to be used is a decision each
mycologist must make based on the fungus under
study, the key diagnostic characters of the fungus
to be observed, and microscopy to be used.
It may require application of a staining agent
to observe colorless, transparent, or lightly colored fungi or fungal structures and improve the
contrast and differentiation of the fungal structures in the samples to be examined. For a pigmented fungus, a mounting medium providing a
neutral background without altering the color or
the chemical composition of the specimen is
sufficient and satisfactory for microscopic observation, such as water and lactic acid.
The function of a proper mounting medium is
to increase contrast by providing a refractive
index higher than that of a glass slide and by providing a refractive index close to that of the specimen/sample to achieve optimal transparency.
A mounting medium chemically compatible with
the specimen is preferable. A stain or a mounting
medium should retain its optimal condition, not
drying out during microscopic study or storage.
Many stains and mounting media are available
for fungal microscopic studies, water, KOH, lactic acid, lacto-fuchsin, lacto-cotton blue, modified
lacto-cotton blue (MLCB), and lacto-phenol blue
are most commonly or routinely used by mycologists and lab technicians (see Note 1). A number
of publications provide detailed information on
the stains and mounting media [11–13]. For information on special agents, such as nuclear staining
agents, and the stains to differentiate fungi in animal tissues, see references [11, 13, 14].
It should be pointed out that some mycologists
employ a special staining agent, mounting
media, or the technique they developed for their
Microscopy: Principles and Application
Optical microscope is an indispensable instrument to a mycologist. Microscopes really extend
the vision of mycologists and biologists into a
microcosmic world to study fungi microscopically. Microscopy is using microscopes to observe
samples, specimens, or structures that are invisible to the unaided eye.
Both stereo (dissecting) and compound microscopes are necessary instruments for fungal
observation and analyses.
The objective of the microscopy is to produce
a quality image showing more details of the specimen within its designed capacity.
Stereo Microscopes
The stereo microscope, also known as a dissecting microscope, is an optical microscope variant
designed for low-magnification observation of
the surface of a sample/specimen using incident
light illumination rather than transillumination
with magnification range of × 5 to × 60. It employs
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two separate optical paths, with two objectives
and two eyepieces providing slightly different
viewing angles to the left and right eyes so as to
produce a three-dimensional view of the sample/
specimen under the microscope.
There are two major types of magnification
systems in stereo microscopes: fixed magnification
and zoom magnification systems. In the former
one primary magnification is achieved by a pair
of objective lenses with different discrete
magnifications. The zoom magnification is able
to continuously change magnification within a
designed range of × 5 to × 60. Recently, newer
versions of stereo microscopes have been
improved with motor controlled zooms integrated
with digital imaging systems.
The observation of fungal materials under stereo microscope is essential before a compound
microscope is used for microscopic examination
[18]. The significance of using a stereo microscope is often not fully appreciated. A number of
morphological characters of fruiting bodies, conidiophores, and conidial orientations on the fungal colonies on natural substrates or culture media
are diagnostic under a stereo microscope. For
instance, the orientation of conidial chains and
the conidial mass shapes of Aspergillus and
Penicillium are diagnostic features for identifying some fungi of these genera to species [10,
19]. The morphological features of fruiting bodies are critical for identifying most fungi to genera or to species, such as Alternaria, Aspergillus,
Chaetomium, Penicillium, and powdery mildews.
These characteristics are undisturbed for observation under a stereo microscope, but can be lost
or at least altered due to disturbance in a wet
mount preparation. The patterns of sporulation
and conidial chains are very important characters
to differentiate species of Alternaria [20, 21].
The stereo microscope is used to observe and
locate fungal materials on the plant tissues or on
the culture media. Removal of conidiomata and
ascomata from plant materials or colonies with a
needle or a piece of tape are conducted under a
stereo microscope with an increased change of
success of picking right fungal materials to prepare wet mounts for observation under a compound microscope. It is often used to prepare the
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samples or subsamples for further observation
under a compound microscope.
It is relatively easy to use a stereo microscope.
Observation starts from low power and proceeds
to high power. Stereo microscopes are normally
equipped with an internal light source projecting
light from upper back and bottom. For better
quality of photomicrography, an external fiber
optic light source is sometimes necessary to
enhance the characters with better lighting from
an optimal angle. Occasionally, simultaneous
lighting from above and below is necessary to
observe certain fungal structures, such as conidial
orientation from a culture.
Compound Microscopes
A compound microscope is a microscope that
uses multiple lenses (4–6 objectives) to collect
light from the sample and then a pair of eyepiece
lenses to focus the light into the eye or camera
with a magnification range of × 40 to × 2000.
Slayter [22] indicated that the quality of the
image produced by a compound microscope
depends on the magnification, the resolution of
the microscope, and the contrast produced in the
image.
Microscope objectives are characterized by
two parameters: magnification and numerical
aperture [23]. Magnification is achieved by objectives (object lenses) and eyepieces (ocular lenses)
chosen for your microscope with a magnifying
range of × 4 to × 100. For most compound microscopes, a set of objectives ranges from four to six
lenses. The objective lenses are available with
magnifications of × 4, × 10, × 20, × 40, × 60, × 100
(oil immersion lens). Eyepieces of × 10, × 15, and
× 20 are available. A pair of × 10 eyepieces is
most commonly used. The total magnification =
objective × eyepiece.
Numerical aperture ranges from 0.10 to 1.25,
corresponding to focal lengths of approximately
40–2 mm, respectively. Numerical apertures can
be achieved with oil immersion as high as 1.6, at
which the highest resolution can be reached.
The resolution of an optical microscope is the
shortest distance between two points on a specimen that can be distinguished as separate entities
[24]. The resolving power of a microscope is the
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Microscopic Methods for Analytical Studies of Fungi
119
Fig. 7.1 Spores of Arcyria sp. captured in the same viewing field with bright field (a), phase contrast (b), and differential interference contrast (c)
most important technical specification or feature
of the optical system and determines the ability to
differentiate between fine details of a specimen.
The maximal resolution for an optical compound
is 0.2 mm (200 nm) [25]. The wavelengths of visible spectrum of light are from 390 to 750 nm.
The resolution of an image is limited by the
wavelength of light used to illuminate the sample.
When objects in the specimen are much smaller
than the wavelength of the light, they do not
interfere with the waves so that these objects are
indistinguishable.
Using a microscope with a more powerful
magnification does not increase its resolution
beyond 0.2 mm; it just enlarges the image.
However, objects closer than 0.2 mm will become
blurry and only be observed as an indistinguishable entity.
Contrast is the difference in visual properties
that makes an object distinguishable from other
objects and the background. Contrast is determined by the difference in the color and brightness of the object and other objects within the
same field of view under a microscope.
The contrast of a fungal sample is determined
by several factors: type of specimen, mounting
medium, degree of optical aberration, proper setting of aperture diaphragms, contrast mechanism
employed, and characteristics of the detector.
Proper usage of contrast can improve the quality
of photomicrographs.
Illumination Techniques
Three illumination techniques—bright field,
phase contrast, and differential interference contrast (DIC)—commonly used in microscopic
studies of fungi (Fig. 7.1).
Figure 7.1 illustrates the differences of spores
of Arcyria sp. in three images captured in the
same viewing field under differing contrast illumination techniques: bright field, phase contrast,
and DIC.
Bright Field Microscopy
Bright field microscopy is the simplest and the
most basic illumination technique (see Fig. 7.1a).
This illumination technique uses transmitted visible light. The advantages of the technique are the
simplicity and the minimal sample preparation
required. Its limitations are the low contrast of
most biological samples and the low resolution
due to the blur of out of focus material. It often
has difficulty to observe colorless or transparent
fungal spores and other structures clearly due to
low contrast, especially septation without aid of
staining.
Despite its limitations, the majority of the
laboratories providing services to Indoor Air
Quality and Indoor Mold Industry use bright field
illumination according to the survey conducted
by the task force of AIHA (unpublished data),
but the colorless specimens/spores are routinely
stained to increase the contrast in these labs.
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For a research laboratory, the choice of illumination techniques relies on the quality to show the
characteristic of fungal structure to be observed.
Phase contrast and differential interference
techniques make observation of unstained biological materials possible.
Phase Contrast
Phase contrast is a widely used technique showing differences in refractive index as difference in
contrast [26]. It allows observing colorless or
transparent biological materials or certain fungal
structures, such as septation in colorless spores
without the aid of staining agents. It was developed by Frits Zernike, a Dutch physicist, in the
1930s during his study of diffraction gratings. He
was awarded the Nobel Prize in 1953 for his
invention of the phase contrast microscope [27].
It necessitates a special set of objectives with
phase rings in the back focal plane of the objective. A matching set of phase rings are placed in
the condenser and these rings are marked for
their setting position. The phase objective is
required to match the specified phase setting on
the turret condenser to enable best observation
possible. The major limitations of phase contrast
technique are halo artifacts around objects and
the requirement of thin specimen preparations
(see Fig. 7.1b) [28].
Differential Interference Contrast
DIC (also known as Nomarski interference contrast) was created and introduced to microscopy
by George Nomarski, a Polish physicist and
optics theoretician in 1950s [29]. DIC is an excellent technique, but much more expensive for rendering contrast in colorless or transparent
specimens. Contrast is very good and the condenser aperture can be used fully open, thereby
reducing the depth of field and maximizing resolution. DIC uses a special prism (Nomarski prism,
Wollaston prism) in the front focal plane of the
condenser and one in the rear focal plane of the
objective [30]. It also needs a polarized light
source which is generated by two polarizing
filters installed in the light path, one below the
condenser (the polarizer), and the other above the
objective (the analyzer). This technique is widely
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used to study live biological specimens and
unstained tissues. Under DIC technique, biological specimens including fungi show a pseudocolor that deviates from their natural color.
The technique produces a three-dimensional
effect of the specimen’s surface which is actually
a monochromatic shadow-cast image showing
the gradient of optical paths for both high and
low spatial frequencies in the specimen (see
Fig. 7.1c) [31]. A DIC turret condenser usually
has a bright field illumination setting as well as
the DIC setting to match each objective. A DIC
turret condenser in some models has a bright field
and a set of phase illumination settings. DIC produces much higher resolution images than phase
contrast and bright field.
Compound Microscope Calibration,
Adjustment, Use, and Care
All compound microscopes should be equipped
with an ocular micrometer or micro-grid for measuring fungal structures during microscopic
observation, if necessary. The ocular micrometer
must first be calibrated with each objective before
it can be used. Each microscope must be individually calibrated routinely at least once a year. For
the same magnification on the microscopes of
different brand, or even different models of the
same brand, the measurements by micrometer
with the lenses may vary slightly. The ocular
micrometer in the eyepiece is measured against a
stage micrometer or a graduated slide with each
objective. The length of each division of the ocular micrometer is calculated, recorded, and documented for each objective. The calibration result
for each objective should be recorded and taped
on the microscope for quick reference.
A microscope should be used properly. It
involves proper alignment and setting-up. The
first thing is to understand your microscope and
to know the features and functions as well as the
limits of the microscope. The aperture should be
set based on the objective lens to be used. For
instance, when using a × 60 objective lens, the
aperture should be closed down to the 0.3 position on a bright field microscope to achieve the
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Microscopic Methods for Analytical Studies of Fungi
121
optimal contrast. A wide-open aperture will result
in low contrast and wash out the details in the
image against an over bright background. The
field of view should be centered. Use your microscope with care. Start with your lens on the lowest magnification to locate the area that you want
to observe and move to higher magnification lens
in a sequence. All objectives are parfocal and
parcentered by design to minimize refocusing,
when switching lenses. Understand the depth of
focus of each objective on your microscope. It
determines your workable space associated your
specimen and objective. Never crush your lens
into a slide when you focus or switch objectives.
Such an action will scratch the lens. A dirty lens
should be cleaned with lens paper and lens cleaning solution. Immersion oil should be cleaned
from the × 100 objective, when observation with
the oil lens is complete.
Eyepieces on a microscope should be adjusted
individually so that both eyes of an observer will
focus on the same plane (parfocal). If two eyes
are focused on different focus planes, it is easy to
lead to eye fatigue. To some individuals it may
result in dizziness or headache. Make yourself
comfortable ergonomically by adjusting the position and height of your microscope and chair to
avoid ergonomic fatigue or injury. A chair that is
adjustable in height and with back support is
preferred.
where the photo(s) is saved. All photomicrographs
should be annotated with scale bars. If your system does not have the feature to insert a scale bar
to a photo, the magnification or all magnifying
factors should be recorded in details. Always
save your photos to a folder with a clear and
meaningful title so that you can retrieve them
later. How often we experience frustration by the
difficulties of retrieving a photo due to hazy
memories!
When it is necessary to measure fungal structures during microscopic observation, it is recommended to measure enough numbers of a
structure under study. Scan the whole sample first
and make sure to measure individuals that represent the full range of a population. Randomly
choose the individual subject to measure. Avoid
any bias in the process of measurement. Make 30
measurements for each structure, if possible, so
that a statistical data analysis can be conducted
later. Keep in mind that the best resolution of a
light microscope is 0.2 mm. Accuracy of a measurement < 0.2 mm is meaningless. This is the
reason why for new taxon description, all measurements are rounded up to closest 0.5 mm or
whole number in some publications.
Observation and Measurement of Fungi
under Light Microscope
Microscopic observation can be used in many
areas of mycological studies, such as fungal taxonomy, identification, fungal development/biology, pathogen and host relationship, etc. Two
factors, proper preparation of samples (slide) and
proper usage of microscope determine whether a
microscopic study can obtain the data necessary
for your objectives. Take detailed notes when you
make your observations, especially a key or diagnostic character or a detail that is not able to be
captured by photomicrography with sufficient
quality. All photos should be annotated in your
notebook including the digital filing information
Photomicrographic System and Usage
Drent [32] indicated that digital photomicrographic systems are dominant in image capturing
for microscopy which demands high resolution,
retaining true color and proper usage of limited
light conditions. Digital cameras can capture
images with quality comparable to traditional
film photography. It is the norm to use digital
photomicrographs in academic papers for publishing in peer-reviewed journals including mycological journals. Photographic software programs
make editing and post image-capture process a
much easier operation. Making a photoplate is no
longer a time-consuming and laborious process.
A simple plate with four to six images can be
done within an hour. Digital images can be captured, stored, and retrieved much more easily
than traditional photomicrography with 35-mm
film. The photomicrograph system is either
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mounted on the trinocular or the ocular tube. The
ocular setup is more economical.
Color shift under a microscope can be corrected by white balance or by a neutral density
filter slide [33].
We have to be careful to edit the images that
are for scientific research. Adjusting contrast and
brightness, correcting shifted color, and cropping
out excessive edges are acceptable, but any
Photoshop alterations to the contents of images
for aesthetic and other reasons are questionable
such as removing and replacing background.
Such an action may compromise the integrity of
scientific research.
Capturing a good image with a high resolution, a proper contrast, and true color relies on
two factors or skills: (1) using the photographic
system properly, and (2) preparing a good wet
mount with your research fungal material. Read
the manual of your photographic system and
become familiar with the functions and features
of your gear prior to operation. You will avoid
unnecessary frustration and improve your photomicrographic skill. You may enjoy it during the
process.
TIFF, GIF, and JPG are the common file formats used for taking photos, especially JPG.
These formats are able to provide the quality necessary for publication without any problem. The
key requirement for publication is ³ 300 dpi. For
a photo to be published at a print size of 4 by
4 in., the photo digital size is 1.44 mb at 300 dpi
resolution. Medium quality setting for a camera
with photo size 0.5–1.5 mb is adequate. A quality
photo is determined by being in focus and having
good resolution, proper contrast, nice clarity, and
neutral color. Size of a photomicrograph does not
need to be over 2 mb in most cases. RAW is not
necessary, unless the photomicrograph needs to
be enlarged to poster size, or you need the quality
of professional photography. Working with large
size photos will significantly slow down your
computer and your work progress or you would
need to acquire a very powerful personal computer with high quality and speed. Storage and
digital filing for larger photos could be another
issue, as they will fill up your hard drive space
rather quickly.
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Microscopy for Airborne Fungal Spore
Analysis
Analysis of airborne fungal spores provides
important information to the indoor and airborne
fungi industry and IAQ professionals to help their
indoor mold investigation. Both qualification and
enumeration of fungal spores are included in the
analysis. Analysis of airborne fungal spores is
often referred to as “spore count.” It is covered
here as a separate section to discuss its procedures.
Lab materials for spore counts are similar to
those described in previous sections. No other
special material is needed.
For spore count analysis a compound microscope is commonly equipped with × 20, × 40, × 60,
and × 100 objectives [34, 35]. Objectives of × 4
and × 10 are rarely chosen for a microscope for
spore count. A × 20 lens can serve for locating
sample trace, scanning the whole trace, and identifying large spores and hydrophilic fungal spores,
such as Stachybotrys, Memnoniella, Chaetomium,
Alternaria, and Epicoccums. Objectives of × 40 or
× 60 are used for overall spore enumeration and
identification [36–39]. It is the preference of a lab
or an analyst to decide which one to use. More
and more labs tend to use a × 60 lens for better
observation of details of fungal structures and not
significantly reduced viewing area and depth of
field. The × 100 objective is occasionally used to
observe fine, diagnostic characters or structures
on small spores (< 6 mm). A number of research
labs used a × 100 objective for their research [39–
42]. Such a practice is not economical for a commercial lab because of the time restraints.
The Environmental Microbiology Proficiency
Analytical Testing task force of American
Industrial Hygienist Association conducted a survey several years ago. The survey found that the
majority of accredited environmental laboratories conduct fungal spore counts under
magnifications of × 200 to × 600 and a few laboratories use × 1000.
For spore count the ocular micrometer is used
to measure the size of fungal spores and the
dimensions of sample traces. It can also be used
as a guidepost during the analysis of the sample.
7
Microscopic Methods for Analytical Studies of Fungi
Fungal Spore Identification
It is not difficult to identify common fungal
spores to genus level, such as Alternaria,
Cladosporium,
Pithomyces,
Epicoccum,
Ganoderma, Chaetomium, Fusarium, Curvularia,
Torula, Nigrospora, Spegazzinia, etc. It is a challenge to identify some spores to genus level. This
is the reason why the Aspergillus/Penicilium-like
group is used to accommodate colorless, ellipsoid, ovoid, or subglobose spores that often
belong to more than a half dozen of different genera, Aspergillus, Pacilomyces, Penicilium,
Acremonium, Geosmithia, and Merimbla. For the
spores of Basidiomycota and Ascomycota are
often identified as basidiospores and ascospores.
Among these spores, a few may be able to be
identified to genus, such as Ganoderma and
Leptosphaeria. There is no doubt that it is necessary to conduct more research on fungal spore
morphology and their taxonomic values for fungal spore identification. Possession of a good
collection of mycological literature is vital to
fungal spore identification.
Procedures to Identify and Quantify
Spore Traps
Ideally, 100% of a sample trace should be analyzed. Analysis of spore count samples cover the
traces ranging from 25% to 100%. Only a small
number of labs analyze their samples by covering
100% of the trace. A majority analyzes a portion
of the sample by subsampling in the trace due to
time or economic restraints. For a normal spore
load, it is minimal to analyze 25% of the trace.
Occasionally the analyzed area might be reduced
to < 25%, if a spore load is extraordinarily high
(> 800 spores/sample) and has a relatively even
spore distribution. Be extremely cautious when
< 25% of the trace is analyzed. This practice is
allowed in some commercial labs with qualitycontrol measures in place. For research this practice should be avoided.
A number of methods are available for examining spore trap samples, such as transverse pass,
and longitudinal pass methods, zigzag field,
and random fields (Fig. 7.2). These methods
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illustrated in Fig. 7.2 are straightforward. The
methods have two different approaches: using the
entire field of view or using a micrometer to guide
passes and the counting process. The micrometer
method has its obvious advantage. Because the
view area is much narrower than the width of the
field of view, the analyst does not need to turn
his/her eyes to read the entire field of view. It is
much easier to focus on the area within the
micrometer. This method normally imposes less
occupational stress on eyes [43].
For pass methods when spores/conidia touch
the edge of the micrometer that is guiding the
counting pass, always count those that touch the
left edge of the micrometer and exclude any that
touch the right edge (Fig. 7.3) [43].
For random and zigzag field methods, randomly scan across the trace (3–5 fields per pass)
in a zigzag pattern. The number of fields counted
should be equivalent to 25% of the trace area or
more. Count all spores within the entire field of
view. Try to scatter fields randomly over the
entire trace. For spores present on the perimeter
of the field, count only the ones attaching or on
the perimeter on the left-hand side (Fig. 7.4), not
the ones on the right-hand side. Move the field to
the left so as to reveal the entire spore for a more
precise identification. This should be done when
all spores in the field of view are analyzed except
for the ones on the left edge. Thus, the spores
with the partial view can be identified.
Collecting Reference Data and Calculating
Trace and Coverage Area
• With a calibrated microscope, the full length
of the ocular micrometer under × 40, × 60, and
× 100 lenses can be calculated by multiplying
by 100. The length of the objective you choose
to use will be the width of passes covering
through the samples, if pass method is used.
If field-of-view method is used, the diameters
of field of view should be measured.
• Obtain dimensions of the adhesive band for the
sample deposit for each sample type from the
manuals or from the manufacturers. For example, the trace size of Air-O-Cell is 1.055 mm
× 14.4 mm (15.19 mm2); Burkard samplers,
14 mm × 2 mm (28 mm2); and Allergenco samplers, 14.5 mm × 1.1 mm (16 mm2).
124
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Fig. 7.2 Fungal sample trace reading methods. (a) Transverse passes. (b) Longitudinal passes. (c) Zigzag fields.
(d) Random fields
Fig. 7.3 Spore-counting criteria for the pass methods.
Only count the spores touching the left boundary line of
the micrometer pass. Do not count the ones touching the
right boundary line. Two vertical solid arrows indicate the
direction of a pass guided by the ocular micrometer. W is
the width of the micrometer and counting pass. The
unshaded spores are counted, and the shaded spores are
not counted
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Microscopic Methods for Analytical Studies of Fungi
125
Fig. 7.4 Spore-counting criteria of random and zigzag
field methods. The spores (unshaded) on the left half of
the perimeter of the field of view and other unshaded
spores are counted, and those (shaded ) on or touching the
right half are not counted
• Use the data collected in steps 1 and 2 to
calculate the number of passes necessary to
cover the entire trace for using pass method.
The diameter of the field of view is used to
calculate the number of fields needed to cover
the entire trace, if using the field-of-view
method. Using these results, the numbers of
passes or field-of-views necessary to cover a
portion of trace area, such as 30%, can be
calculated.
• Mark the trace with a permanent marker.
• Apply a little bit of clear nail polish to each of
four corners of the sample.
• Remove the coverslip with a pair of tweezers
and turn it over.
• Place the sample on a slide with trace facing
up and allow the nail polish to anchor the sample on the slide.
• Add a drop of staining agent on the sample.
• Place a 22 × 22 mm coverslip on the sample.
• Write down the lab ID number and the sample
ID number on the right-hand side of the slide.
Procedure of Analyzing Samples
Collected with Air-O-Cell Cassettes or
Similar Devices
Sample Preparation
• Check the sample label against the Chain of
Custody sheet.
• Fill up the data sheet with client information,
sample information (number, time, location,
air volume, samplers).
• Remove or cut the seal.
• Pry the cassette open with a weighing spatula
or a penny.
Sample Analysis
• Place the prepared slide on the stage of the
microscope.
• Scan the whole trace with the × 20 objective. If
Stachybotrys, Memnoniella, and Chaetomium
spores are found during scanning, start the
first pass/field-of-view from that location.
When the first pass/field-of-view is completed,
move to the end of the slide and start the 2nd
pass/field-of-view.
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• Move the slide to the marking place as the
starting point.
• Switch to × 40 or × 60 objective.
• Start to count and identify spores with a
multiple channel counter following the reading methods you have chosen. For less common spores that are not listed on the counter,
mark each of the spores on the data sheet
directly. Use a micrometer to guide the passes
or use field-of-view to count spores.
• Write down the location with two stage rulers
when a spore has to be measured. Move the
spore to the center of the field to measure.
Once the measurement is complete, use the
coordinates to return the location where
counting stopped to resume the counting
process.
• Take notes on the information of spores in
clumps, presence of conidiophores, hypha/
mycelium, dust load, animal hair, arthropod
parts, pollen grains, etc.
• When the other end of the trace is reached,
write down the numbers of each fungal taxon
counted on the data sheet.
• Enter the data to a computer and calculate the
concentrations of airborne fungi.
• Generate a lab report, when analyses of all
samples from the same project are complete.
The equation for calculating spore
concentration is as follows:
Spores / m 3 = total spores × (number of fields in the whole trace/number of fields counted) ×
(1,000 L / (air flow rate × sampling time)).
Procedure for Analyzing Samples
Collected with Allergenco Sampler or
Similar Devices
This sampler can collect twelve discrete samples
on the same slide.
Sample Preparation
• Remove sample slide from the slide box.
• Check the sample label against the Chain of
Custody sheet and make sure the information
matches. Check the number of samples on the
slide and the sequence of the sample collection. Mark the sample which is the first one
collected.
• Fill up the data sheet with client information,
sample information (number, time, location,
air volume, samplers).
• Mark one end of all traces on the slide with a
permanent marker
• Add two drops of staining agent on the sample
slide, if 12 samples were collected on the same
slide.
• Place a 40 × 22 mm coverslip on the sample.
For £ 6 traces on a slide, a 22 × 22 coverslip
will be used.
• Write down the lab ID number and the sample
ID number on the right-hand side of the slide.
Sample Analysis
• Same as the samples for Air-O-Cell. Choose
the longitudinal pass method to analyze multiple samples on the same slide.
Procedure of Analyzing Samples Collected
with Burkard Sampler or Similar Devices
The Burkard 7-day Recording Volumetric Spore
Sampler
(Burkard
Manufacturing
Co.,
Rickmansworth, UK or Burkard Scientific
Uxbridge, Middlesex, UK) collects airborne
samples on Melinex tape coated with adhesives.
The tape is attached to a slow rotating drum. The
drum rotates 2 mm/h. The flow rate of this sampler is 10 L/min. This is a great sampler for outdoor aeromycological studies. It is too bulky for
use indoors.
Sample Preparation
• Mark the beginning end of the Melinex tape.
• Remove the entire tape from the drum
carefully with a pair of fine forceps.
7
Microscopic Methods for Analytical Studies of Fungi
127
• Place the tape on the cutting block (or a glass
plate).
• Cut the tape with a sharp razor blade or a surgery scalpel into seven pieces. Each piece covers 24 h of sampling (48 mm in length).
• Prepare the slides by writing the date, starting
time of the samples (sampler # if using more
than one trap) onto the slides with a permanent marker.
• Lay a bead of 10% Gelvatol in distilled water,
about 45 mm long, down the center of the
slide. This works well as an adhesive to hold
the tape on the slide. The exposed segment of
tape can then be rolled onto the Gelvatol,
using fine pointed forceps, taking care not to
get bubbles under the tape. Once in place, the
tape can be readjusted to make sure it is
adhered perfectly straight. The start of the tape
should consistently be placed at the same end
of the slide.
• Wait 24 h to allow the Gelvatol to dry. At this
stage, the slides can be stored for later mounting and analysis.
• Mounting the slides. Place 5–7 drops of melted
glycerin jelly (in a 50°C hot water bath) on the
exposed tape and carefully cover the tape with
a 22 × 50 mm coverslip. Warming the slides on
a slide warmer at 48°C. Press the coverslip
gently to spread the glycerin jelly evenly.
should be conducted in a clean area (i.e., in
a hood or a clean Petri dish) to avoid
contamination.
• Wait approximately 10–30 min for the clearing process.
• Add a mounting agent.
• Place a 25 × 25 mm coverslip gently over the
sample.
• Count 25–50% or a fixed area of the filter. Or
draw a fine cross on the back of the slide and
randomly choose one quarter of the sample to
analyze.
For 37-mm filter cassettes, they are too big to
mount in a slide directly. Cut the filter into four
equal pieces with an alcohol-wiped scalpel blade
by cutting a cross over it. Randomly choose one
piece and mount it on a slide for clearing and
staining. The rest of the steps are the same as for
previous samples.
Sample Analysis
• Put a sample slide under a microscope.
• Identify and count fungal spores using one
single longitudinal traverse, or 12 bi-hourly
transverse traverses depending on the objectives of the investigation or research.
• Remaining steps are the same steps as for
Air-O-Cell. Choose longitudinal pass method
to analyze multiple samples on the same
slide.
Procedure of Analyzing Samples
Collected with 25-mm Mixed Cellulose
Ester (MCE) Filter Cassette
• Remove the MCE filter from the cassette with
the filter forceps and mount onto a slide.
• Apply a clearing agent (such as triacetin)
containing a dye to the filter. This process
Notes
Commonly Used Mounting and Staining
Agents/Reagents
Water
Water is routinely used as a mounting medium in
microscopic observation/examination in mycological research. It serves two purposes: (1) rehydration of the fungal samples/specimens [18], if
necessary and (2) retention of the natural color of
the fungus and its structures [7, 44]. When
observing living and fresh materials, application
of water as a mounting agent will retain fungi’s
natural shape, size, and color of the structures so
that these characters can be accurately recorded.
It is essential to mount fungal materials in water
to observe mucoid, noncellular appendages or
mucoid layer on ascospores. These glutinous
structures and noncellular appendages are invisible in lactic acid. The disadvantage of water as a
mounting agent is that water evaporates rather
quickly. It requires adding water regularly to the
wet mount slide to avoid the material drying out.
If the observation is not complete, the wet mount
may be sealed with nail polish. Some fungi are
hydrophobic. Adding a wetting agent to the water
128
(e.g., Triton X 100, Tween 80, or Span 60) will
improve the quality of the wet mounting and
avoid formation of air bubbles among the fungal
structures under the coverslip on the slides [15].
However, water is not regularly used as a mounting medium for microscopic analysis in commercial laboratories.
Lactic Acid (85%)
This chemical is commonly used as a mounting
medium in microscopic observation of fungi.
It retains the natural color of fungal structures
with minimal alteration. It desiccates rather
slowly. A wet mount without application of any
sealant can keep its quality for several weeks
before noticeable deterioration occurs. If a wet
mount slide is sealed with nail polish or other
sealant, the slide can be kept for several months.
When distorted fungal materials in their morphological characters due to dehydration are mounted
in lactic acid and left on a lab bench for overnight
to 24 h, the lactic acid is able to rehydrate the fungal materials and results in recovery of morphology of some fungal structures.
Gelatinous fungal structures and noncellular
appendages are invisible in lactic acid. If the fungus under observation may develop mucoid structures, it should be mounted in water to make sure
that the mucoid structures will not be missed.
When lactic acid mounting does not produce
the quality you need, a drop of a staining agent
can be added to a side of the coverslip to allow
the staining agent to diffuse inside the coverslip
without disturbing the mounting. A piece of paper
towel can be placed at the opposite side of the
coverslip to draw out excessive mounting medium
and to expedite the diffusion process.
Lacto-Fuchsin
Carmichael [45] published this agent (dissolve
0.1 g acid fuchsin in 100 mL 85% anhydrous lactic acid). It is a widely used stain and mounting
medium. It turns the cytoplasmic elements a deep
pink. Dhingra and Sinclair [11] considered it to
be superior to cotton blue. To obtain the best
observation, freshly prepared mounts should be
used, especially for photomicrography. Because
De-Wei Li
it increases the contrast of some fungal structures
and provides excellent clarity, it can be used for
temporary and semipermanent mounts for most
taxonomic groups of fungi [13]. It is very useful
to observe septation in spores and conidiogenesis
of hyaline fungi. For dematiaceous hyphomycetes or fungi with unique color/pigments, it is
not necessary to use this agent or any other staining agents, since the fungi will lose their natural
color. This agent can be diluted by adding lactic
acid to the solution to obtain a lighter pink color
to fungal materials.
Cotton Blue in Lactic Acid
This is a widely used staining-mounting medium
by staining the cytoplasm of fungal cells. The
composition of this staining medium includes
only two ingredients: 0.01 g cotton blue and
100 mL 85% lactic acid. The procedure to make
this medium is as follows: add lactic acid to a
beaker with a magnetic stirrer; place the beaker
on a hot plate; and add cotton blue powder to the
lactic acid, which is being heated and stirred until
the cotton blue has dissolved. After the solution
is cool, filter out the undissolved particles [13].
The solution can be diluted with 85% lactic acid,
if a lighter blue color is required. Permanent
slides can be made with this staining-mounting
medium by applying two coats of nail polish to
seal them.
Lactic Acid in Glycerol
To make this mounting medium, mix lactic acid,
glycerol, and distilled water in the ratio of 1:2:1
(v/v/v) [13].
Modified Lacto-Cotton Blue
The ingredients of this staining agent are: 3 mL
cotton blue stock (1.0 g cotton blue and 99 mL
85% lactic acid), 250 mL glycerol, 100 mL 85%
lactic acid, and 50 mL deionized water [46].
The stain retains the natural color of pigmented
fungal materials, such as spores/conidia and
stains hyaline and thin-walled spores and other
structures subtly. It is developed for observing
airborne fungal spores collected on a slide,
which includes spores of multiple fungal taxa
7
Microscopic Methods for Analytical Studies of Fungi
129
without overstaining the sample. Occasionally,
the resultant color may be found to be too light
for certain spores. The contrast and clarity produced by this agent are not as good as lactofuchsin. Sime et al. [46] indicated that this stain
is able to rehydrate desiccated and deformed
spores so as to regain their original shapes. The
rehydration process takes several hours or overnight. No doubt, the rehydration will assist in
identification of these; otherwise, deformed
fungal spores under a microscope may hinder
identification.
The amyloid reaction includes two subtypes
of reactions:
• Euamyloid reaction—test material turns blue
without potassium hydroxide (KOH)pretreatment.
• Hemiamyloid reaction—test material turns
red in Lugol’s solution, but shows no reaction
in Melzer’s reagent; when pretreated with
KOH, it turns blue in both reagents [48].
Melzer’s reagent is the most widely used
reagent in studies of Agaricales and Boletales.
• KOH (5–10%), potassium hydroxide [13].
• Deionized water 1,000 mL.
• Potassium hydroxide 50–100 g.
KOH is not a staining agent. It is used to soften
and clear fungal tissues as a simple and quick
method in microscopic preparations. It is rather
advantageous to examine thick, gelatinous/mucoid
material or specimens containing keratinous material, such as skin scales, nails, or hair. The cellular
material and background keratin in these samples
are digested by this reagent to reveal the fungal
material by improving visibility of fungal structures under a compound microscope. KOH is also
an agent used for determining chemical reactions
of fleshy macrofungi (mushrooms).
KOH is a very strong corrosive reagent.
Extreme care should be taken when using it or
preparing this reagent. Proper personal protection gear will help avoid personal injury.
Polyvinyl Alcohol in Lactic Acid (PVLG)
It is a permanent mounting medium. It has a minimal distorting effect on fungal material and sets
rapidly. It requires a 2- to 3-day time period for
clearing of spore content in mounted slides when
used as a clearing agent. It is composed of 1.66 g
polyvinyl alcohol, 1 mL glycerol, 10 mL lactic
acid, and 10 mL distilled water [11]. To make this
agent, mix all liquid ingredients in a dark bottle
first; then add the polyvinyl alcohol to the wellmixed liquid ingredients. The polyvinyl alcohol
dissolves slowly. Thus, a water bath at 70–80°C
is necessary to hold the bottle until the solution
becomes clear in 4–6 h. This solution can be
stored in a dark bottle for a year without deterioration of the quality.
Melzer’s Reagent
This reagent is composed of 1.5 g potassium
iodide, 0.5 g iodine, 20 mL distilled water, and
22 g chloral hydrate [13]. Melzer’s is routinely
used in a microscope slide preparation for observing amyloid reactions in fruiting bodies of
Ascomycota and Basidiomycota [44, 47]. It is
not a staining agent. It is a reagent to trigger amyloid reactions in certain fungi resulting in color
change.
• Melzer’s-positive reaction (amyloid)—test
material turns blue, bluish gray to black.
• Dextrinoid reaction (pseudoamyloid)—test
material turns brown to reddish-brown.
• Melzer’s-negative (inamyloid)—test material
does not show color change, or turn to faintly
yellowish-brown.
Acknowledgments I am grateful to Dr. James LaMondia
for his pre-submission review and to Diane Riddle for her
editorial assistance.
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8
Scanning Electron Microscopy
for Fungal Sample Examination
Eduardo Alves, Gilvaine Ciavareli Lucas,
Edson Ampélio Pozza,
and Marcelo de Carvalho Alves
Abstract
This chapter presents general and specific methods for preparing fungi,
plant, and seed tissues infected with fungi, and nematodes parasited by
fungi using scanning electron microscopy (SEM). Conventional methods
with chemical fixation as well as cryofixation are included, with details for
subsequent steps of sample preparation to get optimal fungal specimens
for visualization and examination through SEM.
Keywords
Scanning electron microscopy • SEM of seed-borne fungi • SEM of plant
pathogenic fungi • SEM of nematophagous fungi • Ultra-structure •
Methods for SEM
E. Alves
Department of Phytopathology, Ultra-structural Analysis
and Electron Microscopy Laboratory—Ufla,
Lavras, Minas Gerais 37200-000, Brazil
G.C. Lucas
Department of Phytopathology, Ultra-structural Analysis
and Electron Microscopy Laboratory—Ufla,
Lavras, Minas Gerais 37200-000, Brazil
Departamento de Fitopatologia, Universidade
Federal de Lavras, Caixa postal 3037, Lavras,
Minas Gerais 37200-000, Brazil
E.A. Pozza
Departamento de Fitopatologia, Universidade
Federal de Lavras, Caixa postal 3037, Lavras,
Minas Gerais 37200-000, Brazil
M. de Carvalho Alves (*)
Department of Soil and Rural Engineering,
Campus of the Federal University of Mato Grosso,
Av. Fernando Correa da Costa, 2367, Boa Esperança,
Cuiaba, Mato Grosso 78060900, Brazil
e-mail: marcelocarvalhoalves@gmail.com
Introduction
Fungi are small, generally microscopic, eukaryotic, usually filamentous, branched, spore-bearing organisms that lack chlorophyll [1]. They can
be obligate parasites, nonobligate parasites, or
biotrophs. Owing to these characteristics these
microorganisms can develop several interactions
with plants, animals, or the environment and can
be used to produce food and enzymes for industrial processes [2].
Scanning electron microscopy (SEM) has
caused a revolution in the study of the microscopic
world. Its advantages include bi-dimensional aspect
of the images with high depth of field; large increase
in magnitude from 10 to 1,000,000 times; rapid
image processing, digitalization, and acquisition;
ease of preparation of samples and operation; and
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_8, © Springer Science+Business Media, LLC 2013
133
134
accessible costs [3]. The images generated by
SEM are also used to enlarge the possibilities of
teaching–learning interaction based on the application of virtual reality techniques for visualization of these microorganisms [4] and to study
details of fungi taxonomy [5].
When using SEM, the goal is to investigate
the external features of a specimen. However, the
SEM can be used to probe internal cellular detail
removing the overlying material, by fracturing,
cutting, or tearing the specimen [6]. The objective of the observation will also affect how the
specimens must be prepared.
The SEM is a helpful tool to study fungi, their
interaction with other organisms, and their use in
industrial processes. It permits study of several
aspects of the morphology, such as surface details,
fungi parasitism, and saprophytism. This tool has
been used since the beginning of the 1970s to
study these microorganisms [7], and to date has
produced a lot of knowledge about fungi and
their interactions.
SEM analysis of biologic material, like fungi,
requires optimal preparation of the samples. The
prevention of degeneration processes and changes
in the material is necessary during microscopic
observation. Both problems would lead to the
formation of undesirable artifacts.
Some methods to prepare study fungi and their
interactions with others organisms are presented
in this chapter. Most of these protocols are used
frequently in the Ultra-structural Analysis and
Electron Microscopy Laboratory, a multiuser
facility at the Federal University of Lavras,
Brazil. The goal is to present the protocol applied
to fungi studies, with greater coverage about
specimen preparation and SEM application. For
further information about the application and
operation of the SEM, see references [3, 6, 8].
E. Alves et al.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
25.
26.
27.
28.
containers, centrifuge tube (15 and 50 mL),
Erlenmeyer flasks, and tissue culture plates).
Fixative solution 1: modified Karnovsky’s
solution (glutaraldehyde 2.5%, paraformaldehyde 2.0% in cacodylate sodium buffer
0.05 M, pH 7.2, CaCl2 0.001 M).
Buffer cacodylate 0.05 M.
Fixative solution 2: aqueous 1% osmium tetroxide solution 0.05 M pH 7.2.
Distilled water or sterile distilled water.
Acetone dehydration series (25, 50, 75, 90,
and 100%).
Ethanol dehydration series (25, 50, 75, 90,
95, and 100%).
30% Glycerol.
Poly-l-lysine-coated coverslip with diameter
of 13 mm.
Double-stick tape (3 M).
Double-stick carbon tape.
Aluminum stubs.
Aluminum foil.
Critical point dryer.
Sputter coater.
Desiccator container.
Silica gel.
Liquid nitrogen.
Styrofoam box (about 10 × 15 cm).
Plastic box a little bit small than Styrofoam
box.
Parafilm.
Coverslip 13 mm of diameter and 22 mm
square.
Ariel (a commercially available washing
powder—Proctor and Gamble).
Ultramicrotome or microtome.
Cryo-transfer system.
Stereomicroscope.
SEM.
Methods
Materials
Routine Protocol
These materials are used to prepare fungi for
SEM observation. The necessary materials can
vary depending on the protocol used.
1. Razor blades and scalpel.
2. Specimen collection supplies (microcentrifuge tubes, pipettes, dishes (3 and 9 cm),
The method presented below is for preparing
fungi or yeasts from solid media or from any
substrate for routine observation using conventional approaches. This method enables the
study of fungi on the surface of the substrate,
8
Scanning Electron Microscopy for Fungal Sample Examination
135
Fig. 8.1 Scanning electron micrographs of rust fungi.
(a) Puccinia nakanishiki urediniospores (lemon grass
rust). (b) Phakopsora pachyrhizi germinated urediniospore with appressorium (soybean rust). (c) Prospodium
bicolor teliospore (Tabebuia sp. rust). (d) Hemileia vastatrix urediniospore (coffee rust). Specimens were prepared
as described in section “Routine Protocol”
the morphology of hyphae, spores production,
pre-invasion events of the pathogenesis in plants
(Figs. 8.1 and 8.2), and interaction between
fungi and others organisms such as nematodes
and insects.
1. Squares or discs with about 0.5 cm of a solid
medium with mycelia of a fungus or yeast
species (after the incubation period necessary
to produce the structure that you would like to
see) or a piece of leaves, roots, stems or others
parts from infected plant or organisms must be
immersed in a microcentifuge tube with
fixative solution (modified Karnovsky’s
fixative 2.5% glutaraldehyde–2.5% paraformaldehyde, 0.05 M cacodylate buffer, CaCl2
0.001 M) at pH 7.2 and kept for 24 h or more
in a refrigerator (see Note 1).
2. Then the specimens are washed in cacodylate
buffer (three times, for 10 min each wash).
3. Post-fix in 1% osmium tetroxide aqueous solution in water for 1–4 h at room temperature.
4. Rinse three times in distilled water.
5. Follow dehydration in crescent series of acetone solutions (25, 50, 75, 90, and 100%, once
for concentrations up to 90% and twice for the
100% concentration) for 10 min each.
6. Afterwards, the samples are transferred to a
critical point dryer to complete the drying process with carbon dioxide as a transition fluid.
7. The specimens obtained are mounted on aluminum stubs (same time could be necessary to
use stereomicroscope), with a double-stick
carbon tape pasted on a film of aluminum foil
(see Note 2). Take care to keep the area to be
observed upward.
8. Coat with gold in a sputter. Keep the material
in desiccator with silica gel until observing.
9. Observe in a SEM.
136
E. Alves et al.
Fig. 8.2 Scanning electron micrographs of plant pathogenic fungi. (a) Colletotrichum gloeosporioides conidia
germinated with appressoria on coffee leaf. (b) Verticillium
sp. mycelia on culture medium. (c) C. gloeosporioides
acervuli on coffee leaf. (d) Cercospora coffeicolla conidiophores and conidia on coffee leaf. Specimens were prepared as described in section “Routine Protocol”
Sample Preparation to Observe Fungi
Inside Plant Tissue or Inside a Fungal
Fruiting Body
Glycerol Method
This is a simple method that needs just a few
materials to be developed and it is possible to get
good results (Figs. 8.3 and 8.4).
1. Fix the sample as described at step 1 in section
“Routine Protocols.”
2. Move the material to 30% glycerol (see Note
3) at least 30 min.
3. Transfer the material for a container with liquid
nitrogen. Wait for the specimen deep to the
bottom of the container.
4. Take each specimen one-to-one with a forceps
and put over a metal surface, inside a plastic
container with liquid nitrogen placed inside a
styrofoam box. Immediately, fracture the specimen in many parts as possible with a scalpel.
5. Take the parts fractured and put in a microcentrifuge tube with distilled water.
6. Follow steps 3–9 in section “Routine Protocol.”
It is possible to use SEM to see internal cellular
detail inside a fungal fruiting body or inside plant
tissue infected by fungi, if overlying material is
removed. This material can be removed by fracturing, cutting, or tearing into the specimen and
removing part of the tissue that is covering the
fungi inside the material. This can be accomplished in the following three ways.
Cutting the Tissue in Liquid Nitrogen
(Cryofracture)
This technique permits one to get smooth and
clean sections that can become possible to see
inside some materials. The process is simple and
could be done in two ways.
8
Scanning Electron Microscopy for Fungal Sample Examination
137
Fig. 8.3 Scanning electron micrographs of plant pathogenic fungi inside plant tissue. (a) Cross section of grape
leaf with uredium of the rust fungus Phakopsora euvitis.
U-urediniospore; P-paraphysis. (b) Cross section of
soybean leaf with telium of the rust fungus Phakopsora
pachyrhizi. (c) Cross section of coffee leaf petiole showing C. gloeosporioides hyphae inside the cell. (d) Hyphae
of P. euvitis inside grape leaf sponge parenchyma.
Specimens were prepared as described in section “Glycerol
Method”
Fig. 8.4 Scanning electron micrographs of the fruiting
body of Diaporthe phaseolorum f. sp. Meridionali in soybean stem. (a) General view of the ascocarp. (b) Detail of
asci with ascospores. Specimens were prepared as
described in section “Glycerol Method” (Courtesy of
Regiane Medici)
Ethanol Method
This method is a simple, rapid, and inexpensive
way to observe fine detail inside the tissue. It is
adapted from reference [9], as follows:
1. Fix tissues in 2% glutaraldehyde/0.1 M cacodylate buffer, pH 7.2.
2. Wash in 0.1 M cacodylate buffer + 5%
sucrose for 30 min.
3. Cut tissue into strips 1 mm × 4 mm (for leaves)
or small parts for fungi fruiting bodies.
4. Post fix in 2% OsO4/0.1 M cacodylate buffer,
for 2 h.
5. Wash in 0.1 M cacodylate + 5% sucrose for
30 min.
6. Dehydrate in ethyl alcohol, 70, 95, 100%
(2×), 15 min each.
138
E. Alves et al.
Fig. 8.5 Scanning electron micrographs of rust soybean
fungus (Phakopsora pachyrhizi) inside soybean leaves tissue. (a) Intern face of epidermis. Arrow hyphae; C-sponge
parenchyma cell. (b) Sponge parenchyma with hyphae of
the fungus. Arrow hyphae. Specimens were prepared as
described in section “Removing the Leaf Epidermis”
(Courtesy of Elisandra B. Zambenedetti Magnani)
7. While submerged in the final change of absolute alcohol, insert material into Parafilm
sleeves (make by rolling 2 cm strips of
Parafilm around an applicator stick). Crimp
sleeves shut on both ends (see Note 4).
8. Take this material with forceps and held
under liquid nitrogen until frozen.
9. The tissue which is visible through the
Parafilm and frozen ethanol is fractured with
a single-edge razor blade or scalpel held in a
hemostat and pre-cooled in liquid nitrogen.
10. Pick the fractured fragments up with forceps
and return to a fresh volume of absolute
ethanol.
11. Remove the Parafilm sleeve.
12. Then the specimen is critical point dried
using liquid CO2 as a transition fluid.
13. The fractured face (which is distinctly
smoother and shinier when viewed in a stereomicroscope than the other surface) is oriented
upward and secured to a specimen stub.
14. The specimens are coated with layer of
vaporized gold and can be observed in a
SEM.
treatment to remove the resin and then coated
with gold [10].
1. In one ultramicrotome, cut thick section
(1 mm) from a tissue in a resin block.
2. Take the sections with a gold wire loop to
transfer to a glass coverslip measuring 13 mm
of diameter. Allow the sections to dry flat
against the glass.
3. Put a drop of resin solvent and gradually
replace the solvent by drawing off solvent
from one side with filter paper while you add
fresh solvent at the other side with a pipette.
4. After 20 min, examine the sections in a stereomicroscope. When the traces of resin are
removed, wash the sections with methyl alcohol by filter paper/pipette method. Allow the
coverslip to dry in a dust-free place.
5. Put the coverslip on stub covered with a film
of aluminum foil using a double-stick tape.
6. Coat it in the sputter for examination in the
SEM.
Preparing Thick Sections
One alternative process to observe inside the
tissue is to make thick sections in ultramicrotome (1–2 mm). Those sections could be assembled in coverslip or on stubs and submitted to a
Removing the Leaf Epidermis
This method could be used to see the fungal
colonization inside the leaf blade in the internal
part of the epidermis and in the parenchyma
(Fig. 8.5) [11]. It is modified from the leaf fracture method [12].
1. Leaves from the species in study are inoculated, in random marked places, with a 50 mL
8
Scanning Electron Microscopy for Fungal Sample Examination
139
Fig. 8.6 Scanning electron micrographs of yeasts on polystyrene membrane. (a) General view. (b) Detail of yeast
cells. Specimens were prepared as described in section “Use of Polystyrene Surface to Study Yeasts in SEM”
2.
3.
4.
5.
6.
droplet of the spore suspension from the
fungus tested. They are kept in dew chamber
for 24 h.
The inoculated places from leaves are harvested after a time necessary to start the colonization (this time can vary from 12 to 96 h,
depended on the fungus), and cut into 3 × 3 mm
squares.
The collected material is prepared as described
up to step 7 (critical point dry) in section
“Routine Protocol.”
After mounting the specimen on stubs, take a
piece of double-stick tape and put over the
specimens. Push softly and then make a fast
upward movement to remove the epidermis.
Place the stripped part on a new stub and coat
with gold in a sputter with both specimens (it
will form two parts: the stripped epidermis
and the parenchyma part that was kept on the
surface of the first stub).
Observe in a SEM.
Use of Proteases to Clean and Reveal
Details of Interfaces Between Plant
Cells and Fungal Structures
This technique could be used to remove all cytoplasmic remnants from sectioned host cells to
examine morphology of fungal haustoria, inside
leaf cells as well as inner surface of the host cells,
adapted from references [13, 14]. This technique
was associated successfully with the process
described in section “Glycerol Method.”
1. Firstly, cut the plant tissue as described for
glycerol liquid nitrogen technique (steps
1–3).
2. Put the material in a 5% (w/v) aqueous solution of Ariel (a commercially available washing powder—Proctor and Gamble) or other
washing powder contented protease and incubate overnight (approximately 14 h) at 30°C.
3. Following incubation, the material is removed
from Ariel solution and washed for 30 min in
multiple changes of distilled water.
4. Follow steps 3–9 of the section “Routine
Protocol.”
Use of Polystyrene Surface to Study
Yeasts in SEM
This technique was developed to study bacterial
adhesion [15]; however, it is used to study adhesion, polysaccharides production, and morphology of yeasts (Fig. 8.6) grown in liquid culture or
fermentation medium in SEM (see Note 5). This
protocol is split into three major steps:
1. Prepare covered glass slides (PCGS) following the method described as follows [16]. Take
one-half of a polystyrene Petri dish and dissolve in 50 mL of amyl acetate. Clean glass
slides are dipped into the polystyrene solution
and dried overnight in a laminar flow hood to
140
Fig. 8.7 Scanning electron micrographs of Aspergillus
spp. spores on coverslip with poly-l-lysine. (a) A. tubingensis. (b) A. japonicus. Specimens were prepared as
2.
3.
4.
5.
6.
7.
8.
remove traces of the solvent and to prevent
surface contamination.
Sterilization of the PCGS inside a 50-mL centrifuge tube (Falcon tubes) for 20 min at
120°C.
In aseptic conditions, transfer 25 mL of liquid
medium and inoculate the yeast.
The centrifuge tube is transferred into an
Erlenmeyer, which is incubated inside optimal
conditions and time necessary for yeast growing in rotator agitation.
After that, remove the polystyrene surface
from the slide by passing a blade along the
edges of the slide, cut the area with the yeast,
and put it into a dish (3-cm Petri dish) with
fixative solution described in step 1 of section
“Routine Protocol.” Keep the polystyrene
membrane immersed in fixative solution with
the surface of the yeast facing upward.
Follow steps 2–4 in section “Routine
Protocol.”
Follow dehydration in increasingly more concentrated ethanol solutions (25, 50, 75, 90, 95,
and 100%, once for concentrations up to 95%
and twice for the 100% concentration) for
10 min each. The dehydration needs to be in
ethanol series because the acetone is solvent
for polystyrene.
Follow steps 6–9 in section “Routine
Protocol.”
E. Alves et al.
described in section “Study of Yeasts or Fungi Spores on
Poly-l-Lysine Coverslips” (Courtesy of Daiani Maria da
Silva)
Study of Yeasts or Fungi Spores
on Poly-L-Lysine Coverslips
This technique can be used to study yeast in liquid culture or fungi spore morphology and is
adapted from reference [6] and processed as follows (Fig. 8.7).
1. Prepare circular coverslips (13 mm of diameter) coated with poly-l-lysine. Use a swab to
pass poly-l-lysine on coverslips and wait
10 min to dry.
2. For adhering yeast cells or fungi spores, transfer 50 mL of suspension (yeast cells or spores)
to a coated coverslip and wait for 10 min.
Protect the slide from the evaporation.
3. With a fine-tip pipette take out the liquid and
gently put in 100 mL of the fixative solution
described in section “Routine Protocol” and
follow steps 2–9. Take care when pouring off
the previous solution to put on a new one, and
do not let the specimen dry out. Keep the cell
or spore layer uppermost at all times.
Fixation of Fungal Structures with
Osmium Tetroxide Vapor
This methodology can be used for imaging
soft fungi that can lose their spores (such as
Aspergillus and Penicillium species) (Fig. 8.8),
8
Scanning Electron Microscopy for Fungal Sample Examination
141
Fig. 8.8 Scanning electron micrographs of agar fungi
culture prepared using osmium tetroxide vapor technique
as described in section “Fixation of Fungal Structures
with Osmium Tetroxide Vapor.” (a) Aspergillus sp. (b)
Penicillium sp.
for entomopathogenic fungi on insects [17], exudation of conidia from fruiting bodies or present
any types of alterations due to conventional methods. It is modified from reference [18].
1. Prepare the material to be processed (it can be
an agar plug from a fungal sporulation area).
Put with care on a glass surface and keep the
fungal layer uppermost.
2. Put the specimens inside a 9-cm Petri dish.
Use a piece of moisturized filter paper to make
a dew chamber.
3. In a fume hood, place 1 mL of 2% glutaraldehyde solution in an opened container and put it
inside the Petri dish with the specimens. Keep it
closed at room temperature (22–24°C) for 2 h.
4. Move the specimens for other Petri dish, place
1 mL of 2% osmium tetroxide solution in an
opened container, and close the dish.
5. Keep the Petri dish covered with aluminum
foil inside the fume hood for 12–48 h (see
Note 6).
6. After fixation the material must be kept inside
a desiccator with silica gel for 1–2 days. The
OsO4 vapor fixation combination with silica
gel desiccation can reduce disruption satisfactorily, but specimens were not as well preserved as with conventional method.
7. The specimens obtained are mounted on aluminum stubs, with a double-stick carbon tape
pasted on a film of aluminum foil.
8. Coat with gold in a sputter.
9. Observe in a SEM in no more than 3 days.
Desiccator Method
In our lab we sometime have a problem keeping
fungi spores (such as: Aspergillus (Fig. 8.9) and
Penicillium species) attached to conidiophores to
be observed in SEM. We have observed that the
largest loss of spore has occurred at critical point
dry process. Then we developed the protocol
below with good results.
1. Take some plug from Petri dish content, a
sporulated fungal culture. Remove as much
underlying agar as possible.
2. Place some excised plug in a small Petri dish
(3 cm in diameter) with the fixative solution
described in section “Routine Protocol” and
follow the steps 2–5.
3. Afterward, the samples are transferred to a
desiccator containing silica gel to complete
the drying process.
4. Follow steps 7–9 in section “Routine
Protocol.”
Ultrasonication to Remove Fungi
Appressoria
Some pathogenic fungi, before invasion of a
plant, produce an appressorium. Under this
structure is formed a narrow hypha peg used
for penetration. To study the hole in the plant tissue that the fungus has used to penetrate through
to the epidermis it is necessary to remove the
142
E. Alves et al.
Fig. 8.9 Scanning electron micrograph of agar Aspergillus sp. culture prepared using the technique described in section “Desiccator Method”
apressorium. The technique presented below is
used with this objective (Fig. 8.10) [11].
1. Leaves from the species in study are inoculated, in random marked places, with a droplet
of the spore suspension from the tested fungus. They are kept in dew chamber for 24 h.
2. The inoculated places from leaves are harvested after a time necessary to start the
appressoria formation (this time can vary
depended on the fungus), and cut into 5 × 5 mm
squares.
3. Put the material in microcentrifuge tube with
fixative solution (modified Karnovsky’s
fixative 2.5% glutaraldehyde–2.5% paraformaldehyde, 0.05 M cacodylate buffer, CaCl2
0.001 M) at pH 7.2 and kept for 24 h or more.
4. Then specimens are washed in cacodylate buffer (three times, for 10 min each wash).
5. Move the material to a beaker with cacodylate
buffer and put the beaker into an ultrasound.
Sonicating the material for 10 s.
6. Follow steps 3–9 described in the section
“Routine Protocol.”
Fixation with Tannic Acid for Fungi
In this protocol the tannic acid is used as a
mordent for osmium tetroxide, according to the
methodology modified from reference [19]. It
can improve the fixation and increase the secondary electron emission by specimen surface. It could
be necessary for samples with low conductivity.
1. Place the fungal tissue in 2.0% glutaraldehyde
aqueous solution with 0.2% of tannic acid and
keep for 3 h.
2. Move the sample for a new container with
2.0% glutaraldehyde aqueous solution with
2% of tannic acid and keep for more 3 h.
3. Rinse in distilled water three times for 10 min
each.
4. Post-fix in 1% OsO4 aqueous solution for 1 h.
5. Rinse in distilled water for three times and follow steps 5–9 described in section “Routine
Protocol.”
Cryofixation and Freeze-Substitution
Technique for Preparing Fungal
Specimens for Scanning Electron
Microscopy
This technique could be used to improve the
quality of the fixation process [20]. This process
provides a much-improved structural and biochemical preservation of cell that is close to its
living state relative to aqueous fixation protocols.
It can be developed using many expensive
8
Scanning Electron Microscopy for Fungal Sample Examination
143
Fig. 8.10 Scanning electron micrographs of rust soybean
fungus (Phakopsora pachyrhizi) on soybean leaves. (a)
General view of urediniospore in germination forming
appessoria. (b) General view of urediniospore in germination with appessorium removed. (c) General view of ured-
iniospore in germination with appessorium not removed
totally yet. (d) Detail of a hole seen after removal of the
appressorium. Specimens were prepared as described in
section “Ultrasonication to Remove Fungi Appressoria”
(Courtesy of Elisandra B. Zambenedetti Magnani)
machines, but the following procedure is a simple
and economic method:
1. Prepare a solution 1 with 1% tannic acid and
1% glutaraldehyde in anhydrous EM-grade
acetone in a polypropylene cryovials and
one solution 2 with 1% osmium tetroxide in
anhydrous EM-grade acetone and put both
in a −85°C freezer for 1 h prior to the
cryofixation.
2. Take some agar plugs or pieces of plant tissue with fungal structures and put in 30%
glycerol and keep for 30 min.
3. Take the plugs and deep one-to-one in styrofoam box with liquid nitrogen and wait for
cooling.
4. In a hood fume collect the plugs and transfer
to a vial with the solution 1 kept in a box
with ice and transfer to a −85°C freezer and
leave there for 48–72 h.
Wash the specimen three times, at low-temperature (−85°C freezer), with 100% acetone
(P.A.) for 15 min.
Transfer the sample to solution 2 in the same
condition of step 4.
Wash again for three times in acetone as
developed in step 5.
Transfer the vial to −20°C in a freezer for 2 h.
Transfer the sample to a 4°C refrigerator for
2 h.
Afterward, the samples are transferred to a
critical point dryer to complete the drying process with carbon dioxide as a transition fluid.
Follow steps 7–9 in section “Routine
Protocol.”
5.
6.
7.
8.
9.
10.
11.
144
Fig. 8.11 Scanning electron micrographs of fungus
Magnaporthe grisea prepared using cryo-chamber system
as described in section “Use of Cryochamber in SEM to
Study Fungi.” (a) Detail of conidia and conidiophores. (b)
Conidia germinating on a plastic coverslip. (c, d) Image of
E. Alves et al.
two cultures of the fungus to show the difference to produce spores between the strains using this technique
(Images taken in Bioimaging Lab at University of
Delaware—USA advised by Dr. Kirk J. Czymmek)
Use of Cryochamber in SEM
to Study Fungi
The accessory for SEM, called cryo-transfer, permits to observe samples at low temperature. It is
possible to get excellent preservation and fast
preparation of delicate samples to be observed in
SEM, which could not be possible using other
techniques avoiding artifact formation. For
example, this method could be used to measure
the ability to produce spore by fungi in agar plates
[20, 30] or to observe activities of nematophagous fungi in soil [21] and the fungi spore germination and penetration on plants or artificial
hydrophobic surfaces (Fig. 8.11).
1. Agar plugs from a medium of fungi culture,
pieces of leaves, artificial hydrophobic membranes or nematodes with fungi in a small
2.
3.
4.
5.
plastic container (see Note 7) are taken
mounted using a double-stick carbon tape to a
metal stub and plunged into liquid nitrogen for
cryofixation for 20 s. It is possible to use
Tissue Tak with carbon graphite to paste the
specimens on stubs.
Frozen specimens are placed in a cryo-chamber of a cryo-transfer system maintained at
−170°C and transferred to a cold specimen
stage of a SEM.
Sublimation of superficial frozen water is performed by heating and maintaining the cold
specimen stage at −60°C for 10 min.
The specimens are sputter-coated with gold
(approximately 30 nm in thickness) in the
cryo-chamber.
Observe with the electron microscope at
20 kV.
8
Scanning Electron Microscopy for Fungal Sample Examination
145
Fig. 8.12 Scanning electron micrographs of fungi in
seeds prepared using technique described in section
“Scanning Electron Microscopy as a Complementary
Methodology to Identify Seed-Borne Fungi.” (a, b)
Fusarium oxysporum, presenting microconidia, (c) false
heads of microconidia formed on short conidiophores
along the hyphae, (d) terminal and intercalary phialides,
(e) clusters of conidia and (f) hyphae
Scanning Electron Microscopy
as a Complementary Methodology
to Identify Seed-Borne Fungi
describes the material and protocols to prepare
seeds for studying seed-borne fungi in SEM.
Techniques of observation of fungi in seeds with
light microscope and stereomicroscopy can be
supplemented by alternative methods with greater
precision, such as SEM (Fig. 8.12). This topic
Material
1. Materials for blotter test incubation method
for seed health analysis.
2. Eight repetitions of 50 seeds for each
species.
146
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
E. Alves et al.
Absorbent paper.
Petri dishes or equivalent containers.
Transparent lid for light.
NUV light.
Freezer.
Sodium hypochlorite solution at 1%
concentration.
2,4-D salt (sodium 2,4-dichloro-phenoxyacetate) at 5–10 ppm concentration.
Incubator with temperature and light
regulation.
Agar 0.2%.
Laminar flow hood.
Autoclave.
Dishwasher.
Microscopes, compound and stereomicroscope.
Mannitol or other osmotic compound at
osmotic potentials of −0.6 to −1.0 MPa.
Protocol for Blotter Test Incubation
Method for Preparing Seeds for Health
Analysis in SEM [5]
1. Seeds are submitted to the standard blotter
test. The blotter test preparation is conducted
in accordance with the International Seed
Testing Association criteria [22]. For instance,
it will be presented a protocol for maize, common bean and cotton seeds, but other seeds
could be prepared following this protocol with
slight alterations.
2. Seeds of maize are incubated initially 24 h in
room condition, then kept 24 h in a freezer
(−20°C), followed by 5 days under NUV light,
with 12 h photoperiod, at 20°C. To prevent
seed germination of common bean and cotton
during the incubation period, the substrate
paper must be moistened with 2,4-D salt solution, 5–10 ppm, and 0.2% agar to prevent
seeds from rolling during handling.
3. After that, seeded dishes are placed under
NUV light with 12-h photoperiod at 20°C for
7 days.
4. Seeds are individually examined with a stereomicroscope from 30 to 80 magnification and
slide mounts are prepared for observation
using light microscope, in order to confirm
fungi identity.
5. If the doubt about the fungi is kept then the
seed is prepared for SEM as described in section “Routine Protocol.”
Protocol for Blotter Test Incubation
Method with Water Restriction for
Preparing Seeds for Health Analysis
in SEM [5]
This technique to get seeds to be prepared for
SEM is similar to that from the preceding protocol, with the addition of the water restriction
technique [23].
1. Lots of healthy seeds are chosen in order to
ensure the occurrence of the inoculated fungal
species.
2. The pure colony of each pathogen is scraped
with a Drigalsky loop in order to obtain a suspension of conidia and mycelium with 1 mL
solution.
3. The solution is transferred to Petri dishes with
15 cm in diameter, with potato dextrose agar
(PDA) + mannitol medium, at −1.0 MPa.
Mannitol concentrations are obtained through
Van’t Hoff equation [24].
4. The Petri dishes are incubated in chamber
with 20°C and photoperiod of 12 h, for 5
days.
5. The seeds are disinfected in a solution of
sodium hypochlorite 1% per 3 min, then
washed with sterile water and dried in the
shade per 24 h. Then, 35 g of seeds of cotton
(Gossypium hirsutum), soybean (Glycine
max) and common bean (Phaseolus vulgaris) are distributed in single layer on the
colony of Colletotrichum gossypii var.
cephalosporioides, Colletotrichum truncatum and Colletotrichum lindemuthianum,
respectively.
6. A suspension of conidia with concentration of
1 × 106 conidia/mL is sprayed on the single
layer of seeds to ensure the inoculation.
7. After that, the plates returned to the chamber
of 20°C with photoperiod of 12 h, per 144 h.
8. Follow steps 4 and 5 in the section “Protocol
for Blotter Test Incubation Method for
Preparing Seeds for Health Analysis in
SEM.”
8
Scanning Electron Microscopy for Fungal Sample Examination
147
Fig. 8.13 Scanning electron micrographs of nematophagous fungi culture prepared using the technique described
in section “Preparation of Samples to Study
Nematophagous Fungi.” (a) General view of the fungus
trapped the nematode. (b) Detail of a trap
Preparation of Samples to Study
Nematophagous Fungi
Preparation of Samples to Study Nuclei
and Chromosomes of Fungi in SEM
This protocol describes the process to prepare
nemathophagous fungi for SEM from in vitro
trails. It is modified from reference [25]
(Fig. 8.13).
1. In a fume hood transfer an agar disk of nematophagous fungus from single spore culture
for a Petri dish with medium and put the dish
into a growth chamber at 25°C, in dark for 5
days.
2. Pour on 1 mL of a nematode suspension (about
100 specimens) on the dish and incubate again
under same conditions. Observe if 50 or more
nematodes are captured, then pour on fixative
solution described in the section “Routine
Protocol” to cover the mycelia.
3. Keep in refrigerator for 72 h to complete the
fixation.
4. Then the dish is washed in cacodylate buffer
(three times, for 10 min each wash).
5. Agar disks are taken from the medium carefully, moved for a vial and post-fix in 1%
osmium tetroxide aqueous solution in water
for 1–4 h at room temperature.
6. Follow steps 4–9 in section “Routine
Protocol.”
This technique is important to determine whether
fungal nuclei and chromosomes have structures
distinct from those of plants and animals, and
give information about the manner of chromatin
compaction into condensed chromosomes during
nuclear division in fungi. The protocol presented
her is for Cochliobolus heterostrophus and
Neurospora crassa based on reference [26] but
this technique can be used for other fungi with
some changes as reviewed in reference [27].
1. Take 200 mL droplet of conidial suspension
(1.5 × 105/mL) containing 30 mM hydroxyurea (HU) and 3% (w/v) glucose and incubated
to germinate on a glass coverslip under optimal conditions for the fungus.
2. After washing with distilled water to remove
HU, 200 mL of 3% glucose is added to the
germinated conidia adhering to the slide and
incubation is resumed for 30 min.
3. After washing away the medium with water,
germinated conidia on the slide are incubated
in newly added 200 mL Vogel’s medium containing 50 mg/mL thiabendazole for more than
2 h. Thiabendazole treatment is used to arrest
nuclear division at metaphase.
148
4. After completing incubation, the germlings of
fungus is rinsed with water and treated with
0.05% glutaraldehyde (buffered with 50 mM
cacodylate, 2 mM MgCl2, pH 7.2) for 30 s.
5. The slide is then immersed in the fixative
(99.5% methanol:glacial acetic acid = 7:3) for
2 h at room temperature. Bursting of the germling cells occurred upon immersion in the
fixative to discharge the nuclei and chromosomes from cells and spread them on the glass
slide surface.
6. Follow steps 1–9 in section “Routine Protocol,”
with a slight difference at step 7, where the
coverslip is put over the double-stick tape on
stub.
Other Methods to Grow and to Take
Fungi Material for Preparing Samples
for SEM
Using Microculture of Fungi in Coverslip
1. In a Petri dish, place two sheets of filter paper
and pour on distilled water to prepare a moisture chamber.
2. Take a glass slide for microscopy and put two
agar disks, with the fungi mycelia took from
the edge of a good fungal culture. Put these
disk about 1.5 cm away one from the other
with the mycelia up.
3. Place a coverslip on the two disks. Cover the
dish, and incubate under optimal conditions
for the fungus. Wait the fungus colonization
on the coverslip.
4. Remove the coverslip with the attached mycelia and spores and put in a small Petri dish
(3 cm), with fixation solution from section
“Routine Protocol.” Keep the mycelia layer
uppermost.
5. Follow steps 2–9 in section “Routine Protocol,”
with a slight difference at step 7, where the
coverslip is put over the double-stick tape on
stub.
Using Filter for Mycelia Preparation
This method could be used to take mycelia and
conidia from a fungus growing in an agar media
for preparation for SEM. We have used this
E. Alves et al.
technique to evaluate the effect of fungicides,
plant extracts, essential oils, or other substances
on mycelial and conidia of fungi [28].
1. The substance that will be evaluated is transferred through an agar disk covered with
mycelia of fungus. The dish is transferred into
a growth chamber in optimal conditions for
the fungus to grow.
2. After fungus have grown, pour distilled water
on the dish and prepare a hyphal and conidial
suspension, using a spatula to remove the
mycelia and filter in Millipore membrane
PTFE hydrophilic of pore size 0.45 mm and
diameter of 13 mm.
3. Take the Millipore membrane and prepare
for SEM as described in section “Routine
Protocol.”
Using Cellophane Membrane to Prepare
Fungal Sample
This technique could be used to study the hyphal
interactions of one biological control fungi with
one plant pathogenic fungi or for collect hyphae
from one fungus to study morphologic characteristic modified from reference [29].
1. A cellophane membrane is placed on an agar
media in a Petri dish.
2. An agar disk covered with mycelia of antagonistic fungus is put on one end of the dish and
the plant pathogenic fungus on the other. The
fungi are placed to grow in a growth chamber
under optimal conditions for both fungi.
3. After growth, the cellophane from the interaction area is cut, fixed, and prepared for SEM
following steps 5–8, as described in section
“Use of Polystyrene Surface to Study Yeasts
in SEM.” If the goal is to observe just one fungus then a good area (with characteristic structures of the fungus) could be selected in any
part of the cellophane.
Notes
1. The fungi must be in Petri dishes on an appropriate media to obtain typical and good growth
and sporulation. When cutting the agar disk,
remove as much underlying agar as possible.
8
2.
3.
4.
5.
6.
7.
Scanning Electron Microscopy for Fungal Sample Examination
Other fixative solution could be used, but in
our laboratory we have used this solution as
standard. The sample could be kept in this
solution, in a refrigerator, for a long time (6
months or more depending on the type of
material) for SEM preparation.
In our laboratory, in order to make it easier to
clean the stubs, we are using a piece of aluminum foil to cover this support. An aluminum
foil square (about 2.5 × 2.5 cm) is cut and
placed on the stub. It is a good idea to make
certain that the foil is affixed well.
Glycerol is a cryoprotector, and we have used
it because it is cheaper and easier to use. 20%
sucrose and 15% dextran could be used.
The Parafilm sleeves are used so that the section does not move.
Stainless steel, plastic, or glass surface chips
of 10 × 20 mm could be used to study yeast in
place of the polystyrene membrane. The chips
can be placed inside an Erlenmeyer with liquid culture and kept under agitation for a time
necessary to grow the yeast. Collect the chips
and prepare as described in section “Use of
Polystyrene Surface to Study Yeasts in SEM.”
The time depends on the size and type of the
material. The fixation process occurs slowly
owing to the osmium vapor. If the material
becomes black, it is a good signal. If it does
not turn black, it could be necessary to change
the osmium tetroxide solution as often as is
necessary.
Fill a small plastic container (ca.
1.5 × 0.5 × 0.3 cm) with 3 mm soil layers. The
fungus is added to the soil together with nematodes. The specimens are incubated in a
moisture chamber with optimal temperature,
for 3–5 days. After incubation, the plastic is
frozen and prepared as described in step 1.
With cryo-SEM, it is possible to study soil
colonization and nematode parasitism by the
introduced fungus, as well as the formation
and action of traps.
Acknowledgements The authors gratefully acknowledge CNPq (Conselho Nacional de Desenvolvimento
Científico e Tecnológico), CAPES (Coordenadoria de
Aperfeiçoamento de Pessoal de Nível Superior) and
149
FAPEMIG (Fundação de Amparo à Pesquisa do Estado de
Minas Gerais—Brazil) for financial support to Ultrastructural analysis and electron microscopy laboratory of
the Ufla, Brazil.
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9
High-Resolution Imaging and Force
Spectroscopy of Fungal Hyphal Cells
by Atomic Force Microscopy
Biplab C. Paul, Hui Ma, Laelie A. Snook,
and Tanya E.S. Dahms
Abstract
Various forms of microscopy applied to image fungal hyphae have been
limited by either diffraction or the ability to image viable specimens. The
advent of the atomic force microscope offered the opportunity to image
live fungal hyphae under ambient conditions at very high resolution. The
force spectroscopy capabilities of the microscope facilitate physical and
chemical characterization of the fungal cell surface. In this chapter, we
describe the detailed protocols that have allowed high-resolution imaging
and force spectroscopy of fungal spores, germinants, and hyphae for both
fixed and viable specimens.
Keywords
Adhesion • Atomic force microscopy • Force spectroscopy • Cell wall
ultrastructure • Hyphae • Spore • Viscoelasticity
Introduction
Conventional imaging techniques for probing
the ultrastructure of the fungal cell surface—
scanning electron microscopy (SEM), for example—require rigorous sample preparation, which
B.C. Paul • T.E.S. Dahms (*)
Department of Chemistry and Biochemistry,
University of Regina, Regina, SK, Canada S4S 0A2
e-mail: tanya.dahms@uregina.ca
H. Ma
Department of Chemistry, National University of
Singapore, Singapore
L.A. Snook
Department of Human Health and Nutritional Sciences,
50 Stone Road East, Guelph, Ontario, N1G 2W1 Canada
can alter the native state of the cell. The development of cryo-SEM allowed cell imaging without
the need for critical point drying and gold coating,
but since electron microscopy requires vacuum,
the imaging of cells under ambient conditions
remained illusive. The advent of atomic force
microscopy (AFM) presented the opportunity to
overcome these limitations and provide the highest possible topographic and lateral resolution of
cell surfaces with little or no sample preparation.
In addition to its ability to provide ultrastructural
information comparable to that obtained by cryoSEM, AFM can also be used to probe chemical
and physical surface characteristics. Integration of
AFM with optical microscopes compensates for
the microscope’s relatively small field of view
(100 × 100 mm). AFM has become a very popular
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_9, © Springer Science+Business Media, LLC 2013
151
152
tool in microbiology for observing cell morphology, cell surface ultrastructure, chemical and physical properties of microbes and eukaryotic cells.
The heart of the AFM instrument is the microfabricated tip on the end of a metal-coated cantilever. Three piezoelectric micropositioners (X, Y, and
Z) maintain a constant distance between the tip and
sample (Z), as the tip is raster scanned (X, Y) over
the sample surface (tip scanning) or vice versa
(sample scanning). The optical lever, consisting of
a laser reflected from the top surface of the cantilever into a four quadrant photodetector, is the most
common AFM detection method [1]. Changes in
topography and tip–sample interactions give rise to
cantilever deflection or lateral movement registered
by the photodetector as a vertical or horizontal displacement, respectively. This signal is sent to the
computer for data collection and to the electronic
control unit (ECU), which adjusts the height (Z) of
the AFM tip through a fast feedback loop. Vertical
and horizontal displacement of the laser in the photodiode is registered as topography and tip–sample
interactions, respectively. For rough samples, lateral force images are convoluted with topographic
signals as the tip encounters large changes in the
slope, giving rise to the “edge effect,” [2] which
improves contrast and helps delineate surface feature boundaries for live samples [3].
Immobilization of the sample onto a solid surface is mandatory for imaging cells, fixed or viable, by AFM. Otherwise, raster scanning of the
AFM tip can dislodge the sample. Imaging in
noncontact mode can alleviate this problem, but
it comes at the cost of resolution [3]. Therefore,
methods described in this chapter relate to contact mode imaging of both fixed and viable cells.
In contact mode AFM imaging, specimen
immobilization will prevent movement during
scanning. There are various immobilization techniques, including physical trapping [4], electrostatic interaction by coating glass coverslips [5],
and chemical tethering of the cell to the substrate
[6]. Early studies by Kasas and Ikai [7] used
physical trapping to immobilize cells on filter
paper, while more recent studies have introduced
a micro-patterned surface for this purpose [8].
The latter methods are appropriate for spherical
cells such as certain bacteria or yeast. Most rodshaped bacteria imaged by AFM [9] have been
B.C. Paul et al.
immobilized through electrostatic interactions or
by culturing cells on chemically functionalized
solid surfaces. The different surface characteristics of conidia (hydrophobic) and hyphae (hydrophilic) make it difficult to use a chemical method
to adhere the entire fungal body to a solid support.
Thus, prior to developing a method for imaging
live filamentous fungal hyphae [4], the majority
of AFM studies had been restricted to spores or
yeast [10]. Zhao et al. [11] determined the elasticity of both hydrated and dehydrated Aspergillus
nidulans by electrostatically immobilizing fixed
cells on poly-l-lysine-coated coverslips; however,
imaging growing filamentous fungi requires the
continuous supply of media, which is not possible
with this method. Ma et al. [3] devised a method
to image live fungal hyphae by AFM. Dague et al.
[12] recently imaged the dynamic germination of
Aspergillus fumigatus spores immobilized on
filter paper by AFM and monitored the effect of
antimicrobial agents on germination, but their
work has not been extended to live hyphae. Here
we present sample preparation techniques for fungal spores, fixed hyphae, and live hyphae, and the
methods used for their AFM imaging and analysis
by force spectroscopy (FS).
Materials
1. Complete medium [13, 14].
2. Sterilized distilled water.
3. Ultrapure water (18 MW, Thermo
Scientific).
4. Ethanol (95%, Fisher Scientific).
5. Acetone (Fisher Scientific).
6. Hydrogen peroxide (Fisher Scientific).
7. Sulfuric acid (Fisher Scientific).
8. Octadecyltrichlorosilane ((CH3(CH2)17SiCl3)
NaNO3, Sigma-Aldrich).
9. Compressed air duster (Grand and Toy).
10. Incubator (Queue).
11. Kim wipes (Kimberly-Clark).
12. Filter paper (Whatman, USA).
13. Glass coverslips (22 mm × 22 mm, Fisher
Scientific).
14. Petri dish (100 mm × 15 mm, polystyrine,
disposable, sterile Petri dish, VWR
International).
9
High-Resolution Imaging and Force Spectroscopy of Fungal Hyphal Cells…
15. Dialysis tubing (Spectra/Pore® MWCO8000).
16. Triton-X-100 (Sigma-Aldrich).
17. Cantilevers (Ni3N4 AFM tip; spring constant:
0.05–0.5 nN/nm; tip radius: 10 nm; Bruker).
18. Standard sample (AFM Calibration Grating,
Bruker).
Experimental Methods
Here we describe the sample preparation and
microscopy methods developed for AFM imaging and force spectroscopy of fungal spores, and
germinating and mature fungal hyphae.
Preparation of Fungal Spores
for Imaging
Previous studies reported that the spore surface is
composed of hydrophobic rodlet (RodA) layers
composed of hydrophobin proteins [15], making
the cell surface hydrophobic [16]. Since the glass
substrate surface is hydrophilic, the hydrophobic
spore would not favorably adhere to the glass
coverslip. For this reason, the glass surface was
chemically modified with silanes to facilitate
spore adherence to coverslips through hydrophobic interactions. Physical confinement in a porous
membrane by filtering the spore suspension can
also be used to image spores by AFM [12], but
with the drawback of having the spores exposed
to water, which can lead to surface modification
through water imbibition and swelling [4]. Thus,
imaging spores under dry conditions is the only
way to examine its native ultrastructure and
physio-chemical surface properties. Steps
involved in silanization, all of which are conducted in a fumehood, are described in the following section.
Cleaning Coverslips
Fungal spores are approximately 1–3 mm in
diameter [17]; therefore, contaminants on the glass
coverslip surface will lead to imaging artifacts
and also can adversely affect the silanization
153
process required to adsorb spores onto the glass
surface. Prior to coating, coverslips must be
cleaned.
1. Dip coverslips in 1 M HCl for 2 min (see
Note 1).
2. Wash coverslips in deionized water and let
air dry.
3. Soak coverslips in Piranha solution (5 mL of
30% H2O2 + 15 mL of 18 M H2SO4) for 1 h
(see Note 2).
4. Remove the coverslip and wash with copious
amounts of deionized water.
5. Dip coverslip in methanol for 2 min and let
air dry.
6. Dip coverslip in acetone for 2 min and let air
dry.
7. Store the clean coverslips in a dust-free
container.
8. Clean coverslips can be sterilized with 70%
ethanol or by autoclaving.
9. Clean coverslips can be coated with either
poly-l-lysine or a silanization reagent.
10. Coated coverslips can be sterilized with 70%
ethanol (see Note 3).
Silanization
1. Immerse glass coverslips in a solution of 2%
octadecyltrichlorosilane (see Note 4).
2. Rinse coverslip in 100% hexanes (3×) with
gentle shaking, 1 min each.
3. Air dry over night in the fume hood.
Spore Sample Preparation
1. Touch the silanized coverslip to the conidiating culture (spores are produced after 3–4
days culture), so that a thin layer of conidia
adhere to the coverslip.
2. Incubate the sample for 1 h at room temperature, allowing spores to settle and form a
strong hydrophobic interaction with the
coverslip.
3. Remove any nonadhering spores or dust using
compressed air. Nonadhering spores on the
coverslip will make AFM imaging extremely
difficult.
The sample is now ready for AFM imaging in
contact mode.
154
Preparing Fixed and Dehydrated Fungal
Hyphae for Imaging
A. nidulans is a filamentous fungus that forms
mycelium when it grows for 2–3 days. The height
of the mycelium is beyond the limit of many
AFM Z-piezo ranges (10–100 mm). While it is
possible to view hyphae at low resolution by
CCD or light microscopy and visually line up the
AFM tip on the surface of structures that exceed
the AFM Z piezo limit [18], it is easier to probe a
sample by AFM with less height variability.
Therefore, two-dimensional hyphal samples were
grown for imaging by AFM, whereby hyphae
were grown for a short period of time between
two laterally aligned coverslips.
Formaldehyde Fixation
1. Prepare spore suspensions by pipetting 10 mL
of sterile water onto the conidiating culture.
2. Collect the water from the culture that now
contains spores and add it to 90 mL of sterile
water in an Eppendorf tube.
3. Mix the spore suspension by vortex.
4. Serially dilute the spore suspension to reduce
the spore count (~1/100 final dilution).
5. Add 20 mL of the spore suspension to 980 mL
of complete medium (CM) and mix thoroughly by vortex.
6. Pipette 200 mL of the spore suspension onto
a sterile, clean glass coverslip laying in a
sterile Petri dish.
7. Place a second sterile, clean glass coverslip on
top of the first one very carefully so that the
medium remains between the two coverslips.
8. Add 2 mL of liquid CM to the Petri dish area
surrounding the coverslips to avoid dehydration. Ensure that the additional 2 mL of liquid media added does not come into contact
with the coverslips.
9. Incubate the sample at 37 °C for 16 h.
10. Carefully remove the top coverslip and wick
off the remaining media from the bottom
coverslip using a Kimwipe, being careful not
to touch the mycelia.
11. Wash the bottom coverslip with 50 mM
warm (37 °C) phosphate buffer (dilute
B.C. Paul et al.
30.75 mL of 1 M K2HPO4 and 19.25 mL of
1 M KH2PO4 to 1 L with 18 MW water).
12. Treat with fixative solution (see Note 5)
(200 mL; 3.7% formaldehyde, 2% of
Triton-X-100, in 50 mM phosphate buffer,
pH 7.0) and incubate for 10 min.
13. Wash carefully (3×) with deionized water by
depositing 100 mL at a time on one corner of
the coverslip, allowing the solution to disperse, and wicking the solution from the
other side of the coverslip with a Kimwipe.
14. Dry the sample at room temperature in a covered Petri dish.
Fixation by OsO4
To compare AFM and SEM images, samples can
be fixed with OsO4 and then critical point dried
and gold-coated [3]. A detailed protocol is given
below.
1. Grow hyphae on coverslips following steps
1–10, described in section “Formaldehyde
Fixation.”
2. Attach the coverslip to the lid of a Petri dish
using double-sided tape and invert each plate
over approximate 200 mL of OsO4 (4% aqueous) in the Petri dish. Incubate for at least 2 h
at RT until the highly volatile OsO4 completely
penetrates the hyphae (agar is dark from Os
accumulation).
3. Freeze the fixed cells by plunging into cold
anhydrous acetone (60 mL, −80 °C) and dehydrate for at least 4 h.
4. Then warm the sample slowly to room temperature (2 h at −20 °C, 2 h at 4 °C, 2 h at RT)
to avoid condensation.
5. Dehydrate the sample (Emitech K850 Critical
Point Dryer; Quorum Technologies, UK). In
this method acetone is exchanged for liquid
CO2 (in a pressure chamber), which is heated
through its phase transition temperature,
becoming gas at the critical point.
6. The sample can then be imaged, or gold sputter coated (Emitech SC7620 Sputter Coater;
Quorum Technologies, UK) for direct comparison with traditional SEM data. Goldcoated samples are more robust, and so can be
imaged for several days after preparation.
9
High-Resolution Imaging and Force Spectroscopy of Fungal Hyphal Cells…
7. Mount the coverslip on the AFM sample
holder for imaging. Gold-coated samples, with
a harder surface, offer superior contact mode
AFM images.
Preparing Viable Fungal Hyphae
Many microbes have been imaged live by AFM;
however, viable filamentous fungi have not been
as well explored with this method. Many previous studies focus on unicellular, spherical yeast,
which are amenable to physical trapping for AFM
imaging and force spectroscopy [19–21]. Fixed
and rehydrated A. nidulans fungal hyphae and
spores were probed by FS and imaged [11, 22],
and the surface dynamics of A. fumigatus spores
have been probed in liquid medium during germination [16]. However, to date only our laboratory has studied the cell wall ultrastructure and
physical properties of viable hyphae, from spore
to germling [4, 12, 23, 24]. Below we describe
the steps used to prepare samples of live germlings and hyphae for AFM imaging.
1. Place a piece of sterile (boiled) dialysis tubing
membrane on a Petri dish containing agar
media.
2. Inoculate the membrane with the fungal spore
suspension (1 mL of 1/100 dilution). (Follow
step 1 of section “Spore Sample Preparation”
for spore suspension.)
3. Incubate at 37 °C for 16 h.
4. Following adequate hyphal growth, remove
the membrane from the Petri dish for transfer
to the AFM.
Imaging Spores and Fixed Hyphae
by AFM
1. After the 30-min warm-up, mount the sample
onto the AFM stage using double-sided tape
(see Note 6).
2. Locate the specimen visually with the CCD
camera or optical microscope (see Note 7).
3. Bring the AFM cantilever very close (~50 mm)
to the sample surface, but far enough away to
155
avoid crashing the tip into the surface and
breaking the cantilever.
4. For instruments with an optical lever design,
the laser must be aligned on the portion of the
cantilever directly above the AFM tip. Ideally,
this region would correspond to an optimal
signal (see Note 8).
5. Adjust the mirror between the laser and detector to give an optimum signal in the fourquadrant photodetector, which monitors the
position of the AFM cantilever and thus tip
response.
6. Many AFM instruments allow the sensor
response, which is a plot of distance vs. cantilever deflection, to be tested. For a properly
aligned laser and a new AFM cantilever, the
sensor response should be linear. If not, disengage the tip, readjust the laser and mirror, and
then reengage the tip. If it is not possible to
obtain a linear sensor response, it is likely that
the cantilever is old, and it is worth using a
new cantilever.
7. Use the line scan function in topography mode
to evaluate instrument feedback. Each time
the same line of the sample is scanned, the
topographic patterns should overlap. If not,
adjust the input gains, scan speed and other
parameters until this is achieved.
8. Once instrument feedback has been established, which is crucial for AFM imaging, a
large field of view (100 × 100 mm) can be
imaged at low resolution (200 × 200 pixels)
to identify spores or hyphae of interest (see
Note 9).
9. The zoom function allows a smaller field of
view (<500 nm × 500 nm) for imaging the surface of single spores or hyphae at higher resolution (up to 1,000 × 1,000 pixels).
These methods have been used to resolve 10 nm
hydrophobin rodlets on the native spore surface
(Fig. 9.1a) and 25 nm subunits on the hyphal surface of viable specimens (Fig. 9.1b) [4].
AFM Imaging of Live Hyphae
After growing hyphae on dialysis tubing membrane following the steps described in the section
156
B.C. Paul et al.
Fig. 9.1 Atomic force microscopy (AFM) images of
A. nidulans spore and fixed hyphal cell surface. (a) Highresolution AFM image (1.5 mm × 1.5 mm, 1,000 × 1,000
pixel) reveals the surface structure of a dry spore; and
inset is the image of the 10 nm-wide rodlets. (b) AFM
image (80 mm × 80 mm) of hyphae fixed with OsO4 and
critical point dried at low resolution (200 × 200 pixel). The
inset (1.5 mm × 1.5 mm, 1,000 × 1,000 pixel) shows the
ultrastructure of the same hyphal cell wall surface resolving 25 nm surface subunits. Images were adapted and
reprinted with permission from reference [4]
“Preparing Viable Fungal Hyphae,” the sample is
mounted on the AFM and imaged following the
steps described below:
1. Attach the dialysis tubing membrane to a glass
coverslip with double-sided tape.
2. Insert a small piece of filter paper into the
dialysis membrane tube (see Note 10).
3. Mount the coverslip onto the sample holder of
the AFM with double-sided tape.
4. Add media dropwise to the filter paper to
deliver media nutrients through the membrane to the hyphae during imaging (see
Note 11).
5. Make sure that the dialysis tubing remains
moist; otherwise it will dry out into a highly
convoluted structure that is too rough for AFM
imaging (Fig. 9.2) (see Note 12).
6. Follow the steps described in steps 2–8 of the
section “Imaging Spores and Fixed Hyphae
by AFM” for imaging live hyphae.
Force Spectroscopy of Live
and Fixed Hyphae
AFM is not only a tool for nanometer-scale
imaging, but can also be used for force spectroscopy, in which the tip approaches and retracts
from the sample surface. A force curve is generated by plotting cantilever deflection vs. tip distance from the surface in the Z direction, and this
is converted to force vs. distance using the spring
constant of the cantilever. Newer instruments
have combined the raster scanning and force
spectroscopy capabilities to produce force
maps. The same principles apply, but rather than
probing only a single point beneath the tip, force
curves can be collected over a large surface area
of the cell. When the AFM tip exerts a small
amount of force, the degree of cell indentation
will depend on the viscoelasticity of both the cell
and cantilever spring constant [25]. The slope of
9
High-Resolution Imaging and Force Spectroscopy of Fungal Hyphal Cells…
157
Fig. 9.2 Schematic of the assembly for AFM imaging of
hyphae on dialysis tubing. Dialysis tubing is fixed on a
coverslip with double-sided tape then placed on the AFM
stage. Media is delivered from underneath the dialysis
tubing through a piece of filter paper. More sophisticated
environmental chambers on new AFMs can be used for
media delivery
the approach curve into the cell surface in
conjunction with appropriate models can be used
to measure the spring constant of the cell, cell
wall, or envelope, while the last portion of the
retraction cycle reflects tip–sample adhesion. For
acquisition of force spectroscopic data for both
live and fixed hyphae in specific regions, it is
necessary to image the sample first using the
method described above, followed by the steps
given below:
1. Determine the spring constant of the cantilever based on its resonant frequency [26], or
calibrate the AFM cantilever using the method
described by Gibson et al. [27] to ensure accurate force data (see Note 13).
2. After mounting the sample on the AFM stage,
steps described in the section “Imaging Spores
and Fixed Hyphae by AFM” should serve as a
guide for imaging.
3. Switch to force spectroscopy or force mapping mode, allowing measurement at any
given point or across the entire surface, respectively (see Note 14).
4. For A. nidulans hyphal samples, the tip is initialized 1 mm from the sample surface.
5. Adjust the approach speed of the AFM tip during force curve acquisition, which will vary
with each AFM system. For force mapping,
the approach speed will determine the force
map acquisition time [28].
6. Image and acquire FS data for the sample surface, substrate surface, and a hard surface as a
reference.
Determination of Hyphal Viscoelasticity
Viscoelasticity can report on whole cell turgor
and cell wall integrity. The spring constant of
the hyphal cell, Kw can be determined from the
following equation using the slope, S, of the
approach force–distance curve (Fig. 9.3, section
b-c). A force curve from a hard surface is used to
determine the slope of the approach portion,
designated Sh.
K w = K c S / (Sh − S)
(9.1)
where Kc is the spring constant of the cantilever
used to acquire the force curve of the hyphal
sample determined according to step 1 in section “Force Spectroscopy of Live and Fixed
Hyphae.”
Once Kw has been calculated, the viscoelasticity of the hyphal cell wall can be calculated
according to the following equation [11]:
E = 0.80(K w / h)(R / h)1.5
(9.2)
158
B.C. Paul et al.
Fig. 9.3 Force curve from the live hyphal cell surface of
A. nidulans. Representative force vs. distance curve of
live A. nidulans hyphae. The slope of the b–c region can
be used to determine the mechanical properties of cell
using appropriate models. The e–f region describes adhesion between AFM tip and sample surface in pN, which
will depend on the chemical properties of AFM tip and
sample
where h is the cell-wall thickness measured by
TEM and R is the radius of hyphal cell measured
by AFM or TEM.
Such models hold for the conditions for
which they have been developed, in this case
hyphae with intact cell walls and relatively small
surface subunits. However, the model may no
longer fit data from hyphae with compromised
cell walls [13].
as a function of polarized hyphal growth [4] or
cell wall biosynthetic enzyme mutations [13].
Determining Surface Adhesion
The adhesion force between tip and sample
depends on the chemical nature of the tip and
sample surface. Adhesion force measurements
are useful to determine the relative hydrophobicity, hydrophilicity, or electrostatic properties of
the sample [29]. If chemically functionalized
tips are used, the force of binding between the
two molecules (FS) or the surface distribution of
a particular molecule (FM) can be determined.
The last portion of the retract curve is used to
determine the adhesion force in pN between the
sample and AFM tip as the distance between
points e and f of the force–distance curve (see
Fig. 9.3). Adhesion values can provide valuable
insight on carbohydrate remodeling, for instance
Notes
1. If preparing a large number of coverslips, a
home-built rack can be used for dipping,
since it is important to keep coverslips separate to ensure both proper cleaning and subsequent coating of the surface.
2. Caution: Piranha solution is extremely caustic and volatile. Work in a fume hood doublegloved.
3. Coated coverslips should not be autoclaved,
as the coating will degrade at high
temperatures.
4. Immersion of the coverslip in octadecyltrichlorosilane allows formation of covalent
bonds between silane and silicate groups of
the glass surface, resulting in the formation
of a self-assembled hydrocarbon monolayer
on the coverslip surface. Hence, the coverslip surface is transformed from being
hydrophilic to hydrophobic.
9
High-Resolution Imaging and Force Spectroscopy of Fungal Hyphal Cells…
5. Mix 10 mL of 37% formaldehyde and
0.2 mL of 1.7 M Triton-X-100 and dilute to
100 mL with 50 mM potassium phosphate
buffer.
6. For instruments with optical lever detection,
the laser will increase cantilever temperature
over time, so it is best to align the laser onto
the end of the cantilever to allow it to stabilize (~30 min). The piezoelectric scanners
perform best when warm (many labs who
use AFM for QC find images improve over
time, Asylum, personal communication), so
it is worthwhile to allow the AFM to raster
scan for at least 30 min with the tip out of
feedback prior to imaging.
7. It is very useful to first examine AFM samples by light microscopy to visualize the position of spores, germlings, hyphae, or mycelia.
Areas of interest can be marked underneath
the glass coverslip using a thin-tipped marker
to make them visible under the CCD camera.
It is very difficult to view spores with the
CCD camera (×200), so a large area of the
coverslip (100 × 100 mm) must first be imaged
at low resolution (200 × 200 pixels) to locate
spores. If the AFM is mounted on a light
microscope with ×40 or ×60 objectives, at
this magnification (×400–×600) it will be
possible to view the spores, eliminating the
need to image large areas at low resolution or
premark areas of interest on the coverslip
using a separate light microscope.
8. It can be difficult to position the laser on the
cantilever by viewing through a CCD camera.
One way to tell that the red laser is actually
hitting the surface of the cantilever is if a diffraction pattern appears below the cantilever.
9. Look for an area on the coverslip with limited three-dimensional growth and adequate
space between hyphae to allow access to a
single hypha. Make sure the mycelium is not
positioned beneath the free end of cantilever.
The AFM head can be tilted (front down,
back up) slightly to avoid the latter situation
which will ultimately prevent the tip from
engaging in feedback.
10. Cut the filter paper to a width slightly smaller
than that of the dialysis tubing, and to a
11.
12.
13.
14.
159
length slightly longer, such that part of the
filter paper remains outside the membrane
for media delivery.
During live imaging, 2 drops (~10 mL) of
media must be delivered to the sample every
30 min to keep the hyphae viable. Care
should be taken when delivering media, since
excess liquid on the membrane surface will
create extreme capillary forces that interfere
with AFM feedback. As a result, it can be
difficult to establish and maintain feedback
while imaging live samples, requiring AFM
imaging parameters to be adjusted more
frequently.
During live hyphal imaging, if the AFM tip
is pulled to the surface of the dialysis tubing
by capillary forces, feedback will be destabilized, so extra care should be taken during
media delivery. In some cases, reducing the
set point can also solve this problem.
Varying geometry and characteristics of
AFM probes may cause image distortion and
overestimation of the lateral size (X and Y) of
small features. To minimize experimental
error, AFM probes can be calibrated using an
AFM calibration grating with known dimensions, or nanometer-scale gold spheres [4,
30]. Accurate determination of physical
parameters such as hardness, friction or
adhesion requires an accurate cantilever
spring constant, which can be derived from
the spectrum of its thermal vibration in air.
Note that most modern instruments use a
mode called force mapping, which combines
force spectroscopy with raster scanning, creating a map of the sample surface describing
tip–sample interactions and sample compression. Force spectroscopy still remains a useful method for fast surface analysis.
References
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of Aspergillus nidulans studied with atomic force
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Dahms TE (2011) Quantifying the importance of
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Use of Fourier-Transform Infrared
(FTIR) Microscopy Method
for Detection of Phyto-Fungal
Pathogens
10
Vitaly Erukhimovitch and Mahmoud Huleihel
Abstract
Reliable and rapid identification of the fungal pathogens that cause plant
diseases is playing an important role in their control strategies. The available methods for identification of fungi are time-consuming and not always
very specific. Fourier-transform infrared microscopy is proving to be a
reliable and sensitive method for detection of molecular changes in cells.
Fungi pathogens display typical infrared spectra that differ from spectra of
substrate material such as potato, which make it is possible to detect and
identify such pathogens directly from the infected tissue.
In addition, although different strains of the same fungi species display
very similar infrared spectra, there are specific spectral differences between
them that might be successfully used, with the assistance of advanced statistical methods, for the identification of these fungal strains.
Keywords
Fungal pathogens • Fungal detection • Fungi • Fourier-transform infrared
microscopy • Spectral biomarkers • Potato
Introduction
V. Erukhimovitch
Ben-Gurion University of the Negev,
Analytical Equipment Unit, POB 653,
Beer-Sheva 84105, Israel
e-mail: evitaly@bgu.ac.il
M. Huleihel (*)
Department of Virology and Developmental Genetics,
Ben-Gurion University of the Negev, POB 653,
Beer-Sheva 84105, Israel
e-mail: mahmoudh@bgu.ac.il
Fungal pathogens are considered one of the most
common causes of severe diseases in various
plants. Infection with fungal pathogens can lead,
in many cases, to a great deal of economic damage [1]; consider, for instance, Colletotrichum
coccodes, a major pathogen of potato and tomato
[2]. Infected seed tubers are a major source of
contamination of field soils and storage areas. In
many cases, it is difficult to detect low levels of
contamination in the early stages of infection [3].
Early identification enables one to precisely target
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_10, © Springer Science+Business Media, LLC 2013
161
162
V. Erukhimovitch and M. Huleihel
a pathogen with the most effective treatment,
thereby preventing large economic damage. Most
commercially available identification systems are
based on the physiological and nutritional characteristics of fungi. Such identification systems are
usually time-consuming (2–4 weeks) and not
always very specific. Polymerase chain reaction
(PCR)-based methods developed for the detection
and identification of plant pathogenic fungi are
rapid and sensitive [4, 5]. Primers, designed to
conserve regions of the internal transcriber spacer
regions within ribosomal gene clusters, have been
used to detect and identify plant pathogenic fungi
[6]. Although this method is promising, it is not
yet in large-scale use and is expensive.
Fourier-transform infrared (FTIR) spectroscopy is considered valuable because of its sensitivity, rapidity, low expense, and simplicity. These
factors, together with the large information
already known about spectral peaks obtained
from FTIR spectra of living cells [7], make it an
attractive technique for detection of pathogens.
This technique was used for the detection and
characterization of cancer cells [8, 9], cells
infected with viruses [10], and microorganisms,
including some fungi [11–16].
Materials
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
Zinc sellenide crystals.
Sterile distilled water.
Vortex.
Water bath.
Incubator shaker.
Microcentrifuge.
Pipators.
Sterile tips (in different sizes) for the appropriate pipators.
Disposable polypropylene microcentrifuge
tubes (2 mL screw-capped).
Potato Dextrose Agar (Difco).
Potato Dextrose Broth medium (Difco).
Fungi: Different fungi strains were obtained
from ATCC for examination by FTIR
microscopy.
Infected tissues (such as potatoes) with
known strains of fungi.
Method
Procedures Used for Preparation
of Fungi Samples
FTIR microscopy method is applied in our studies for the detection and identification of different
fungal species and strains that are isolated and
purified from culture media or directly examined
on the infected tissues.
1. Fungi growing on solid media: The examined
fungi are cultivated for several days (3–6 days)
on Potato Dextrose Agar (PDA, Difco) at
27 °C. Small aliquots of fungi are picked up
from the growing fungi on the agar with a bacteriological loop, suspended in 500 mL of sterile distilled H2O, pelleted by centrifugation at
2,000 rpm for 5 min. Each pellet is washed
twice with sterile distilled H2O (by suspending it with 500 mL of sterile distilled H2O and
pelleting by centrifugation at 2,000 rpm for
5 min) and suspended with 50 mL of sterile
distilled H2O.
2. Fungi growing in liquid media: The examined
fungi are cultivated and identified using classical microbiological techniques [2]. Briefly,
samples of the fungi are cultivated and maintained in Potato Dextrose Broth media (Difco).
These cultures are grown for 3–10 days at
continuous shaking conditions and at a temperature of 27 °C (the growth time depends on
the fungi species).
3. Samples of these fungi are purified from these
media by spinning about 1 mL of medium
containing fungi at 2,000 rpm for 5 min, washing twice with sterile distilled H2O and the
pellet is suspended in appropriate volume
(about 50 mL) of sterile distilled H2O.
4. Direct examination of fungi from infected tissue: Small epidermal aliquots of samples are
scratched from the infected (or uninfected)
areas on the surface of tissue (such as potato
tubers), suspended in 500 mL of sterile distilled H2O, pelleted by centrifugation at
1,000 rpm for 5 min. Each pellet is washed
twice with H2O and re-suspended with 50 mL
of sterile distilled H2O.
10
Use of Fourier-Transform Infrared (FTIR) Microscopy Method for Detection…
163
Fig. 10.1 A representative photo of the FTIR microscopy instrument used in this study
Preparation of Slides for FTIR
Microscopy Examination
Since ordinary glass slides exhibit strong absorption in the wavelength range of interest to us, zinc
sellenide crystals, which are highly transparent to
IR radiation, are used. A drop of 5 mL of the
obtained suspension (containing the fungal sample as described above in steps 1–3) is placed on
the zinc sellenide crystal, air dried for 20–30 min
at room temperature (or for 5–10 min by air drying in a laminar flow) until all the water had evaporated, and then examined by FTIR microscopy.
FTIR Spectra Measurement
FTIR measurements are performed in transmission mode with a liquid nitrogen-cooled MCT
detector of FTIR microscope (Bruker IRScope
II) coupled to an FTIR spectrometer (Bruker
Equinox model 55/S, OPUS software) (Fig. 10.1).
Figure 10.2 demonstrates a principal scheme of
FTIR microscope. The spectra are obtained in the
wave number range of 600–4,000 cm−1 in the
mid-IR region. A spectrum is taken as an average
of 128 scans to increase the signal/noise ratio,
and the spectral resolution was at 4 cm−1 with
Backman Harris 4-Term adopization function.
Since the obtained samples are heterogeneous in
many cases, appropriate regions are chosen by
FTIR microscope out of different impurity (salts,
medium residuals, etc.). The optimal aperture
used in this study was 100 mm, since this aperture
gave the best signal/noise ratio. At lower apertures, the quality of the spectra is bad owing to
high levels of noise. In addition, at apertures
lower than 20 mm, there is diffraction of the IR
beam. Baseline correction and normalization are
obtained for all the spectra by OPUS software.
Baseline correction is performed by the rubber
band as follows: each spectrum is divided up into
ranges of equal size. In each range, the minimum
y-value is determined. The baseline is then created by connecting the minima with straight lines.
Starting from “below” a rubber band stretched
over this curve constituted the baseline. The baseline points that do not lie on the rubber band are
discarded. Normalization is performed by a vector method, as follows. The average y-value of
164
V. Erukhimovitch and M. Huleihel
Fig. 10.2 A representative diagram of the sample measurement by FTIR microscopy with different parameters used in
these measurements
the spectrum is first calculated. This average
value is then subtracted from the spectrum so that
the middle of the spectrum was pulled down to
y = 0. The sum of the squares of all the y-values is
then calculated, and the spectrum is divided by
the square root of this sum. The vector norm of
the resulting spectrum is 1. Peak positions are
determined using second derivation.
FTIR Spectra Analysis
The obtained spectra are analyzed for specific
regions which show distinct differences between
normal uninfected and infected tissues with fungi.
Also, similar analysis is done for the differentiation between different fungi strains.
For instance, Fig. 10.3 shows the FTIR spectra of both uninfected and infected samples
obtained from potato tubers. Although there is a
very high similarity between the spectra of the
infected and the uninfected control tissues,
some of the characteristic fungal bands appeared
in the spectra of the infected tissue samples (while
they are missing in the spectra of the control
uninfected tissue samples) as follows:
1. A significant band is found in all examined
infected samples at 1,545 cm−1 (see Fig. 10.3b),
but it is missing in the control uninfected
samples.
2. A peak at 1,405 cm−1 is found in infected
potato samples, but it is missing in control
uninfected samples (see Fig. 10.3c).
These markers seem to be very characteristics
of the fungal spectra compared to the spectra of
uninfected potato epidermal samples as shown
above (see Fig. 10.3).
Statistical Analysis
Cluster Analysis
The obtained spectral results of infected and control uninfected potato tissues are classified using
cluster analysis. Cluster analysis (CA) is an unsupervised technique that examines the interpoint
distances between all the samples and presents
that information in the form of a two-dimensional
10
Use of Fourier-Transform Infrared (FTIR) Microscopy Method for Detection…
Fig. 10.3 FTIR spectra
of purified fungi
(Colletotrichum) grown in
standard medium, control
uninfected potato samples,
and potato samples
infected with fungi
obtained by scratching
technique. The obtained
spectra were examined
in various regions:
(a) of 600–2,000 cm−1,
(b) 1,480–1,600 cm−1,
and (c) 1,340–1,460 cm−1
165
166
plot known as a dendrogram. These dendrograms
present the data from high-dimensional row
spaces in a form that facilitates the use of human
pattern recognition abilities. To generate the dendrogram, CA methods form clusters of samples
based on their nearness in row space. A common
approach is to initially treat every sample as a
cluster and to join the closest clusters together.
This process is repeated until only one cluster
remains. Cluster analysis was performed according to Ward’s algorithm by OPUS software.
V. Erukhimovitch and M. Huleihel
2.
3.
4.
Principal Component Analysis
Principal component analysis (PCA) is a standard
tool in modern data analysis [17, 18]. It is a common approach for the reduction of dimensionality. In the transformed space the data are
uncorrelated, but not statistically independent. It
is widely used in identification problems, with the
assumption that the most separable directions are
those with the highest variance. This is frequently
the case, but not always. It is easy to show a scenario with most separable direction with much
lower variance than the maximal variance one.
Basically, PCA is a mathematical algorithm
that reduces the dimension of the problem dealt
with. In other words, instead of using many variables, the variability in the data is described using
only a few principal components [19].
The first linear combination is called the first
principal component (PC1), and contains in
region III approximately 62.5% of the variance.
The second principal component (PC2) accounts
for most of the residual variance and is perpendicular to the first one. The subsequent principal
components obey the same rules. This method
allows the reduction of our spectra to 36 variables
in the lower and higher wave number regions that
account for almost 100% of the variance [20].
Notes
1. It is important to dry completely the examined
samples because water spectral band may
overlap important spectral bands of the tested
5.
6.
7.
sample. Therefore, air drying in a laminar
flow or with a fan might be helpful and
recommended.
The 5-mL drop of sample (fungi) should be
placed on the zinc sellenide crystal as a concentrated drop and then slightly spread on the
crystal.
When choosing by microscope the aperture of
the sample to be scanned, it is important to
choose an aperture with confluent fungal
cells.
Be careful not to choose possible contaminants such as salts, rather than fungi, for
scanning.
Fungi are usually constructed of vast numbers
of threads called hyphae, which are tangled
together, rather than arranged in an organized
structure. This morphology makes it difficult
to dissolve fungi in water or spread them on
surfaces. FTIR-microscopy measurements
require the smearing of the measured specimen on the crystal surface, thus presenting a
serious challenge to the fungi measurement.
The fungi were torn into small pieces and
mixed as evenly as possible within the distilled water before taking and placing the sample on the zinc sellenide crystals.
After placing the sample containing the fungi
on the zinc sellenide crystals, try to spread the
drop as carefully as possible in order to obtain
homogeneous, thin (about 20 mm) layers.
When the samples placed on zinc sellenide
crystal for FTIR microscopy examination are
obtained from fungi growing on solid or in liquid media, the optimal signal/noise ratio is
achieved using an aperture of 100 mm.
However, when the samples are scratched
from areas on the surface of tissue, part of
these pieces may be with a radius smaller than
100 mm. Therefore, in these cases only large
pieces (than 100 mm) should be selected for
examination with 100-mm aperture. If smaller
pieces are examined, it is necessary to decrease
the aperture size in order to get spectra only
from the examined sample without unrelated
around regions.
10
Use of Fourier-Transform Infrared (FTIR) Microscopy Method for Detection…
References
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New York
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Effect of Colletotrichum coccodes on potato yield,
tuber quality, and stem colonization during spring and
autumn. Plant Dis 83:561–565
3. Bang U (1986) Effects of planting potato tubers
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Diagnosis of Parasitic Fungi
in the Plankton: Technique
for Identifying and Counting
Infective Chytrids Using
Epifluorescence Microscopy
11
Télesphore Sime-Ngando, Serena Rasconi,
and Mélanie Gerphagnon
Abstract
Fungal epidemics, especially in the form of parasitic chytrids, are omnipresent in aquatic environments, infecting diverse organisms. Major target
hosts are algae, primarily diatoms, chlorophytes, and colonial or
filamentous cyanobacteria. Chytrids are also called “zoosporic” organisms
because their life cycle includes dispersal forms, that is, uniflagellate zoospores, and host-associated infective sporangia. They are considered relevant not only for the evolution of their hosts but also for the population
dynamics and successions of phytoplankton communities, thus representing an important ecologically driving force in the food web dynamics.
However, ecological knowledge of microscopic fungal parasites in aquatic
environments is weak, compared to terrestrial ecosystems. We propose a
routine protocol based on size fractionation of pelagic samples and the use
of the fluorochrome calcofluor white (which binds to b-1,3 and b-1,4 polysaccharides) for diagnosing, identifying, and counting chitinous fungal
parasites (i.e., the sporangia of chytrids). The protocol offers a valid
method for the quantitative ecology of chytrid epidemics in aquatic ecosystems and food web dynamics.
Keywords
Direct counting method • Fungi • Sporangia • Parasitism • Environmental
samples
Introduction
T. Sime-Ngando (*) • M. Gerphagnon
UMR CNRS 6023, Université Blaise Pascal,
Clermont II, 24 Avenue des Landais, BP 80026,
63171 Aubière Cedex, France
e-mail: telesphore.sime-ngando@univ-bpclermont.fr
S. Rasconi
Department of Biology, University of Oslo,
Blindernvn. 31, Oslo, 0371, Norway
Fungal infections are recurrent in aquatic ecosystems [1, 2]. The most described aquatic fungi in
freshwater ecosystems belong to Chytridyomycota
(or chytrids). Chytrids infect a wide variety of
hosts, including fishes, eggs, zooplankton, and
other aquatic fungi but especially phytoplankton.
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_11, © Springer Science+Business Media, LLC 2013
169
170
Typical phytoplankton hosts include prokaryotes
and eukaryotes, primarily large size diatoms and
filamentous species [3]. Associated chytrids are
external eucarpic parasites that produce a specialized rhizoidal system within host cells, that
is, the diet conveying system that leads to the formation of the chitinous fruit bodies: the sporangium. This parasitic stage produces numerous
uniflagellate spores, the zoospores, which constitute the dissemination phase of the life cycle [4].
Various approaches have been used to study
fungal parasites but routine techniques for reliably identifying and counting these organisms
are missing in the context of aquatic microbial
ecology [5, 6]. So far, observations of parasitic
fungi were obtained by using phase contrast light
microscopy with live or Lugol’s iodine preserved
samples [7–9]. Such conventional microscopy
allows observation of fungal sporangia or similar
forms (especially in laboratory cultures), but is a
poor approach for characterizing chytrid parasites in natural samples, at the complex community level. For example, a simple light microscopy
observation of fungal rhizoidal systems, that is, a
pertinent criterion for identifying chytrids [4, 10,
11], is very difficult. This situation may help
explain the confusion of chytrids with protistan
flagellates such as choanoflagellates or other bacterivorous flagellates in the group of Bicosoeca,
which are attached to phytoplankton but do not
harm their host [5–7].
Earlier studies on chytrids were restricted to
morphological descriptions and focused on few
species [12–15]. Electron microscopy was used to
describe different life stages and the ultrastructural cytology of fungal zoospores and spore differentiation [16–18], providing the basis for
chytrid taxonomy [19, 20]. Studies on pelagic
chytrids started in the British lakes [21], and different authors have provided descriptions of morphological characters [22–26]. However, few
attempts have been made to include the related
parasitism pathway in the aquatic food web
dynamics, and to understand environmental factors that trigger epidemics as well [27]. Some
authors have also investigated the effects of parasitism on the growth of algal host species and on
the genetic structure of infected populations [28].
T. Sime-Ngando et al.
Parasites are thus considered relevant not only for
the evolution of their hosts but also for the population dynamics such as successions of phytoplankton communities, and for structuring microbial
communities in general [9, 29]. Moreover, chytrids
can represent interesting key intermediates in the
food chain [30, 31]. The nutrients from infected
large-size algae that could not be fed directly by
zooplankton can be transferred from sporangium
to grazers via fungal zoospore production. Fungal
zoospores have suitable dimensions and represent
a valuable food source for zooplankton [32]. The
activity of zoosporic fungi and the related biogeochemical processes can thus be crucial in matter
and energy transfer in aquatic systems [29].
Methodological limitations for the study of the
ecological dynamics of chytrid populations can be
overcome with epifluorescence microscopy coupled to a specific fluorochrome targeting molecular tracers (i.e., some types of polysaccharides) of
the fungal chitinous structures, including sporangium and the rhizoidal system.
The protein stain fluorescein isothiocyanate
(FITC) and, in particular, the chitin stain
calcolfluor white (CFW), were suggested as good
markers that offer useful tools for the investigation of fungal dynamics in aquatic samples [33].
CFW binds to b-1,3 and b-1,4 polysaccharides
such as those found in cellulose or in chitin,
which commonly occur in the fungal cell wall [1,
2]. It fluoresces when exposed to UV light and is
currently used in clinical mycology for direct
microscopic examination of skin scrapings, hairs,
nails, and other clinical specimen for fungal elements [34, 35]. In contrast to FITC, CFW penetrates into infected host cells and is more efficient
for the observation of the complete rhizoidal system of parasites, that is, a pertinent criterion for
chytrid identification [4, 10, 11].
The main objective of this chapter is to provide, in a simplified step-by-step format, a routine protocol based on size-fractionation of
pelagic samples and the use of the fluorochrome
calcofluor white for diagnosing, identifying, and
counting fungal parasites (i.e., sporangia of
chytrids) within phytoplanktonic communities
[3], together with practical advices on how to
apply the method.
11
Diagnosis of Parasitic Fungi in the Plankton…
Materials
1. 25-mm nylon filter.
2. 0.2-mm filters.
3. High-performance concentration/diafiltration
system. As an example, we use the system
Amicon model DC 10LA (Epernon, France)
equipped with a reusable hollow fiber cartridge
(0.2 mm cutoff, surface area of 0.45 m2).
4. 36% Formaldehyde.
5. Calcofluor white (C40H44N12O10S2 Fluorescent
Brightener 28; Sigma catalog no. F3543).
6. 10 N NaOH.
7. Balance.
8. Distilled water.
9. 15 and 0.2-mL tubes.
10. Glass slides and coverslips.
11. Epifluorescence microscope equipped with
appropriate UV filter sets and Neofluar objective lens (optional).
Methods
Concentrations of Cells (See Note 1)
1. Pass the sample (ca. 20 L) through the 25-mm
pore size nylon filter (see Note 2).
2. Collect large phytoplankton cells in the
>25 mm size fraction by washing the filter with
40 mL of 0.2 mm filtered lake water.
3. Fix the concentrate sample with formaldehyde
(2% final conc.), before staining and analysis.
4. Concentrate nanoplanktonic cells in the
<25 mm size fraction (i.e., the 20 L filtrate) ca
20× by ultrafiltration to a volume of approximately 1 L, entry pressure 0.9 bar.
5. Fix about 180 mL of the ultrafiltrate retentate
with formaldehyde (2% final conc.), before
staining and analysis.
Preparation of Calcofluor Stock
Solution
1. Weigh 35 mg of Calcofluor White into a
15-mL tube.
171
2. Add 7 mL of sterile distilled water and 2–3
drops of 10 N NaOH (to increase pH to 10–11).
Calcofluor does not dissolve well in neutral
solutions.
3. Dissolve the calcofluor.
4. Adjust the volume to 10 mL by adding sterile
distilled water.
5. Distribute the stock solution in 0.2-mL tubes
and store in a light-proof tube at −20 °C.
Staining and Visualization
1. In the dark, stain aliquots (about 200 mL) of
concentrated and fixed materials by adding
1–2.5% (vol/vol) of CFW stock solution
directly in solution for 10 min.
2. Mount drop (5–10 mL) of the stained samples
between glass slides and coverslips for observations and counting.
3. In a dark room, examine the slides under an
epifluorescence microscope equipped with an
appropriate set of filters and objective lens.
Shift between white and UV light to visualize
and determine parasites and phytoplankton
cells, and check the viability of the host cell,
e.g., presence of chloroplast.
4. Applied a standard procedure for microscopic
counting (see Notes 3 and 4).
Notes
1. Different approaches were tested to concentrate samples: the total community approach
and the size-fractionated community approach.
For the former approach, 180 mL of experimental samples were fixed with formaldehyde
(2% final conc.) and aliquots were concentrated in three different ways: (1) by simple
gravity, following Utermöhl’s [36] method
before staining the chyrids; (2) by vacuum
pressure on two different filters before staining directly onto filters; and (3) by vacuum
pressure on the same two types of filters but
after staining in solution.
For the Utermöhl method, 100 mL of fixed
samples were settled for at least 24 h. For each
172
of the two filter-vacuum pressure methods,
10 mL × 2 of fixed samples were filtered onto
polycarbonate white filters (pore size 0.6 mm,
catalog no. DTTP02500, Millipore) and nuclepore polycarbonate black filters (pore size
0.8 mm, catalog no. 110659, Whatman), by
using gentle vacuum (<0.2 bar or 20 kPa). For
the total community approach using the classical Utermöhl [36] method, visualization of
fungal parasites was very difficult and most of
the time practically impossible for all the stain
concentrations tested. The main reason was
that staining directly in the Utermöhl chamber
resulted in very poor-quality specimens of
parasites observed in any given sample. Other
disadvantages of the procedure include the
relatively long sedimentation time and the
difficulty of increasing the volume analyzed.
The alternative total community approaches
based on vacuum pressure concentrations on
polycarbonate filters, that is, white (0.6-mmpore-size) and black (0.8-mm-pore-size) filters,
yielded similar quality images of fungal parasites, either when CFW staining was done
before (i.e., in solution) or after (i.e., on filters)
concentrating phytoplankton host cells onto
filters. However, substantial differences were
noted depending both on the type of the filter
and on the concentration of the stain. In general, for the two types of filters, high levels of
background noises were obtained when using
CFW at final concentrations of 3, 10, or 20%,
precluding any accurate assessment of numerical and phenotypic characteristics of both host
cells and their fungal parasites. Staining with
1% CFW final concentration substantially
improved the viewing of chytrids on filters,
with an increasing contrast from the white
DTTP Millipore to the black Whatman filters.
However, none of the membrane-retaining
approaches yielded satisfactory images of morphological and cellular features of the host
cells (e.g., presence of chloroplast, viability of
the host cell). Accordingly, the proposed protocol is based on the size-fractionation approach
using 1–2.5% vol/vol CFW final concentration
(from the stock solution), which substantially
enhanced the observational results.
T. Sime-Ngando et al.
2. The approach is efficient since it is based on
the concentration of large initial volumes and
size-partitioning of samples, a step that we
judged necessary in order to yield good analytic images of infectious sporangia for accurate diagnosis and identification of parasites.
In addition, this approach yielded satisfactory
images of morphological and cellular features
of the host cells, for phytoplankton
identification based on phenotypic features
and viability of the host cell, through the integrity of cell wall and the presence of chloroplasts, which are fundamental parameters to
assess the intensity of the disease. We consider
this protocol optimal for the diagnosis and
quantitative assessment of phytoplanktonic
chytrid infections in natural samples. Finally,
the approach was designed to freeze-conserve
particulate DNA samples for quantifying the
propagule stages (i.e., zoospores) of chytrids
via FISH targeting of specific rRNA oligonucleotide probes (see Chap. 5).
3. To estimate the infectivity parameters of ecological interest in phytoplankton population,
several algorithms are used according to formula proposed by Bush et al. [37] These
parameters include the prevalence of infection
(Pr), that is, the proportion of individuals in a
given phytoplankton population having one or
more sporangia or rhizoids, expressed as Pr
(%) = [(Ni/N) × 100], where Ni is the number
of infected host cells and N the total number
of host cells. The second parameter is the
mean intensity of infection (I), calculated as
I = Np/Ni, where Np is the number of parasites
and Ni is the number of the infected individuals within a host population.
4. We propose a third parameter concerning the
prevalence of infection of cells in colonial (or
filamentous)
species
(PrCF).
PrCF
(%) = [(Ni/N) × 100], where Ni is the number
of infected host cells in parasitized colonies
(or filaments) and N the total number of parasitized host colonies (or filaments).
Acknowledgements SR and MG were supported by
PhD Fellowships from the French Ministère de la
Recherche et de la Technologie (MRT). This study
11
Diagnosis of Parasitic Fungi in the Plankton…
receives grant-aided support from the French ANR
Programme Blanc # ANR 07 BLAN 0370 titled DREP:
Diversity and Roles of Eumycetes in the Pelagos.
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Fungal Cell Wall Analysis
12
Pilar Pérez and Juan C. Ribas
Abstract
Fungal cell wall is a rigid structure mainly composed of polysaccharides
(up to 90 %) and glycoproteins. It is essential for survival of the fungal
cells, because it protects them against bursting caused by internal turgor
pressure and against mechanical injury. Because of its absence in mammalian cells, it is an attractive target for antifungal agents. Thus, for various reasons, it might be important to know how the cell wall is synthesized,
and how to analyze its composition. We provide here information about
in vitro analysis of the biosynthetic activities of the main fungal wall and
describe some methods for rapid analysis of cell wall composition by
using specific enzymatic degradations. We also describe some additional
methods that can be occasionally used to analyze fungal wall properties or
composition. These methods provide powerful tools to evaluate changes
in fungal cell walls and will be useful for screening new compounds for
antifungal activity that might cause inhibition of cell wall biosynthesis
and/or alter the structure of the fungal cell wall.
Keywords
Cell wall • Polysaccharides • Glucan • Chitin • Mannan • Antifungal
drugs
Introduction
P. Pérez (*) • J.C. Ribas
Instituto de Biología Funcional y Genómica (IBFG)
CSIC, Universidad de SalamancaC, Zacarías González
s/n 37007, Salamanca, Spain
e-mail: piper@usal.es
The fungal wall is responsible for the cell shape,
provides mechanical protection, and supports
the internal osmotic pressure of fungal cells. In
addition, it acts as a filter for large molecules,
and its rigid structure is useful for penetration
into and colonization of insoluble substrates.
The cell wall is also the surface of interaction
between pathogenic fungi and their host. Indeed
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_12, © Springer Science+Business Media, LLC 2013
175
176
P. Pérez and J.C. Ribas
Fig. 12.1 Structure and composition of fungal cell wall.
Transmission electron micrograph (TEM) of a fission
yeast cell. A TEM detail of the cell wall is presented in the
lower panel with a scheme of the organization and com-
position of the two main cell wall layers—electron dense
and electron transparent layers—and the inner plasma
membrane-bound glycoproteins
the host defense response is usually directed
against the cell wall. This structure is not simply
a rigid exoskeleton but has the elasticity necessary to permit morphological changes during
fungal growth and life cycle.
To build the walls, fungal cells need to synthesize wall components, export them across the
plasma membrane, and assemble them outside the
cell. The wall is composed basically of polysaccharides (70–90%) and glycoproteins (10–30%).
Although composition varies among fungal species, and may even vary within a single fungal
isolate, depending upon the growth conditions,
most walls have a common structure [1]. When
observed by transmission electron microscopy
(TEM) the cell walls show a dark external layer
formed by glycoproteins and an internal layer
more transparent to the electrons, which mainly
contains fibrillar polysaccharides (Fig. 12.1). The
major fungal wall fibrillar components are: glu-
cose homopolymers, b(1,3)-d-glucan with some
b(1,6) branches, that constitutes 48–54% of total
cell wall polysaccharides; chitin, a b(1,4)-Nacetylglucosamine
polymer;
and
a(1,3)
(1,4)-d-glucan. Chitin accounts for only 1–2% of
yeasts wall [2, 3], whereas filamentous fungi, such
as Neurospora or Aspergillus, contain 10–20%
chitin in their walls [1]. In both yeasts and
filamentous fungi, chitin forms microfibrils from
interchain hydrogen bonding that have enormous
tensile strength and significantly contribute to the
overall integrity of the cell wall [4].
The wall polysaccharides are formed at the
plasma membrane by synthase enzymes and
extruded into the periplasmic space where they
bind to each other [5–7]. The linkages among the
different components, which results in a tightly
linked network, are generated by transglycosylation [8, 9] and are responsible for the mechanical
strength of the cell wall [5–7, 10].
12
Fungal Cell Wall Analysis
The formation and remodeling of the cell wall
involves several biosynthetic pathways and the
concerted actions of numerous gene products
within the fungal cell. Many of the genes involved
in cell wall synthesis or regulation have been
cloned by complementation of mutants altered in
wall structure or defective in the biosynthesis of
cell wall components. Those mutants were isolated in many different ways, reflecting the complexity of functions involved in cell wall integrity
and cell viability. Moreover, many of the genes
and enzymes critical for assembly and biogenesis
of fungal walls remain unidentified or poorly
characterized. The main studies on fungal wall
composition and biosynthesis have been performed in Saccharomyces cerevisiae [3, 11] but
can be extended to other fungi.
Cell Wall Components
Glucan
Glucan is the main structural polysaccharide of
the wall, and it represents 50–60% of this structure’s dry weight. The majority of glucan polymers are composed of glucose units with b(1,3)
bonds (65–90%), although there are also some
b(1,6), b(1,3)(1,4) and b(1,4) glucans. Usually
the main backbone is b(1,3)-d-glucan with b(1,6)
branches (Fig. 12.2). The b(1,3)-d-glucan is synthesized by a complex of enzymes known as glucan synthases located in the plasma membrane.
These enzymes catalyze the formation of linear
glucan chains composed of, approximately, 1,500
b(1,3)-bound glucose residues. In these linear
chains, new glucose units bind, forming b(1,6)
branches in variable proportion depending on the
organism—from almost linear to highly
branched—which can bind to other glucans, to
chitin or to glycoproteins, providing a great
mechanical resistance to the wall, which is essential to maintain the fungal cell integrity (see
Fig. 12.2).
The genes coding for the putative b(1,3)-dglucan synthase catalytic subunit were initially
identified in S. cerevisiae and named FKS1 and
FKS2 [12, 13]. The Fks protein family of b(1,
177
3)-d-glucan synthase is very well conserved in
fungi and plants. Orthologs of these genes have
been described in the main fungal genera such as
Schizosaccharomyces, Candida, Aspergillus,
Cryptococcus or Pneumocystis [1, 3, 14]. Besides
the catalytic subunit, fungal glucan synthases
(GS) require GTP-bound Rho1 GTPase for
their activity [15, 16]. This family of enzymes
use uridine-diphospho-glucose (UDP-Glc) as
substrate and catalyze the reaction 2 UDPGlc → [Glc-b-1,3-Glc].
A second b-linked glucan contained in most
fungal walls is the b(1,6)-glucan. This polymer is
shorter than b(1,3)-glucan, it does not form a
fibrillar structure, and acts as a flexible glue by
forming covalent cross-links to b(1,3)-glucan,
chitin, and glycoproteins [6].
Some fungi contain a(1,3)(1,4)-glucan in their
cell wall. However, the corresponding in vitro
a(1,3)-glucan synthase activity has not been
described yet. A putative catalytic subunit was
first described in Schizosaccharomyces pombe
[17, 18]. Ags1/Mok1 is a multidomain integral
membrane protein with a predicted domain highly
similar to starch synthase in the inner side and
another domain similar to a-amylase and other
proteins implicated in glycogen metabolism in
the outer side. S. pombe contains five genes coding Ags/Mok proteins, and genomes of other
fungi, including several human fungal pathogens
in which cell wall a-glucan accounts for around
35% of the total wall polysaccharides, contain
sequences of predicted proteins homologous to
these genes [19, 20].
Chitin
Chitin is a b(1,4)-linked homopolymer of
N-acetylglucosamine present in the cell walls of
all fungi studied to date with the exception of S.
pombe. Chitin represents 1–2% of the dry weight
of the yeast cell wall whereas in the filamentous
fungi it can reach up to 10–20% [1]. Chitin is
synthesized from N-acetylglucosamine units by
the enzyme chitin synthase (CS) that deposits
microfibrils of chitin outside of the plasma
membrane. This family of enzymes use uridine-d
178
P. Pérez and J.C. Ribas
Fig. 12.2 Schematic representation of the synthesis and
organization of the b-glucans forming the central core of
the fungal cell wall. Linear b(1,3)-glucan chains are initially synthesized in the plasma membrane by the glucan
synthase. Then cell wall transglycosidases form b(1,6)branched b(1,3)-glucan that is also linked to b(1,6)-glucan
through the side chains and to chitin via b(1,4) linkages.
Proteins are covalently attached to b(1,6)-glucans (GPICWP) through the GPI remnant or to b(1,3)-glucans (PIRCWP) through a glutamine residue following a
transglutaminase reaction
iphosphate-N-acetylglucosamine (UDP-GlcNAc)
as substrate and catalyze the reaction 2 UDPGlcNAc → [GlcNAc-b-1,4-GlcNAc]. Chitin biosynthesis has been mainly studied in S. cerevisiae,
which has three chitin synthases (CS1–3) responsible for the synthesis of chitin [21] at different
times and places during cell growth. The number
of chitin synthase genes varies from 1 to 20
according to the fungal species. The large family
of chitin synthase (CS) enzymes fall into seven
classes according to the evolution of their amino
acid sequences [22]. The multiplicity of enzymes
suggests that they have redundant roles in chitin
synthesis and makes it difficult to find functional
significance to the different classes [23].
Glycoproteins
Glycoproteins represent 30–50% of the dry weight
of the S. cerevisiae or Candida walls, and around
20% of the dry weight of S. pombe and the
filamentous fungi walls. These wall proteins have
diverse functions, participating in the maintenance
of the cellular form, taking part in adhesion processes, transmitting signals to cytoplasm, and
remodeling the components of the wall. The glycoproteins present in the cell wall are extensively
modified with both N- and O-linked carbohydrates,
predominantly or exclusively formed by mannose
residues known as mannan. In some cases, the
mannan backbone presents single residues or side
12
Fungal Cell Wall Analysis
chains of different sugars, galactomannan, rhamnomannan, glucogalactomannan, rhamnogalactomannan, etc. [1, 24, 25]. Most cell wall proteins are
attached through a glycosylphophatidyl inositol
(GPI) remnant to b(1,3)-glucan or chitin, via a
branched b(1,6)-glucan linker [11].
179
3.
Analysis of the Cell Wall Synthases
We will present here the methods described for
in vitro measurements of the enzymatic activities
responsible for the biosynthesis of the two main
structural wall polysaccharides: b(1,3)-glucan
synthase (GS) and chitin synthase (CS). Both are
integral membrane proteins, localized in the
plasma membrane with their catalytic sites facing
the inner side of the membrane.
4.
5.
b(1,3)-Glucan Synthase (GS)
The original method to detect the in vitro b(1,3)glucan synthase activity was described nearly
three decades ago [4, 26, 27]. This method has
been modified, simplified and improved but
essentially the basis of the protocol remains unaltered [28, 29].
Membrane Extracts Preparation
The source of enzyme activity is a crude membrane extract partially purified from the total cell
extract.
1. Cell cultures (100 mL) are collected at early
log-phase (A600 0.7–1.0) and centrifuged 5 min
at 4 °C, 3,000 × g (5,000 rpm in a GSA-type
rotor).
2. Cells are suspended in 30–40 mL cold buffer
A (50 mM Tris–HCl pH 7.5, 1 mM EDTA,
and 1 mM b-mercaptoethanol, centrifuged
5 min at 4 °C, 3,000 × g (6,000 rpm in a SS34type rotor), transferred to a 1.5-mL tube (with
screw cap), washed with 1 mL cold buffer A
(1 min at 16,000 × g, 13,200 rpm) and resuspended in 100 mL cold buffer A containing
50 mM GTPgS (GTPgS is more stable than
GTP, not hydrolysable and therefore, better
GS activator). Glucan synthase is very labile
6.
and GTPgS is very useful to preserve the
enzyme activity.
Cells are broken with glass beads (0.5 mm
diameter, filling in all the liquid with glass
beads and discarding by gentle drop out the
excess of beads not entrapped by liquid capillarity) in a FastPrep apparatus (Q-Biogene,
MP Biomedicals, Thermo Scientific) during
15 s at a speed of 6.0 and at 4 °C if possible.
Alternatively, the cells can be broken in glass
tubes with glass beads by 6 or 7 cycles of vortexing for 30 s and cooling down in ice for
another 30 s. Due to enzyme instability, the
rest of process must be done at 4 °C and the
sample must be kept on ice.
Broken material and glass beads are diluted
with 30 mL buffer A. Beads and cell debris are
removed by low speed centrifugation, 5 min at
4 °C and 3,000 × g.
The supernatant is then centrifuged at 36,000–
38,000 × g for 30 min at 4 °C and the membrane pellet is resuspended carefully by using
a glass stick and a vortex to extend the membrane material throughout the entire bottom
surface of the tube. Then 25 mL of buffer A
containing 33% glycerol and 50 mM GTPgS is
added and the membranes are homogenized
by vortexing with the glass stick throughout
the tube surface. The process is repeated with
another 25, 50, and 50 mL of the same buffer
until the membranes are homogenized in 100–
150 mL and stored at −80 °C.
A homogeneous emulsion of membrane
extract is critical for a reproducible GS assay.
The amount of protein is quantified by using
the Bradford dye-binding assay (Bio-Rad)
with bovine serum albumin as standard. The
protein concentration of the enzyme extract is
usually kept at 3–5 mg/mL.
GS Assay
The GS mixture contains 5 mM UDP-[14C]glucose (200 cpm/nmol) (PerkinElmer), 150 mM
GTP or GTPgS, 0.75% bovine serum albumin,
2 mM EDTA, 75 mM Tris–HCl pH 8.0, and 5 mL
of enzyme extract (15–25 mg of protein) in a total
volume of 40 mL. The correct amount of protein in
the assay is critical: higher protein concentration
180
does not result in proportional increase of GS
activity; therefore, the relative GS activity per milligram of protein decreases.
The reaction mixture is incubated for
30–60 min at 30 °C and stopped by addition of
1 mL of 10% trichloroacetic acid (TCA). The
samples are kept at least 30 min at 4 °C, filtered in
Whatman GF/C glass fiber filters, and washed
three times with 1 mL of 10% TCA and twice
with 1 mL of ethanol. The filters are placed into
vials, 2 mL of liquid scintillation is added and the
radioactivity of the filters is measured in a
Beckman scintillation counter. One unit of GS
activity is the amount of enzyme that catalyzes the
incorporation of 1 mmol of glucose into glucan
per min at 30 °C. The specific activity is expressed
as milliunits per mg of protein and the reactions
are always performed in duplicate. The GS
enzyme is very labile and therefore, the data for
each GS assay must be reproducible and calculated from at least three to four independent
experiments.
When the membrane extract is obtained in the
absence of GTPgS, we can measure basal and
maximal GS activity by omitting or adding GTP
to the reaction mixture. Alternatively, the GS
assay may be done at pH 7.0–7.5 and may contain 25–30 mM potassium or sodium fluoride [4,
12, 30–37] and 0.5% Brij-35 [38, 39].
The detergents 2% Tergitol NP-40 (with 2 M
NaCl) or 1.0% CHAPS are used for GS fractionation and solubilization of the regulatory subunit
[12, 15, 35, 40–43], and 0.5% CHAPS, 0.1%
cholesteryl hemisuccinate, or 0.1–0.2% CHAPS,
0.5–1.12% octyl glucoside are used for partial
solubilization of GS microsomal fractions [30,
32, 34, 35, 44].
The GS reaction product can be confirmed to
be b(1,3)-glucan, not a contaminant product such
as glycogen, by degradation with Zymolyase100T (AMS Biotechnology) or Kitalase (Wako
Pure Chemical Industries). Zymolyase-100T
is a preparation partially purified by affinity chromatography from Arthrobacter luteus that
contains b(1,3)-glucanase, protease, and
mannanase activities, but it does not contain
b(1,6)-glucanase and a-glucanase activities.
Similarly, Kitalase is a preparation from
P. Pérez and J.C. Ribas
Rhizoctonia solani with b(1,3)-glucanase,
protease, hemicellulase, pectinase, and amylase
activities, but it does not contain a(1,3)-glucanase activity. Kitalase is also named as Lysing
enzymes from Rhizoctonia solani (SigmaAldrich) or as Yeast Lytic Enzyme from
Rhizoctonia solani (MP Biomedicals).
The degradation mixture contains the GS reaction product (40 mL), either 20 mg Zymolyase100T, 50 mM citrate-phosphate pH 5.6 or 25 mg
Kitalase, 50 mM potassium-acetate pH 5.0, and
either 0.2% Tween 20 or Triton X-100 (1% detergent produces lower degradation) in a volume of
300 mL. The mixture is incubated 15–24 h at
30 °C with shaking, stopped with 10% TCA and
processed as a standard GS assay. Zymolyase or
Kitalase degradation in the absence of detergent
is not complete, with a 15–20% residual product,
likely due to the protection conferred by membrane vesicles. Other enzyme complexes result in
only partial degradation of the reaction product.
Microtiter-Based Fluorescence Assay
This method has been described as an alternative
to the use of UDP-[14C]-glucose for the GS assay
[39]. This method takes advantage of substituting
radioactive substrate for the fluorochrome aniline
blue that is specific for linear b(1,3)glucan.
Aniline binding is proportional to the amount of
linear glucan, and it can be measured in a
microplate fluorescence reader (excitation at
400 nm, emission at 460 nm). The GS mixture
(50 mL total volume, 100 mg of enzyme protein) is
similar to that described above except that UDP[14C]-glucose is omitted and it may contain 0.5%
Brij-35. The reactions are performed in microtiter
plate wells at 30 °C for 30–60 min and stopped
with 10 mL of 6 N NaOH. The glucan product is
solubilized by heating at 80 °C for 30 min followed by the addition of 210 mL of aniline blue
mix (1 mL contains 400 mL of 0.1% aniline blue,
210 mL of 1 N HCl, and 590 mL of 1 M glycine/
NaOH pH 9.5). The plate is incubated at 50 °C for
30 min and at room temperature for another
30 min to allow reaction with the fluorochrome.
Then, the fluorescence is quantified in a
fluorescence reader. Linear b(1,3)-glucans (such
as pachyman, curdlan, or yeast glucan dissolved
12
Fungal Cell Wall Analysis
in 1 N NaOH at 80 °C for 30 min) are used as
standards, in a reaction mixture containing the
same components except the membrane extract.
In Situ GS Assay Using Permeabilized
Whole Cells
This method takes advantage of being a more
direct enzymatic assay, omitting the steps of
membrane extract preparation. In addition, this
method can be applied for GS, chitin synthase, or
other membrane-bound or cytosolic enzymes.
The enzyme activity in permeabilized cells can
yield similar or higher activities than those in cell
extracts [45, 46]. The procedure is as follows:
1. Early log-phase cells are collected by centrifugation (1,500 × g, 5 min) and suspended in
40 mM EDTA, 100 mM b-mercaptoethanol
(3.5 mL per g of cells wet weight).
2. The cells are incubated at 30 °C for 30 min
with shaking, collected by centrifugation at
3,000 × g for 5 min, washed with 5 mL of
1.2 M sorbitol, resuspended (7 mL per g of
cells) in 50 mM citrate phosphate pH 6.3,
1 mM EDTA, 1.2 M sorbitol, and incubated at
30 °C for another 30 min with shaking. Similar
result can be obtained keeping the cells on ice
for 30 min without shaking. Then, the cells are
centrifuged at 3,000 × g for 5 min, suspended
in 30 mL of cold 50 mM Tris–HCl pH 7.5 for
osmotic shock, kept on ice for 5 min and centrifuged at 13,000 × g for 5 min.
3. The cells are suspended (1–1.5 mL per g of
cells) in the buffer of the reaction assay of
choice (50 mM Tris–HCl pH 7.5, 33% glycerol for GS and CS assays) and stored at
−80 °C for weeks. Under those conditions,
more that 90% of the cells are permeabilized
(measured by staining with methylene blue).
Alternatively, the cells can be collected, washed
with ice-cold water, suspended in cold 50 mM
Tris–HCl pH 7.5, 1 mM EGTA, 1 mM b-mercaptoethanol, 0.5 mM phenylmethylsulfonyl fluoride
(PMSF), 33% glycerol, and kept for 10 min in ice.
3.5 M glycerol can be replaced by 1.2 M glycerol,
sorbitol, mannitol, or 1 M KCl. The cells are
washed and suspended (1 g of cells wet weight
per mL) in cold buffer without glycerol or the
corresponding osmolyte [38]. In another protocol
181
the cells are collected, washed with 50 mM
Tris–HCl pH 7.5, 1 mM EDTA, 1 mM DTT, 33%
glycerol, resuspended in the same buffer and permeabilized with 2% toluene/methanol (1.1) at
22 °C for 5 min. The cells are washed twice at
4 °C with cold buffer and resuspended in cold
buffer (1 g of cells wet weight per mL) [47].
The GS assay contains the same mixture than
that of the membrane assay except that the 5 mL
of enzyme extract is replaced by 5 mL of permeabilized cells (5 mg of cells wet weight), in a total
volume of 40 mL.
Chitin Synthase (CS)
This protocol was developed for S. cerevisiae
chitin synthase enzymes CSI, CSII, and CSIII
[48]. The in vitro activities of other fungi may be
different and the protocols may need to be
adapted.
Membrane Extracts Preparation
As for the GS, the source of enzyme activity is a
crude membrane extract. The protocol for membrane extract preparation is similar to that
described for GS except that the buffer is 50 mM
Tris–HCl pH 7.5. Similarly, the membranes are
resuspended in the same buffer containing 33%
glycerol.
CS Assay
CS is a zymogenic enzyme that must be degraded
partially to show its maximal activity. As mentioned previously, three CS activities have been
described in S. cerevisiae, corresponding to three
different proteins. A method to determine the
three activities in the same membrane preparation by the use of several modifications in the
reaction conditions has been described [48]. Most
of the in vitro CS activity corresponds to CSI.
The reaction mixture contains 37 mM Tris–HCl
pH 7.5, 4.8 mM magnesium acetate, 5–10 mL of
membrane suspension (up to 20 mL) and 2 mL of
trypsin at the optimal concentration for activation
(0.1–2.0 mg/mL) in a total volume of 40 mL. The
mixture is incubated 15 min at 30 °C and the proteolysis is stopped by adding 2 mL of soybean
182
trypsin inhibitor at a concentration 1.5 times that
of the used trypsin solution. The tubes are placed
on ice and made 1 mM in UDP-[14C]-GlcNAc
(400 cpm/nmol) and 32 mM in GlcNAc in a total
volume of 46 mL. Samples are incubated for
30–60 min at 30 °C, stopped with 1 mL of 10%
TCA, and processed as for the GS assay (see previous). The specific activity is expressed as
hmoles of GlcNAc incorporated per hour and mg
of protein.
For CSII and CSIII, the reaction mixture
before proteolysis contains 32 mM Tris–HCl pH
8.0, 5 mM cobalt acetate, 20 mL of membrane
suspension, 2 mL of trypsin, and 1.1 mM in UDP[14C]-GlcNAc (400 cpm/nmol) in a total volume
of 46 mL. For CSIII, the reaction mixture also contains 5 mM nickel acetate. After proteolysis is
stopped, the mixture is made 32 mM in GlcNAc
in a volume of 50 mL, incubated for 90 min at
30 °C, and stopped with 1 mL of 10% TCA.
Although the CSI assay detects the three
activities, CSII and CSIII are minor contributors
and therefore they do not alter significantly the
value of CSI activity. However, they can be calculated based in the inhibitor effect of Ni2+ and
Co2+. Ni2+ is a powerful inhibitor of CSI and II but
has little effect on CSIII and Co2+ stimulates CSII
and III but inhibits CSI. In summary:
CSI activity (total CS—CSII + III): CS assay
minus CS assay (+Co2+).
CSIII activity: CS assay (+Co2+, +Ni2+).
CSII activity (CSII + III—CSIII): CS assay
(+Co2+) minus CS assay (+Co2+, +Ni2+).
Microtiter-Based Fluorescence Assay
As for GS, an alternative to the use of UDP-[14C]GlcNAc for the CS assay has been described
[49]. The CS reaction is similar but the radioactive UDP-[14C]-GlcNAc is omitted. The wells of
a microtiter plate are coated with wheat germ
agglutinin (WGA) which binds with high affinity
and specificity to chitin. The procedure involves
the binding of the synthesized chitin to the WGAcoated surface. Then, horseradish peroxidase–
WGA conjugate is added to the mixture. The
WGA of the conjugate will bind to the chitin
previously fixed in the well. The horseradish
peroxidase activity is measured at 600 nm, and
P. Pérez and J.C. Ribas
the amount of chitin is calculated using acidsolubilized chitin as standard. This method is
suitable for the three CS activities.
In Situ CS Assay Using Permeabilized
Whole Cells
Similar to the procedure described previously for
GS [38, 45–47].
Analysis of Cell Wall Polysaccharides
Different methods can be used to analyze the cell
wall polysaccharide composition. All the methods, either for a precise analysis or for a rapid
estimation of cell wall polymers, require a separation of the wall from the rest of the cell components. In general, the current methods have been
adapted for a simple, accurate and rapid analysis
of wall polysaccharides and are all based on
labeling and fractionation of cell wall polysaccharides using chemical and enzymatic procedures. Basically, the methods available have been
established using S. cerevisiae and S. pombe
models, although the techniques used can easily
be adapted for any organism.
Radioactive Labeling and Fractionation
of the Cell Walls
A basic procedure to quantify the cell wall polymers consists in 14C-glucose labeling and fractionation of the cell wall polysaccharides as
follows (Fig. 12.3):
Cell Wall Labeling
1. Exponentially growing cultures are adjusted to
5 × 106 cells/mL, 5–7 mL are supplemented with
[U-14C]-glucose (3 mCi/mL) (HartmannAnalytic)
and incubated for at least one doubling time (3 h)
to allow 14C incorporation into the cell. One doubling time means 50% labeling of cell material.
If a stronger labeling is required, the cultures can
be incubated for longer times, the 14C-glucose
can be increased (up to 18 mCi/mL), and the glucose concentration in the culture medium can be
reduced to 1 or 0.5%.
12
Fungal Cell Wall Analysis
183
Fig. 12.3 Overview of [U-14C]-glucose radioactive labeling and purification of the fungal cell wall
2. Cells (1–2 × 107 cells/mL) are harvested by
centrifugation (5,000 × g for 5 min).
3. Centrifuged cells from exponentially growing
cultures without radioactive glucose (300 mL
of a concentrate of 109–1010 cells/mL) are
added to minimize lost material.
4. Cells are washed twice with 1 mM EDTA,
transferred to 1.5–2.0 mL tubes and resuspended in 1.1 mL of 1 mM EDTA, 1 mM
PMSF.
5. Total glucose incorporation is monitored in
two 50 mL aliquots added to 1 mL of cold 10%
TCA and kept at 4 °C for at least 30 min. Then
aliquots are filtered through a fiberglass filter
Whatman GF/C, washed three times with
1 mL of 10% TCA, twice with 1 mL of ethanol, and counted in a liquid scintillation counter. Eventually, the samples can be stored in
10% TCA at 4 °C and analyzed with the samples obtained in further steps.
6. The remaining cells (1.0 mL) are centrifuged
(5,000 × g 5 min), resuspended in 100 mL of
1 mM EDTA, 1 mM PMSF, filled with cold glass
beads (0.5 mm diameter) to completely cover
the cell suspension and broken in a FastPrep
homogenizer (Q-Biogene, MP Biomedicals,
Thermo Scientific) during 3× 20 s pulse at a
speed of 6.0 and at 4 °C. Complete cell lysis is
confirmed by microscopic observation.
7. The broken cells are collected and the glass
beads are washed twice with 1 mM EDTA,
1 mM PMSF to collect all residual material.
8. The broken material is centrifuged at 1,500 × g
for 5 min, washed three times with 5 M NaCl,
then again twice with 1 mM EDTA.
9. The cell wall pellet is resuspended in 1.1 mL
of 1 mM EDTA, 1 mM PMSF, 0.02% Na
azide, and heated at 100 °C for 5 min, to inactivate the intrinsic hydrolytic cell wall enzymes
that would interfere the wall analysis.
Total radioactivity incorporated into the cell wall
is monitored in two 50-mL aliquots that are added
to 1 mL of cold 10% TCA and processed as
described previously.
184
Cell Wall Alkali-Fractionation and
Analysis
The most common method for cell wall fractionation is that used for S. cerevisiae [50, 51], which
can easily be adapted for other organisms. This
method allows the separation of cell wall b-glucan into alkali-soluble and alkali-insoluble fractions. The alkali-soluble fraction contains
b(1,3)-glucan, mannan, and some b(1,6)-glucan;
and the alkali-insoluble fraction contains chitin
and b(1,3)-glucan b(1,6)-glucan linked to the
chitin.
1. The cell wall suspension (1 mL) is extracted
twice with 6% NaOH for 90 min at 80 °C and
centrifuged at 1,500 × g for 5 min.
2. The alkali-extracted supernatant is divided
into four aliquots of 250 mL.
2.1. Two alkali-extracted aliquots are used to
precipitate the mannan with Fehling´s
reagent [52] as follows: unlabeled purified
mannan from S. cerevisiae (Sigma) is
added to the supernatant as carrier (0.1 mL
from a stock of 50 mg/mL in water).
Fehling´s reagent (2 mL) is then added to
the samples, mixed, and left overnight at
4 °C to precipitate the mannan. Fehling’s
reagent is freshly prepared for each experiment by adding one volume of reagent B
(3.5% CuSO4) to one volume of reagent A
(17.3% potassium sodium tartrate dissolved in 12.5% KOH). After centrifugation at 1,500 × g for 10 min, the pellet is
washed with Fehling´s reagent and solubilized in 20–40 mL of 6 N HCl (drop by
drop and mixing until completely solubilized). Then, 100 mL of 50 mM Tris–HCl
pH 7.5 is added and the solution is transferred to a vial with 2 mL of liquid scintillation. The tube is washed twice with
100 mL of buffer to collect the residual
mannan, which is added to the vial containing liquid scintillation and analyzed
(total mannan fraction).
2.2. The other two alkali-extracted aliquots
are precipitated with 2 volumes of ethanol, allowed to dry, dissolved in 100 mL
of water, collected (washing the tube
twice) and analyzed with liquid scintilla-
P. Pérez and J.C. Ribas
tion as the previous samples (mannan + alkali soluble glucan). The
difference between both fractions is the
alkali-soluble glucan (b1,3 + b1,6).
3. The alkali-insoluble residue is washed with
water several times by centrifugation until it
reaches a neutral pH, then suspended in
1.3 mL of water and divided into six aliquots
of 200 mL.
3.1. The radioactivity of two aliquots is
counted directly (alkali insoluble
glucan + chitin).
3.2. Two aliquots are incubated for 24–36 h at
30 °C with 25 mg Zymolyase-100T in
50 mM citrate-phosphate pH 5.6, 0.02%
Na azide in a volume of 300 mL, and two
aliquots are processed similarly but without Zymolyase as a control. After incubation, the four samples are centrifuged and
the pellets are washed twice, resuspended
in 100 mL of buffer, added to 1 mL of
10% TCA, and processed by filtration in
Whatman GF/C glass fiber filters and
scintillation counting as described above.
The residue remaining after Zymolyase
digestion is the chitin fraction. Treatment
of the alkali-insoluble fraction with
recombinant chitinase from Pyrococcus
furiosus (Wako Pure Chemical Industries)
is not needed because the remaining residue is the alkali-insoluble glucan
(b1,3 + b1,6), which can be obtained as a
difference between fractions.
Alkaline extraction is a widespread procedure for
cell wall analysis of many organisms [1, 25, 53–
59]. However, the data are only reproducible
when maintaining the extraction conditions; the
proportions of alkali-soluble and insoluble fractions can change depending on the alkali concentration, temperature, and incubation time. In fact,
a process such as storing the cell walls at −20 °C
results in total alkali solubility of the cell wall
under conditions similar to those described above,
showing only a small residue coincident with the
chitin fraction.
Chemical fractionation of S. pombe cell wall
polysaccharides is similar to that of S. cerevisiae
[60], although the cell wall composition is
12
Fungal Cell Wall Analysis
different and therefore the results differ. The
alkaline extraction procedure is rather harsh in S.
pombe, due to the absence of the chitin responsible for the alkali-insoluble maintenance of some
b(1,3)-glucan, and to a considerably smaller
amount of b(1,6)-glucan than in S. cerevisiae [61,
62]. Therefore, alkaline extraction causes solubilization of nearly all S. pombe cell wall polymers
and as a result, more gentle methods involving
enzymes capable of specifically digesting one
polymer without altering the others, described
below, yield more accurate results.
Cell Wall Enzymatic Fractionation
and Analysis
This protocol was adapted for S. pombe cell wall.
The most commonly used procedure permits
quantification of the three major cell wall polymers, b-glucans, a-glucans and galactomannoproteins. Once cell wall labeling and purification
has been performed as described above, the procedure is as follows:
1. Half of the cell wall material (500 mL) is
divided in:
1.1. Two 100 mL aliquots that are incubated
for 24–36 h at 30 °C with shaking with
25 mg Zymolyase-100T in 50 mM citratephosphate pH 5.6, 0.02% Na azide in a
volume of 300 mL, and two aliquots are
processed without enzyme as control. A
similar option is the incubation with 100–
200 mg of Kitalase in 50 mM potassiumacetate pH 5.0, 0.02% Na azide.
1.2. Two 100 mL aliquots are incubated for
24–36 h at 30 °C with 100 units of
Quantazyme (MP Biomedicals) or 100
units of recombinant b-1,3-Glucanase
Yeast Lytic Type (Wako Pure Chemical
Industries) in 50 mM potassium phosphate monobasic pH 7.5, 60 mM b-mercaptoethanol, 0.02% Na azide in a volume
of 300 mL.
1.3. Two 50 mL aliquots are processed without
enzyme as control.
2. After incubation, the samples are centrifuged
(16,000 × g for 3 min). The supernatant is
removed and 2 × 50 mL aliquots are counted
directly with 2 mL of liquid scintillation. 1 mL
185
of 10% TCA is added to the pellet and the
radioactivity incorporated is determined by
filtration and liquid scintillation counting as
described
previously. As
mentioned,
Zymolyase-100T and Kitalase contain b(1,3)glucanase, protease, mannanase and other
activities but not a-glucanase activity.
Therefore, the residue obtained after
Zymolyase-100T or Kitalase digestion is considered a-glucan and the supernatant, b-glucan plus galactomannan. Quantazyme and the
b-1,3-Glucanase Yeast Lytic Type are two
recombinant endo-b(1,3)-glucanases capable
of digesting b(1,3)-glucan without degrading
the b(1,6)-glucan or a-glucan [5, 6]. Therefore,
the residue obtained after Quantazyme or similar recombinant b(1,3)-glucanase treatment is
considered a-glucan plus b(1,6)-glucan and
galactomannan, and the supernatant is b(1,3)glucan.
3. Half of the cell wall material (500 mL) is
divided in two aliquots for galactomannan
quantification. The wall is solubilized in alkali
(6% NaOH) by adding 250 mL of 12% NaOH
and heating at 80 °C for 1 h. Then, the galactomannan is precipitated from the alkali-solubilized aliquots with the Fehling’s reagent and
quantified as described previously for S. cerevisiae cell walls.
Besides Zimolyase 100T, Kitalase and
Quantazyme, there is an ample variety of enzymes
and enzyme complexes commercially available
that can be used for the cell wall determination of
a specific fungal organism. These enzymes can
provide information either individually or in
combination. In the later case, depending on the
cell wall polysaccharides composition and linkages, the order of enzymes can be important for
maximal degradation of each enzyme. In addition, the enzymes with exohydrolytic activity
present poor or null activity against the cell wall
polysaccharides and therefore, all the enzymes to
be tested should contain endohydrolytic activity.
Some of these enzymes are:
• b(1-3;1-4)-d-glucan hydrolases such as that
from Bacillus subtilis (Biosupplies). This endoglucanase specifically hydrolyzes b-d-glucans
containing both b(1,3) and b(1,4)-d-glucosidic
186
•
•
•
•
linkages in linear sequence. It does not hydrolyze b(1,3)-glucans or b(1,4)-glucans.
a(1,4)-glucanases or a-amylases. These
enzymes are endoglucanases that hydrolyze
a(1,4)-d-glucosidic linkages in polysaccharides containing three or more a(1,4)-linked
d-glucose units. Examples of a-amylases that
can be used are a-amylase from Bacillus subtilis (heat stable, Sigma), from Bacillus licheniformis (Termamyl-120, heat stable up to 90 °C,
Sigma), from Aspergillus oryzae (TakaDiastase, Taka-Amylase A, Sigma), from
Rhizopus sp. (Merck Calbiochem), from porcine pancreas (Sigma), from human pancreas
(Merck Calbiochem) or from human saliva
(Sigma, Merck Calbiochem). No additional
activity for these enzymes has been reported.
Chitinases. These enzymes hydrolyze internal
linkages in the chitin chain, a linear polymer
of b(1,4)-N-Acetyl-d-glucosamine units. A
recombinant Chitinase (Wako Pure Chemical
Industries) from Pyrococcus furiosus is a thermostable and powerful enzyme. Chitinases
from Trichoderma viride (Sigma) and from
Streptomyces griseus (Sigma) are combination of exo and endochitinases, but with less
efficient hydrolytic activity.
Chitosanases. These enzymes catalyze the
hydrolysis of b(1,4) linkages between d-glucosamine (GlcN-GlcN) residues in chitosan.
Chitosanases from Streptomyces sp. (Sigma;
Merck Calbiochem) are available enzymes.
Mannanases. These are a-mannosidases that
cleave terminal a(1,2), a(1,3) and/or a(1,6)linked mannose residues in mannan. The supplied enzymes do not contain contaminant
protease or glycosidase activities. a(1,2;1,3)mannosidase, recombinant from Xanthomonas
manihotis (Merck Calbiochem) cleaves terminal a(1,2) and a(1,3)-linked mannose residues.
Its activity is efficient and reproducible. a(1,6)mannosidase, recombinant from X. manihotis
(ProZyme, AMS Biotechnology), cleaves terminal a(1,6)-linked mannose residues. It is
recommended for use after digestion with
a(1,2;1,3)-mannosidase for increasing the
degradation efficiency. Other available
enzymes are a(1,2;1,6)-mannosidase and
P. Pérez and J.C. Ribas
a(1,2;1.3;1,6)-mannosidase from Canavalia
ensiformis, Jack bean (Sigma, ProZyme, AMS
Biotechnology), and a(1,2)-mannosidase from
Aspergillus saitoi (ProZyme).
• Proteases such as Proteinase K, recombinant,
from Tritirachium album (Roche Applied
Science); Protease S, recombinant thermostable from Pyrococcus furiosus (Sigma);
Turbo3C protease, recombinant, from human
rhinovirus 3C protease; and aminopeptidase
T, recombinant, thermostable from Thermus
aqualicus (Wako Pure Chemical Industries).
• Enzymatic complexes. They can be used individually or in combination with other
enzymes. The enzymatic composition of
some complexes is well characterized
although they must be used cautiously because
they may contain other not-tested activities.
The enzymatic activities of other complexes
are not characterized and therefore can only
be used to test whether the present and absent
activities are helpful for a specific cell wall
analysis. We already mentioned Zymolyase20T and 100T (AMS Biotechnology) and
Kitalase (Wako Pure Chemical Industries,
City Chemical) also sold as Lysing Enzymes
from Rhizoctonia solani (Sigma) or as Yeast
Lytic Enzyme from Rhizoctonia solani (MP
Biomedicals). Uskizyme (Wako Pure
Chemical Industries) is a preparation from
Trichoderma sp. with b(1,3)-glucanase, cellulase, protease, and chitinase activities.
Westase (Cosmo Bio) is a preparation from
the liquid culture supernatant of Streptomyces
rochei. This complex contains mainly b(1,3)glucanase and b(1,6)-glucanase activities
according to the specifications sheet of the
manufacturer, but minor or absent activities
are not reported. Driselase (Sigma, Kyowa
Hakko Kogyo) is a crude powder from
Basidiomycetes sp. containing laminarinase,
xylanase and cellulase activities. Glucanex or
lysing enzymes from Trichoderma harzianum
(Sigma), previously known as Novozyme-234
(Novozymes Corp., discontinued) contains
b-glucanase, cellulase, protease, and chitinase
activities. It also contains a(1,3)-glucanase
activity, although it is not reported in the
12
Fungal Cell Wall Analysis
specifications sheet. Viscozyme (Sigma,
Novozymes Corp.) is a multi-enzyme complex from Aspergillus sp. containing a wide
range of carbohydrases, including arabanase,
cellulase, b-glucanase, hemicellulase, and
xylanase.
Other Cell Wall Chemical Fractionation
and Analysis
Other methods that require more time and effort
consist of combinations of enzymatic cell wall
degradations (and dialysis), chemical degradations, and analytical techniques that permit the
determination of the degradation products. These
methods usually give more precise information
about the type of bonds between the units forming
the polymers. Common chemical degradations
include alkali solubilization, acid hydrolysis, periodate oxidation, Smith degradation, borohydride
reduction, b-elimination, carboxymethylation, and
permethylation. Common analytical techniques
include determination of reducing sugars, of total
sugars, of glucose, of glucosamine, methylation
analysis, gas–liquid chromatography, mass spectrometry, paper chromatography, gel filtration
(size-exclusion) chromatography, ion-exchange
chromatography, thin-layer chromatography
(TLC), affinity chromatography, high-performance
anionic-exchange chromatography (HPAEC),
nuclear magnetic resonance (NMR) spectroscopy,
and X-ray diffraction [6, 7, 53, 57, 60–70].
Colorimetric Determination
of the Cell Wall Chitin
This is a colorimetric method useful to evaluate
the amount of chitin in the cell wall [71]. This
method is less precise than [14C] labeling and
measurement but it is faster and less toxic.
1. Cell cultures (200 mL) are harvested at early
log-phase (A600 of 1.0, aproximately 200 mg
wet weight), washed twice with water, and the
wet cell pellet is weighed.
2. Cells are resuspended in 1.0 mL water and a
volume corresponding to 100 mg of cells is
transferred to glass tubes.
187
3. The walls of 100 mg cells are extracted in
1 mL of 6% KOH at 80 °C for 90 min and the
suspension is cooled down, neutralized with
100 mL glacial acetic acid, transferred to a
1.5 mL tube (washing the glass tube with
water) and centrifuged 1 min at 16,000 × g.
4. The insoluble cell wall material is washed
three times with water, resuspended in 600 mL
of 50 mM potassium phosphate pH 7.5, and
incubated with 1 unit (5 mL) of recombinant
chitinase from Pyrococcus furiosus (Wako
Pure Chemical Industries) at 85 °C for 2 h.
Then, the degraded cell wall material is incubated with 25 mL of Glusulase (PerkinElmer)
at 37 °C for 1 h, stopped at 100 °C for 1 min
and centrifuged 1 min. Glusulase degrades the
chitobiose and small chitin fragments formed
by the endo-chitinase treatment into GlcNAc
monomers.
4. The amount of GlcNAc is quantified colorimetrically by the Reissig method [72] using
different concentrations of GlcNAc (0, 0.02,
0.04, 0.06, 0.08, and 0.1 mmol) as standards.
Each reaction contains 250 mL of water, 250 mL
of 270 mM potassium tetraborate pH 9.5 and
0, 2, 4, 6, 8 and 10 mL of 10 mM GlcNAc. The
samples contain 250 mL of 270 mM potassium
tetraborate pH 9.5, 200 or 150 mL of water and
50 or 100 mL respectively of the degraded cell
wall material. The samples are boiled (100 °C)
during 8 min, cooled down in ice/water and
3 mL of Reissig reagent is added and mixed.
The Reissig reagent must be prepared freshly
and contains 1 g of 3,5-diamino-benzaldehyde,
1.25 mL of 37% HCl and glacial acetic acid up
to 100 mL. The reactions are done in duplicate.
The samples are incubated 40 min at 37 °C and
the generated color is measured at 585 nm
(quartz cuvettes).
Cell Wall b(1,6) Glucan Determination
The most common method for b(1,6)glucan
determination consists of alkaline extraction of
the cell wall and analysis of the alkali-insoluble
b(1,6)-glucan [73, 74], although part of the
188
b(1,6)-glucan is in the alkali-soluble fraction
[63, 75].
1. Isolated cell walls from 50 mL cultures are
obtained as described in page 9 and Fig 12.3,
and extracted three times with 1 mL of 3%
NaOH at 75 °C for 1 h (removes mannoproteins and alkali-soluble glucan).
2. The extracted walls are washed once with
1 mL of 100 mM Tris–HCl pH 7.5, once with
1 mL of 10 mM Tris–HCl pH 7.5, resuspended
in 1 mL of 10 mM Tris–HCl pH 7.5 and incubated with 1 mg of Zymolyase-100T at 37 °C
for 16 h. Approximately 90% of the glucosecontaining carbohydrate is released into the
supernatant. Zymolyase releases the b(1,6)glucan to the supernatant.
3. The insoluble material is removed by centrifugation (13,000 × g, 15 min) and the supernatant is dialyzed against water (6–8 kDa pore
size) for 16 h. The carbohydrate retained after
dialysis is the amount of b(1-6)-glucan. Total
carbohydrate of each alkali-insoluble fraction
(dialysis-retained, Zymolyase-soluble and
Zymolyase-insoluble) is measured as hexose
by the borosulfuric acid or phenol-sulfuric
methods [76, 77]. The alkali-insoluble b(1,6)glucan is determined as percentage of total
carbohydrate (the sum of both the Zymolyasesoluble and insoluble fractions). The alkaliinsoluble b(1,3)-glucan is determined as the
amount of Zymolyase-soluble material before
dialysis subtracted from the amount of alkaliinsoluble b(1,6)-glucan.
The alkali-soluble b(1,6)glucan can be detected
by immunodetection [75, 78]. The alkali-soluble
extract is spotted onto nitrocellulose, dried,
blocked with 5% nonfat milk in TBST (10 mM
Tris pH 8.0, 150 mM NaCl, 0.05% Tween 20)
and probed with anti-b(1,6)-glucan antibody
[70]. After antibody binding, the membrane is
washed three times with TBST, probed with antirabbit IgG horseradish peroxidase secondary
antibody, washed again three times with TBST,
and visualized with an ECL detection kit
(Amersham). The alkali-soluble b(1,3)glucan
can also be detected with the same procedure by
using commercial monoclonal anti-b(1,3)-glucan
antibodies (Biosupplies) and anti-mouse
P. Pérez and J.C. Ribas
secondary antibody. The amount of b(1,6)glucan
or b(1,3)glucan is quantified in dots by using different concentrations of (b(1,6)glucan (pustulan)
or b(1,3)-glucan (pachyman, curdlan, or laminarin) as standards, respectively.
b(1,6)-glucan can also be determined by highperformance liquid chromatography (HPLC), gas
chromatography, mass spectrometry and NMR
analysis of oligosaccharides after alkaline extraction and/or enzymatic degradation of the cell wall
polymers.
Determination of Cell Wall Proteins
The cell wall proteins are glycoproteins highly
modified with O- and N-linked oligosaccharides,
predominantly or exclusively formed by mannose residues known as mannan. In some cases,
the mannan backbone presents single residues or
side chains of different sugars, galactomannan,
rhamnomannan, glucogalactomannan, rhamnogalactomannan, etc. [1, 24, 25].
Most proteins found in the cell wall are water- or
detergent-soluble and are usually secreted to the
medium. Some few cell wall proteins are covalently
linked to the polysaccharides and can be divided
into two groups, proteins covalently attached to
b(1,3)glucan (Pir proteins) through a glutamine
residue, an alkali-labile linkage that can be extracted
by a mild alkali treatment (30 mM NaOH at 4 °C
for 16 h); and proteins covalently attached by a GPI
anchor to the b(1,6)glucan of a b(1,6)/b(1,3)glucan
core that can be removed by b(1,3)glucanases or
b(1,6)glucanases [1, 59, 79–82].
The SDS-soluble proteins are extracted by hot
SDS-mercaptoethanol treatments, twice with
50 mM Tris–HCl pH 7.8–8.0, 2% SDS, 100 mM
EDTA, and 40 mM b-mercaptoethanol for
5–15 min at 100 °C [70, 83, 84]. The proteins are
concentrated and the SDS removed by precipitation with 9 volumes of cold acetone at −20 °C for
2 h. The proteins are dried, resuspended and analyzed by SDS-PAGE or Western blot.
The covalently attached cell wall proteins can
be purified by three methods [82, 84]:
1. The proteins covalently attached to b(1,3)glucan are released by treatment with 30 mM
12
Fungal Cell Wall Analysis
NaOH at 4 °C for 16 h. The reaction is stopped
by adding acetic acid to neutrality followed by
dialysis.
2. The GPI-attached proteins are released by
degradation of the GPI anchor by treatment
with undiluted pyridine hydrofluoride at 24 °C
for 16 h. HF-pyridine is removed by dialysis.
Treatment with recombinant endo-b(1,6)-glucanase (noncommercial) also releases the GPI
proteins.
3. Both groups of covalently attached proteins
can be released by treatment with recombinant
endo-b(1,3)-glucanase (MP Biomedicals;
Wako Pure Chemical Industries), followed by
dialysis.
The mannan oligosaccharides can be analyzed
directly or after protease treatment by enzymatic
treatment with different mannosidases. As the
glycoproteins are highly glycosylated and this
may interfere with the electrophoresis protein
analysis, the N-linked oligosaccharides can be
removed by treatment with endoglycosidase H
(Boehringer, Roche, New England Biolabs) [70,
85]. The proteins can be analyzed by SDS-PAGE
or Western blot with specific antibodies or with
the lectin concanavalin A (Sigma) that specifically
binds to the mannan region [81, 86]. Additionally,
the cell wall proteins can be biotin-labeled
directly in the cell prior to cell wall purification,
with the biotinylation reagent Sulfo-NHS-LCBiotin (Pierce) in 50 mM potassium phosphate
pH 8.0 by incubation for 90 min on ice. The proteins are analyzed by Western blot and visualized
with streptavidin-horseradish peroxidase conjugate (Pierce) [87, 88].
The mannan can be analyzed by incorporation of radioactive orthophosphate into
N-mannosylated glycoproteins. The cell wall is
purified and extracted with SDS to obtain part of
the mannan. Then the insoluble material is treated
with b(1,3)glucanases to release the rest of the
mannan. The phosphate is bound as mannose-6phosphate to both soluble and covalently linked
cell wall mannoproteins [83, 89]. The phosphorylated cell wall oligosaccharides can be characterized after hydrolysis in trifluoracetic acid
(TFA) by Quaternary aminoethyl (QAE)Sephadex A50 chromatography, Bio-Gel P2
189
chromatography, HPAEC, electrospray ionization tandem mass spectrometry (ESI-MS-MS),
and methylation analysis with gas chromatography–mass spectrometry [83].
Nonquantitative Methods for Analysis
of Cell Wall Polysaccharides
Other methods can also be used in order to detect
cell wall differences in structure or composition
without the need of precise polymer quantification.
Among them, those more used are briefly
described as follows.
Sensitivity of Cells to Enzymatic
Degradation
This method is used for a rough analysis of the
cell wall state and is used to corroborate other
results that suggest an altered cell wall [29, 90].
It is also used as a screening for mutations affecting the cell wall. Common enzymes used in this
procedure are Zymolyase-100T in 50 mM citrate/phosphate buffer pH 5.6, Kitalase in 50 mM
potassium-acetate pH 5.0, or Glucanex (Sigma)
in any of these buffers. Zymolyase-100T and
Kitalase mainly degrade b(1,3)-glucan and mannoproteins, whereas Glucanex degrades the
entire cell wall. Increased sensitivity of the cells
to these enzymes can be due to different causes
such as a decrease in the amount of cell wall
b-glucan or an increase in the permeability of the
cell wall to the enzymes, which can be caused by
a decrease in the cell wall surface glycoproteins
or an altered cell wall structure. A specific cell
wall-related mutant strain may show different
sensitivities to Zymolyase, Kitalase and
Glucanex, depending on the importance that the
a-glucan may have in maintaining cell integrity
in this mutant strain. The sensitivity to degradation using a(1,3)-glucanase has never been
assayed due to the lack of a commercially available enzyme, although the purification of recombinant a(1,3)-glucanases (mutanases) from
Penicillium purpurogenum and Trichoderma
harzianum has been described [91].
190
Hypersensitivity or Resistance to Cell
Wall Biosynthesis Inhibitors
This method can be used to detect changes in the
cell wall composition or structure, and to detect
mutations in the enzymes involved in cell wall
synthesis; it is also used in the analysis of genetic
interactions between genes that might be
specifically related to cell wall biosynthesis and
cell integrity. The main inhibitors are:
• Calcofluor white used to stain chitin.
Calcofluor also binds to linear b(1,3)glucan
with high affinity when chitin is not present,
as in the case of S. pombe [92]. The binding of
this dye perturbs the wall structure at low concentration and halts cell growth at high
concentration.
• Echinocandins (caspofungin, micafungin and
anidulafungin, available for clinical use) are a
family of antifungal drugs that specifically
inhibit both, the in vitro b(1,3)-glucan synthase activity and the in vivo b(1,3)-glucan
synthesis [28, 93–97]. Other families of
b(1,3)-glucan synthase inhibitors like papulacandins and the acidic terpenoid enfumafungin
are also used [29].
• Polyoxins and Nikkomycins are chitin synthase inhibitors [23, 94, 96] or recently discovered b(1,6)glucan synthase inhibitors can
also be used for specific assays [98, 99].
• 2-deoxi-d-glucose can be used as competitor
of glucose for assays of cell wall biosynthesis
defects.
P. Pérez and J.C. Ribas
Similarly, aniline blue (0.5 mg/mL final concentration) (Biosupplies), specifically stains
linear b(1,3)glucan, although its affinity for the
cell wall b(1,3)glucan of growing poles is
lower than that of Calcofluor.
• Lectins against the mannan of cell wall glycoproteins such as Concanavalin A, which binds
mannose residues, the lectin from
Bandeiraea simplicifolia that recognizes
specifically terminal galactose residues or
wheat germ agglutinin (WGA) that recognize
specifically chitin. They can be used for
immunofluorescence analysis bound to
fluorescein isothiocyanate (FITC) (Sigma).
Lectins are added (200 mg/mL) to the cells, in
culture medium or in phosphate buffer solution (PBS) if the cells are fixed. After 15 min
in dark, cells are washed and resuspended for
microscopy observation. For the observation
of chitin, since it is internal to the cell wall,
WGA-FITC binding and visualization
requires a previous mild alkali extraction of
the cell wall. The same lectins, labeled with
colloidal gold, can be used for immunoelectron microscopy.
• Antibodies raised against specific proteins or
polymers can also be used (see below). These
antibodies are combined with secondary
fluorochrome-labeled
antibodies
for
immunofluorescence studies.
Electron Microscopy (EM)
Fluorescence Microscopy
The techniques for EM are numerous and have
advanced greatly during the last years:
Direct observation of the cell wall using
microscopy techniques is an important method
for cell wall studies. Fluorochromes, lectins,
and antibodies that stain a specific cell wall
polysaccharide can be used for cell wall
fluorescence microscopy analysis. The most
commonly used are:
• Calcofluor white (Sigma) (25 mg/mL final concentration) stains the cell wall chitin and in its
absence specifically stains linear b(1,3)glucan.
Transmission Electron Microscopy
Cells fixed with glutaraldehyde have been used to
study the cell wall and septum structures of fungal cells. This technique permits observation of
the cell wall as a three-layered structure of polysaccharides with different electron densities (see
Fig. 12.1) [92, 100–109]. Negative staining for
TEM with uranyl acetate is an alternative method
for visualization of fibrils network and other cell
wall structures [110–114].
12
Fungal Cell Wall Analysis
Scanning Electron Microscopic (SEM)
Cells fixed with glutaraldehyde are used to
observe the cell surface clearly and with high
fidelity. An improved technique is ultra-highresolution low-voltage SEM (UHR-LVSEM)
[103, 110–113, 115–118]. SEM microscopy can
be coupled with lectins or antibodies labeled with
colloidal gold particles. This technique served to
detect a cell wall surface completely filled with
particles specific for the mannan carbohydrate of
glycoproteins [116], and to detect the b(1,3)-glucan in the bud scars of S. Cerevisiae [1].
Atomic Force Microscopy (AMF)
This technique is used to measure the mechanical
properties of the fungal cell wall macromolecules
[1, 113, 119, 120].
Cryoscanning and Cryosectioning
Electron Microscopy
This technique uses cryofixation to physically
immobilize the specimen [113]. An improved
method combines high-pressure freezing with
ultra-low temperature and low-voltage SEM
(ULT-LVSEM). It is useful to analyze fractured
and coated cell samples, allowing the observation
of a fine plane and the ultrastructure of both
external and internal cell components [121].
Transmission Immunoelectron
Microscopy (IEM)
Cells are fixed with paraformaldehyde plus glutaraldehyde. This technique has improved with
the method of cryofixation by high-pressure freezing followed by freeze-substitution to retain the
antigenicity [122]. Specific mono- and polyclonal
antibodies in combination with secondary antibodies conjugated with 10-nm gold particles have
been used to locate the different types of b-glucans [122]. The currently described antibodies
against different wall polysaccharides are:
• Anti-b(1,3)-glucan [123] (Biosupplies), a
murine monoclonal antibody without crossreactivity with b(1,4)-glucans or b(1,3;1,4)glucans.
191
• Anti-b(1,3;1,4)-glucan (Biosupplies). It is a
murine monoclonal antibody without crossreactivity with b(1,3)-glucans [124].
• Anti-b(1,6)-glucan. This rabbit antiserum
specifically recognizes this polymer in S. cerevisiae [70, 125] and S. pombe [122] but is not
commercially available.
• Anti-b(1,6)-branched-b(1,3)-glucan
[122,
126] is a rabbit antiserum obtained against grifolan (a type of b(1,6)-branched-b(1,3)-glucan) and does not reacts with linear
b(1,3)-glucan. It is suggested that the hapten
site of the antibody is the monoglucosyl
b(1,6)-glucan-branched moiety of b(1,3)-glucan. It is not commercially available.
• Anti-a(1,3)-glucan. A polyclonal anti-a(1,3)glucan has been used to analyze the cell wall
a(1,3)-glucan but is not commercially available [64, 127]. Additionally, the monoclonal
IgM antibody MOPC-104E (Sigma, Abcam)
has been used to detect the a(1,3)-glucan in
Histoplasma capsulatum yeast cell walls. This
antibody specifically recognizes a(1,3)-glucan because only a(1,3)-linked and not
b-linked glycosyl polysaccharides can block
the antibody and because the cell walls lacking a(1,3)-glucan of an avirulent strain of
Histoplasma capsulatum are not recognized
by the antibody [128–130].
• Anti-GFP murine monoclonal antibody (JL-8
anti-GFP; BD Biosciences, Sigma) has been
used to detect GFP-fused cell wall or plasma
membrane proteins with secondary gold-labeled
anti-mouse secondary antibodies [131].
The lectins mentioned previously, Concanavalin A, which binds mannose residues, and
wheat germ agglutinin (WGA), which binds
GlcNAc residues, are also used for IEM studies
when labeled with colloidal gold particles [109,
132–134].
Acknowledgements We thank D. Posner for language
revision. This work was supported by grants BFU201015641 and BIO2009-10597 from the Dirección General
de Investigación, MICINN, Spain, and grant CSI038A11-2
from the Junta de Castilla y León, Spain.
192
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Cabib E, Bowers B, Roberts RL (1983) Vectorial
synthesis of a polysaccharide by isolated plasma
membranes. Proc Natl Acad Sci U S A 80:
3318–3321
Molano J, Bowers B, Cabib E (1980) Distribution of
chitin in the yeast cell wall. An ultrastructural and
chemical study. J Cell Biol 85:199–212
Roberts RL, Bowers B, Slater ML, Cabib E (1983)
Chitin synthesis and localization in cell division
cycle mutants of Saccharomyces cerevisiae. Mol
Cell Biol 3:922–930
Histopathological Technique
for Detection of Fungal Infections
in Plants
13
Vijai Kumar Gupta and Brejesh Kumar Pandey
Abstract
Microscopic examination of the interaction between pathogenic fungi and
their host plants has been instrumental in deciphering the biology of this
relationship and can serve as a useful diagnostic tool. In this chapter, we
describe the technique of fixing fungal infections of plant samplings for
histopathological experiments. Toluidine blue O’ staining methods coupled
with stereoscopic microscopy are used to scan the infection structures of the
fungus Fusarium spp. and host response in Psidium guajava L. root
tissues.
Keywords
Fungal infections • Histopathological experiments • Microscopy •
Staining techniques • TOLUIDINE blue O’ • Fusarium spp. • Psidium
guajava L.
Introduction
V.K. Gupta (*)
Molecular Glycobiotechnology Group, Department
of Biochemistry, School of Natural Sciences, National
University of Ireland Galway, University Road,
Galway, Ireland
Assistant Professor of Biotechnology, Department
of Science, Faculty of Arts, Science & Commerce,
MITS University, Rajasthan, India
e-mail: vijai.gupta@nuigalway.ie; vijaifzd@gmail.com
B.K. Pandey
Molecular Plant Pathology Laboratory, Central Institute
for Subtropical Horticulture, Indian Council of
Agricultural Research, P.O.kakori, Rahmankhera,
Lucknow, UP 227 017, India
The ability to observe the growth of fungal
structures in host tissues under the microscope
is an important tool in the study of plant pathogenesis. Over the years many staining techniques that highlight fungal structures in plant
tissues have been reported. In particular, technologies such as stereoscopic microscopy have
enhanced our ability to visualize hyphae in plant
tissue [1–4].
The use of certain staining techniques can
facilitate considerably microscopic observations
and experimental research on plant pathology
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_13, © Springer Science+Business Media, LLC 2013
197
198
V.K. Gupta and B.K. Pandey
by allowing plant and fungal tissues to be differentiated. More specifically, staining can aid
examination of fungal colonization and infection processes, such as differentiating hyphae in
life cycles that involve a transition from a
biotrophic to a necrotrophic phase. Staining of
specific tissues also can simplify identification
of fungal inoculum or hyphal presence in asymptomatic plant tissue. The effectiveness of a particular staining technique can vary greatly
depending on the particular fungus and plant
species. Toluidine blue O’ has been used to stain
and identify callose deposition produced by host
plants in response to intracellular infection of
plant cells by fungi in some plant–fungus interactions [5]. Toluidine blue O’ staining techniques was applied to examine the infection
structures of the fungus Fusarium in root tissues
wilt-infected guava plants. The usefulness of
this staining method was based on the visual
contrast between host plant tissue and fungal
hyphae provided by polychromatic dye and resolution, and the relative ease of preparation and
use [6]. This study describes an improved
method for fixation of sampling of fungalinfected plant parts, and staining and observation of fungal infections in plant tissue for
histopathological visualization.
12. Microprocessor-based automatic tissue
processor (Electra, YSl 104, Yorko)
13. Microtome (MICROM—HM 350)
14. Stereoscopic microscope (Leica—LEITZ—
DM RBE)
Methods
The methods presented in the following sections
describe general procedures for fixation, staining, and microscopy of fungal infections of plant
samplings. Modifications that may be needed to
fix the sampling properly from different types
and sources of material are also described.
Killing and Fixation
Roots samples were collected from wilt-affected
and healthy plants. Root pieces 2–4 cm long were
cut and surface sterilized using 0.1% HgCl2,
washed two to three times in sterilized water, and
the excess water absorbed on Whatman filter
paper 41. Then samples were kept in formaldehyde: acetic acid: alcohol (5 mL: 5 mL: 90 mL)
for a minimum of 48 h (see Note 1).
Dehydration
Materials
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
Sterilized water
0.1% HgCl2
Glass slides
Whatman filter paper no. 41
Formaldehyde
Glacial acetic acid
Alcohol
Xylene
Paraffin wax
Toluidine blue O’
DPX-mount
The samples were processed with the
alcohol:xylene series (as per the flow chart
depicted in Fig. 13.1) using an automatic tissue
processor (Yorko) (see Note 2).
Infiltration and Embedding
The samples were embedded in melted paraffin
wax (54–56 °C) for at least 4–8 h in order to completely replace the xylene with paraffin wax in a
square-shaped block (see Note 3).
13
Histopathological Technique for Detection of Fungal Infections in Plants
199
bringing them to xylene through the alcohol:xylene
series. The detailed procedure is given in the flow
diagram depicted in Fig. 13.2 (see Note 5).
Microscopy and Imaging
Samples were mounted in 50% (v⁄v) DPX mount
and viewed under a stereoscopic microscope
(Leica—LEITZ DM RBE) using a Hoya CM500S
filter (IR cut-off 650 nm). Images were captured
using a CCD camera with a Bayer Array RGB
filter for brilliant pictures (Interline transfer frame
readout CCD—ICX252AQ) and Leica DFC
Twain and Leica Image Manager analysis software (soft microscopy with imaging control software system).
Notes
Fig. 13.1 Samples processed with the alcohol:xylene
series. This process removes the water from the plant tissues and facilitates sectioning
Sectioning
Section (10 mm thick) cutting was done using a
microtome. Blocks were prepared in paraffin wax
and thin sections 10 mm thick were cut with the
help of a microtome (MICROM—HM 350S).
At least 20 slides were prepared for each sample
(see Note 4).
Staining and Mounting
The sections were stained in 0.1% aqueous toluidine blue O’ and were mounted in DPX after
1. The FAA solution is prepared based on the
type of material, that is, soft tissue, moderate
tissue, or hard tissue (use 25% ethanol for
very delicate material, 50% for normal use,
and 70% ethanol for very tough material).
The samples were left in the FAA solution at
least 48 h or until they were processed further. This depends on the hardness of the
tissue.
2. This process removes the water from the plant
tissues and facilitates sectioning.
3. Infiltration and embedding of the material was
done in paraffin wax to remove the xylene
from the tissues. The blocks of wax were prepared in L molds in which the material was
embedded.
4. Sectioning of the material was done with the
automatic microtome (MICROM—HM 350S).
5. Sections were stained in 0.1% aqueous toluidine blue O’ and were mounted in DPX after
bringing them to xylene through the
alcohol:xylene series as described by
Jensen [7]. The samples were examined for
anatomical details as per the technique
described by Pandey [3].
200
V.K. Gupta and B.K. Pandey
Fig. 13.2 Sections were stained in 0.1% aqueous toluidine blue O’ and were mounted in DPX after bringing them to
xylene through the alcohol:xylene series
Acknowledgements The authors are very grateful to
Director, CISH; Head, Department of Crop-Protection,
Central Institute of Subtropical Horticulture (CISH),
Lucknow; and Prof. Shakti Baijal, Ex-Dean, FASC, MITS
University, Rajasthan for providing the necessary research
grants.
4.
5.
References
1. Johansen DA (1940) Plant microtechnique. McGrawHill, New York
2. Meyberg M (1988) Selective staining of fungal hyphae
in parasitic and symbiotic plant-fungus associations.
Histochemistry 88:197–199
3. Pandey BK (1984) Studies of chickpea blight caused
by Ascophyta rafiei (Pass) Labr. with special reference
6.
7.
to survival in crop debris [PhD thesis]. Department of
Plant Pathology. G. B. Pant University of Agriculture
& Technology, Pantnagar, UP, India
Saha DC, Jackson MA, Johnson-Cicalese JM (1988) A
rapid staining method for detection of endophytic fungi
in turf and forage grasses. Phytopathology 78:
237–239
Gupta VK, Misra AK, Pandey BK (2012)
Histopathological changes during wilting in guava
root. Arch Phytopathol Plant Protect 45(5):570–573
Available from http://www.tandfonline.com/doi/abs/1
0.1080/03235408.2011.588047
Knight NL, Sutherland MW (2001) A rapid differential
staining technique for Fusarium pseudograminearum in
cereal tissues during crown rot infections. Plant Pathol
Online. First-doi:10.1111/j.1365-3059.2011.02462.x
Jensen WA (1962) Botanical histochemistry: principals
and practices. WH Freeman, San Francisco, London
Development of Media
for Growth and Enumeration
of Fungi from Water
14
Segula Masaphy
Abstract
Fungi are found in water resources as natural primary inhabitants or as
secondary inhabitants that enter the water source as contaminants. Many
of the fungi in water resources can be directly harmful to human, animal,
and plant health, or cause problems in food processing and preparation, or
by producing biofilms in water-distribution systems. Hence, water fungi
are of concern for consumers. The ability to detect fungi in water sources
is therefore important with respect to minimizing the risk of contamination and for safety-management protocols. However, there is no one uniform method for determining fungal load in water. Various new
molecular-based methods are being developed to analyze water resources,
but the traditional colony-based ones are still the methods of choice for
enumeration and characterization of fungal populations in water. Recent
developments in those methods for water mycological quality examination, particularly with regard to media composition, are presented.
Keywords
CFU • Detection • Enumeration • Fungi • Membrane filtration • Biofilm
• Treated/untreated water • Routine analyses
Introduction
An awareness of the importance of fungi in
water destined for human consumption has
emerged in recent years. Although still limited,
S. Masaphy (*)
Applied Microbiology and Mycology Lab,
MIGAL, Galilee Technology Center, POB 831, 11016,
South Industrial Area, Kiryat Shmona, Israel
e-mail: segula@migal.org.il
the number of publications on this topic is
rising, demonstrating the presence of a range of
fungal species, some of which are known to be
directly pathogenic to humans, cause allergic
reactions, or have harmful effects due to their
production and release of toxins into the water
[1–3]. Fungi are also suspected of contributing
to negative organoleptic qualities in drinking
water [4], and to biofilm production in distribution systems [5]. Plant-pathogenic and foodspoilage fungi have also been found in treated
and untreated water [2].
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_14, © Springer Science+Business Media, LLC 2013
201
202
In terms of human health, there is relatively
little information on the role played by fungal
contamination of water in illnesses and infections
in the general public. However, the greater concern is related to immunosuppressed patients,
who may be infected by drinking water, bath
water, or recreational water bodies [6]. Hence,
studies have been aimed at examining the mycological quality of hospital water systems, to assess
the risk of fungal transmission to patients and
possible infection. Potentially health-disruptive
fungi have been detected in tap water [7–12],
water-distribution systems [10, 13], bottled water
[14, 15], bathing water [16], swimming pools [17,
18], surface water sources (rivers, streams, canals,
lakes, and ponds) [2, 7], and groundwater [19].
Species of potentially human-pathogenic
yeasts and molds recovered from drinking water
include Aspergillus fumigatus [20], Fusarium
[21], Penicillium [14], Aureobasidium pullulans
(found in saunas) [22], Absidia, Mucor, Candida
[23], Trichoderma viride [9], and Chaetomium
globosum [4]. Various potentially pathogenic
fungi, including dermatophytes, have also been
isolated from swimming pools, such as
Cladosporium spp., Penicillium spp., Aspergillus
spp., Rhizopus spp., Fusarium and Trichophyton
rubrum, Mucor spp. and Candida albicans [17,
18, 24], among others.
With the increasing concern about fungal contamination in consumed water, fungal examination of different water sources is on the rise, and
it is recommended that water resources be monitored for fungal contamination as part of watersafety management programs. However, the field
of mycological water quality, including methodologies and regulations, is far less established
than the fields of bacteriological water quality
and mycological food quality. In 1975, the methods for fungal detection in water and wastewater
were still only tentative, as laid out in the fourteenth edition of the Standard Methods for the
Examination of Water and Wastewater [25]. Even
today, on a global scale, there are different regulations and methodologies in place for mycological water safety, and no uniform or standardized
method for determining the mycological quality
of potable water resources has been recognized.
S. Masaphy
Water is very heterogeneous with regard to the
fungal diversity found within it. Fungi can be
found in water resources as natural primary
inhabitants or as secondary inhabitants that have
entered the water source as contaminants. As
summarized by Hageskal et al. [26] primary
inhabitants are those that are adapted to aquatic
environments, belonging to the phylum
Chytridiomycota. Secondary inhabitants are all
other fungi, which enter natural water bodies
from the air, soil, and wastewater. The survival
and proliferation of these latter fungi depend on
the characteristics of the water, that is, nutrition
load, temperature, pH, other microorganisms,
and, in some cases, the presence of disinfectants
(such as chlorine in swimming pools) [27].
The origin of the water source and its designated use also vary (e.g., tap water for drinking,
bottled water, swimming pool water, recreational
water, or wastewater), and these need to be considered in the establishment and application of a
detection method. Each of these water bodies
may support different fungal loads and diversities. It is clear that no one defined fungal indicator can give information on all of the different
types of damage that can be expected from each
water source. It is important to detect the individual fungus that is causing a problem and identify it, but quantifying the total fungal load may
give a faster and clearer estimation of the degree
of fungal contamination, so that appropriate
action can be taken.
Traditionally, quantification of fungi in water
sources has been based on culturing the colonies
on general nutritional media, or culturing specific
fungi on more selective media, and presenting the
fungal load as CFU (colony forming units) in a
certain volume of water. Although it is agreed
that not all fungal species in water will grow on a
specific medium under particular growth conditions, culturing and enumeration of as many
members of the fungal community in the water
source as possible can give a good estimation of
the level of fungal contamination. Although
efforts are being made today to develop rapid
molecular procedures for the detection of fungal
contamination (e.g., PCR-based protocols, ergosterol-content determination, the use of gene
14
Development of Media for Growth and Enumeration of Fungi from Water
probes, protein probes, and mass spectrometry),
culturing fungal colonies on agar media is still
the method of choice to enumerate and isolate
fungi of interest. This chapter focuses on examinations of mycological water quality using the
CFU approach, and on the recent developments
in media used for this purpose.
Fungal Culturing Approach
Historically, the examination of fungi in water
bodies has consisted of adopting analytical methods and media used to examine fungal loads in
foods. Hence, the “pour plate” and “spread plate”
techniques, used with solid products, were recommended for water examinations as well. Later, the
“membrane filtration” (MF) technique was also
recommended for examinations of fungi in water.
Techniques
Spread Plate Technique
This is a well-established technique for mycological examinations of food and environmental
samples. This and the pour plate technique, both
established and used for different types of food
and soil samples, were also adopted for water
examination. In this method, a water sample is
spread by glass or plastic spreader on the surface
of agar medium in a Petri dish. With a suitable
medium and incubation conditions, this technique enables the growth of a range of fungal
species, as most fungi are aerobic. The advantage
of this method is that the colonies can be differentiated by appearance, and the cultured fungi of
interest can be further isolated and identified separately. The disadvantage of this method is sample loss on the spreader, and the low species
diversity growing on the agar surface in comparison to the pour plate technique.
Pour Plate Technique
In this technique, the water sample is placed in
an empty Petri dish, to which molten medium is
then added. After mixing the two, the medium is
allowed to solidify, and the culture is incubated
203
for fungal growth. This method is well suited to
the enumeration of fungi since there is no loss of
sample as in the spread plate technique. In addition, this technique allows the germination and
growth of colonies inside the agar medium,
where each colony is surrounded by a homogeneous microclimate in terms of nutrient, oxygen,
and moisture levels. This allows for higher fungal diversity, as fungi can grow on the surface or
at different depths in the medium. The disadvantage is that it becomes difficult to isolate the colony of interest if that colony is growing inside
the medium.
Both the spread plate technique and the pour
plate technique are simpler and less costly than
the MF technique, but require higher quantities of
medium, and more space for plates incubation.
MF Technique
Today, this is the most commonly used technique
in routine microbiological quality examinations
of low-turbidity water (drinking, bottled, swimming pool, and bathing water). The MF technique was initially developed as an alternative to
other methods for bacteriological analyses of
water samples in the late 1950s [28, 29]. In 1975,
it was adopted as a standard procedure for bacterial water examination in the eleventh edition of
the SMEWW [30]. It was only in 1976 that
Qureshi and Dutka [31] examined the MF technique for the recovery of fungi from water, comparing different brands of MFs for this purpose.
In 1978, it was used for the enumeration of
Candida in natural water [32]. The procedure is
based on filtering a volume of water through a
MF, and the fungal units (conidia, chlamydospores, and other hyphal units) are trapped on
the membrane surface. The filter is then transferred onto nutritional agar medium and the fungal colonies are left to develop on the filter
surface. In their comprehensive work on the optimization of methodologies for fungal recovery
from water, Kinsey et al. [2] showed that this
technique recovers a lower number of different
taxa than the direct spread plate technique. In a
work conducted in our lab (S. Masaphy, unpublished data), the recovery of fungi (Penicillium
sp., Aspergillus sp., Candida albicans, and
204
Saccaromyces sp.) artificially inoculated into
bottled water, tap water, and swimming pool water
was similar using MF and spread plate techniques
in terms of fungal units. The MF technique has
several advantages, especially in examining
water with low microbiological loads. Moreover,
this technique enables examining a large number
of samples in a short period of time.
For all three of the aforementioned methods,
successful fungal recovery is related mainly to
selecting the right medium.
Media
Early media used for the detection of fungi in
water sources were similar to those used for the
detection of fungi in other products, since many
of the fungi in water actually reach the water
body from the surrounding environment. Today,
more specific media for water examination are
being developed, taking into consideration the
low-nutritional conditions of the water matrix
and the use of the MF technique. In general,
media supporting a broad range of fungal taxa
while restricting linear expansion of the fungal
colonies and simultaneously inhibiting bacterial
growth are preferred for the detection and enumeration of fungi in environmental samples.
Different media are used, according to the type of
water source and the aim of the fungal detection.
Nutritional characteristics are the most
influential factor in the suitability of a particular
medium for the recovery of water fungi. To detect
a wide range of fungal propagules present in the
water source, a nutrition-rich medium, with the
addition of an antibacterial agent, is used, such as
Sabourand dextrose agar (SDA) [33]. However,
low-nutrition media have also been recommended for fungal recovery from water [2].
Comparing poor and rich media, half-strength
corn meal agar (CMA/2) was recommended by
Kinsey et al. [2] for routine fungal examination
since it provided good results, with recovery of
higher fungal diversity, and is inexpensive. The
rich SDA medium supported higher fungal
counts, but mostly from common hypomycete
S. Masaphy
(Fungi Imperfecti) genera such as Penicillium
and Aspergillus, whereas CMA/2 supported other
genera [2].
Medium pH is also important. Generally, fungi
tend to grow in more acidic media than bacteria;
hence, many of the mycological media are
adjusted to be more acidic, thereby supporting
fungal growth while inhibiting bacterial growth.
This is even more relevant when fungi are being
detected in food products that may themselves be
acidic. Thus, due to adoption of media from the
food discipline, some officially recommended
media are already acidic, such as modified aureomycin-rose
bengal-glucose-peptone
agar
(MARGPA), which has a pH of 5.4 [34]. Other
recommended media, such as SDA, corn meal
medium, and dichloran-18% glycerol (DG18),
have pH values between 5.6 and 6 [35]. In 1962,
Mossel et al. [36] showed better recovery of
molds and yeasts from foods using media with
more neutral pH containing an antibacterial agent
than with media based on acid pH alone. In a
recent work in our lab [37], we also showed that
for recovery of fungi from a range of fungusinoculated water sources, rose bengal-chloramphenicol (RBC) medium with pH 7.2 was superior
to MARGPA.
Another important consideration in fungal
recovery is their rapid growth. Unlike most bacterial colonies, filamentous fungi tend to form
expansive colonies, which may cover small colonies of slower germinating or growing fungi and
yeast. Therefore, chemicals that inhibit hyphal
growth need to be added to limit overgrowth of
the fast-growing colonies. Dichloran and rose
bengal were shown to perform this function and
were incorporated into a nutrition-rich medium
to restrict the linear expansion of hyphal growth
[38–41]. In 1973, Jarvis [42] developed and used
rose bengal-chlortetracycline medium, and in
1979, King et al. [43] showed that introducing
dichloran and rose bengal together (with reduced
rose bengal concentration) allowed greater recovery of molds. More recently, DG18 has been recommended in water examinations and widely
used by Hageskal et al. [9, 10]. This medium was
developed for xerophilic fungi from foods [44],
14
Development of Media for Growth and Enumeration of Fungi from Water
and was recommended by Samson et al. [35] as a
general medium for the isolation and enumeration of fungi in food with water activity (aw) >90.
Askun et al. [45] compared DG18 and RBC
medium for fungal examination in raisins and
obtained higher fungal species diversity with
RBC, although both media gave similar results
for total fungal counts. We compared RBC
medium with MARGPA for the recovery of fungi
from different water sources and found RBC to
be superior.
As to the antibacterial agent, Korburger and
Rodgers [46] showed the positive effect of adding antibiotic to the medium on enumeration of
yeasts and molds, and today, as mentioned
above, media for detection of fungi in water
samples include a wide spectrum of different
antibiotics, such as chlortetracycline (auromycin) and streptomycin [34, 47], in addition to the
hyphal-restricting agent. We found that the antibiotic chloramphenicol is simplest to use as it is
autoclavable.
Assessing and Counting
The observed fungal colonies are counted and
referred to as CFU. There are two important
issues to consider. First, filamentous fungi tend to
spread over the medium, overlapping other slowgrowing fungi. As mentioned, to overcome this
problem, rose bengal or dichloran are incorporated into the medium. However, the concentration of the added compounds is important, as it
can limit fungal growth too severely. The second
issue involves observation of the fungal colonies
on the medium surface. Some of the fungi are
colored due to colored spores, whereas others
appear pale and are difficult to observe. Upon
using rose bengal to limit the overgrowth of fungi,
we found that it also strongly improves the colony count: the filamentous fungi and yeast colonies tend to absorb the rose bengal, giving them a
sharper color and reducing the need for optical
magnification [37]. This is especially true when
MF technique is used, as it is difficult to observe
light-colored colonies on the white filter.
205
Procedures
Media
Some of the more common media used for water
fungal detection and enumeration are presented
here.
1. Rose bengal-chloramphenicol (RBC) agar
(commercially available). Add 5 g peptone,
10 g glucose, 1 g K2HPO4, 0.5 g MgSO4·7H2O,
0.05 g rose bengal, and 15.5 g agar to 1 L distilled water. Heat to dissolve with stirring, and
autoclave. Adjust final pH to 7.2 ± 0.2.
2. Dichloran-RBC (DRBC) agar. Add 5 g peptone, 10 g glucose, 1 g K2HPO4, 0.5 g
MgSO4·7H2O, 0.002 g dichloran, 0.025 g rose
bengal, 0.1 g chloramphenicol, and 15 g agar
to 1 L distilled water. Adjust pH to 5.4–5.8.
3. Aureomycin-rose
bengal-glucose-peptone
agar (ARGPA). Add 5 g peptone, 10 g glucose, 1 g KH2PO4, 0.5 g MgSO4·7H2O, 0.035 g
rose bengal, and 20 g agar to 800 mL distilled
water. Prepare separately: dissolve 70 mg
chlorotetracycline (aureomycin hydrochloride) in 200 mL distilled water, filter-sterilize.
Add to the cooled (42–45 ºC) melted agar
medium. Adjust final pH to 5.4.
4. Modified ARGPA (MARGPA). Add 5 g peptone, 10 g glucose, 1 g KH2PO4, 0.5 g
MgSO4·7H2O, 0.035 g rose bengal, and 20 g
agar to 800 mL distilled water. Prepare separately: dissolve 200 mg chlorotetracycline in
200 mL distilled water and filter-sterilize. Add
to cooled (42–45 ºC) melted agar medium.
Adjust final pH to 5.4.
5. Dichloran-18% glycerol agar (DG18). Add
5 g peptone, 10 g glucose, 1 g K2HPO4, 0.5 g
MgSO4·7H2O, 1 mL dichloran (0.2% in ethanol), 220 g glycerol, 0.1 g chloramphenicol,
and 15 g agar to 1 L distilled water. Adjust to
pH 5.4–5.8.
6. Neopeptone-glucose-rose bengal-auromycin.
Add 5 g neopeptone, 10 g glucose, 3.5 mL
rose bengal solution (1 g/100 mL), and 20 g
agar to 1 L distilled water. Separately filtersterilize chlorotetracycline or tetracycline
206
S. Masaphy
(1 g/150 mL water). Add 5 mL of this solution
to 1 L agar solution immediately before use.
Adjust to pH 6.5.
7. Czapek Dox (CZ) agar. This medium is recommended for Aspergillus, Penicillium, and
similar fungi, among others, but not for total
fungal recovery. Add 30 g saccharose, 3 g
NaNO3, 1 g K2HPO4, 0.5 g MgSO4·7H2O,
0.5 g KCl, 0.01 g FeSO4, 0.5 g KCl, and 15 g
agar to 1 L distilled water. Adjust to pH 7.3.
8. SDA. Add 10 g mycological peptone, 40 g
glucose, and 15 g agar to 1 L distilled water.
Adjust to pH 5.4–5.8.
If dehydrated commercial medium is being
used, preparation should be as per the manufacturer’s recommendations. Otherwise, weigh each
of the medium components into 1 L or 800 mL
sterile distilled water, heat while stirring on a hotplate to near boiling until the agar is dissolved
and the medium is homogeneous. Autoclave
(121 °C for 15 min). Cool medium to 45 °C, and
then pour into the plates. If autoclave-sensitive
antibiotic is to be used, add 200 mL of the filtersterilized antibiotic at relevant concentration to
800 mL cooled medium, mix by stirring, and then
pour into the plates. Keep at 45 °C, and adjust pH
as required with HCl or NaOH.
5. Screw-cap bottles for dilutions, 100–500 mL
volume
6. Borosilicate glass flasks, 250–1,000 mL
volume
7. Sterile pipettes, glass or plastic, of appropriate volumes
8. Graduated cylinder, 100–1,000 mL
9. Sterile L-shaped glass rod or plastic disposable spreader rod
10. Petri dishes (either 50 or 90 mm), sterile,
plastic
11. Forceps, with smooth tips, to handle filters
without damaging them
12. Membrane filtration unit: filter funnel manifolds and filter manifolds (47 mm)
13. Membrane filter: 0.45-mm pore size white
hydrophobic mixed cellulose acetate membrane filter, grid marked, 47 mm, preferably
presterilized
14. Water bath maintained at 50 °C for tempering
agar medium
15. Incubator maintained at 15, 20, or 25 °C,
90% relative humidity
16. Vortex
17. Heating stirrer
18. Colony counter with magnifying glass
Materials
Techniques
1. Sterile distilled water
2. Ethanol or methanol in wide-mouth container for flame-sterilization of the forceps
3. 0.1 M NaOH and 0.1 M HCl
4. Dilution buffer composed of:
(a) Potassium
dihydrogen
phosphate
(KH2PO4) solution: Weigh 17 g KH2PO4
into 250 mL sterile water. Mix to complete
dissolution. Adjust to pH 7.2 with 1 N
NaOH, bring to 500 mL with sterile water.
(b) MgCl2 solution: Weigh 40.55 g of
MgCl2 · 6H2O into 500 mL sterile water.
Mix to complete dissolution.
(c) Working dilution buffer: Transfer
1.25 mL from solution (a) and 5 mL from
solution (b) into 1 L sterile distilled
water. Sterilize solution before use.
Fungi tend to spread unevenly in water bodies. It
is therefore important to mix the water in the
sample bottle vigorously prior to examination.
When using a low volume of sample, it is recommended that several repeats (optimally five) be
examined [34].
Spread Plate Technique
Streak 0.1–0.05 mL water sample onto the center
of pre-solidified agar medium (10–20 mL
medium) in a 90-mm Petri dish using a sterile
pipette. Spread the sample with a spreading rod
(available as disposable plastic rods, but glass
rods, which can be ethanol-sterilized, are preferred). Streak back and forth across the plate,
working up and down several times to distribute
the fungal units as evenly as possible. Use a single
14
Development of Media for Growth and Enumeration of Fungi from Water
rod per sample. Cover the plate and wait several
minutes before inverting it and incubating.
Pour Plate Technique
Water sample (usually 1 mL, but as little as
0.1 mL can be used) is added to an empty 90-mm
Petri dish. Then, 10–15 mL of molten propionate
medium is poured at 45 °C. The medium should
not be poured directly on the water sample. Mix
water sample and medium and let the mixture
solidify. In some cases, antibiotics may be added
to the Petri dish as well before pouring the molten medium. Incubate plates noninverted.
207
it is recommended that the results be read after
defined incubation periods. The SMEWW [34]
recommends incubation of spread plates at 15 °C
for 7 days, or 20 °C for 5–7 days. Slow-growing
fungi may not produce noticeable colonies until 6
or 7 days of incubation. For the pour plate method,
it is recommended that plates be incubated at
20–24 °C for 3–7 days. For the MF technique, the
recommendation is 15 °C for 5 days or 20 °C for
3–7 days. In all cases, plates should be incubated
in a humid (90–95% RH) atmosphere.
Counting
MF Technique
Shake the sample bottle to distribute the fungal
units uniformly. Filter the sample (10–1,000 mL,
ideally 100 mL) through the MF. Rinse the sides
of the funnel with 20–30 mL sterile dilution
water. Turn off the filtration system vacuum and
aseptically remove the MF from the filter base
using sterile forceps. Overlay the MF on the agar
medium surface in a 50-mm Petri dish. Close the
dish and incubate, either inverted or noninverted.
If the fungal counts exceed 80CFU/filtered volume, lower volumes of water should be used, or
the water source can be diluted 1:10 with dilution
water and filtered.
Incubation Conditions
Petri dishes are either incubated inverted or noninverted. Incubating the plate in the inverted position prevents dripping of the condensed water
onto the agar medium surface. Incubation is performed in the dark to avoid overproduction of
conidia that might spread and recontaminate the
agar medium surface. In some cases, however,
incubation in the light, but not direct sunlight, is
preferred to increase (colored) conidiation. The
incubation temperature is usually relatively low
(15–20 °C) to avoid overgrowth by fast-growing
fungi, allowing the slower fungi to germinate and
grow as well. In this case, a longer incubation may
be needed to recover higher counts. It is recommended that the plate be observed as soon as possible. However, for the purpose of standardization,
The count can provide an estimate of cultivable
fungi extracted from the water sample. All
filamentous fungi and yeast colonies may be
counted together or separately. The number of
fungi in the water sample is calculated as CFU/
mL of water. The calculation should take into
account the dilution factor. If counting cannot be
performed immediately, the culture plate can be
kept at 4 °C for 24 h. It is advisable to count colonies in plates that have the optimal number of
20–150 colonies per 90-mm plate for the spread
plate technique, and up to 300 colonies for the
pour plate technique. For the MF technique, it is
suggested that a magnifying binocular microscope be used to count all of the colonies, which
may be hard to see on the white filter background.
Ideal plates for counting should have 20–80 colonies per filter.
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37:959–964
Sabouraud Agar for Fungal Growth
15
Janelle M. Hare
Abstract
This article describes the history, theory, and use of Sabouraud agar for
isolating and growing fungi. This includes an explanation of the role of
each ingredient, the instructions for making this medium, variations upon
the basic recipe, and the various recipes that are commercially available.
Keywords
Sabouraud agar • Fungi • Growth • Dermatophytes • Sabouraud agar
(Modified) • Sabouraud agar Emmons • Glucose
History
Introduction
Sabouraud agar is one of the oldest and most
commonly used media for isolating and growing
fungi. It is effective in selectively isolating fungi
from environmental samples such as air and soil,
maintaining pure fungal cultures, and growing
fungi to distinguish and identify different species,
especially dermatophytes, by color and appearance. This article describes the theory behind the
use of Sabouraud agar, the role of the ingredients
in the medium, the preparation and use of the
medium, observations about the variation in
names and ingredients that can prove a source of
confusion, as well as visual results in the growth.
J.M. Hare (*)
Department of Biology & Chemistry,
Morehead State University, 150 University Blvd,
Morehead, KY 40351, USA
e-mail: jm.hare@moreheadstate.edu
Sabouraud agar medium was developed by the
French dermatologist Raymond J. A. Sabouraud
(pronounced sah-bū-rō¢) in the late 1800s to
support the growth of fungi, particularly dermatophytes [1, 2]. Sabouraud’s medical investigations focused on bacteria and fungi that cause
skin lesions, and he developed many agars and
techniques to culture pathogens such as dermatophytes and Malassezia. The long incubation period (multiple weeks) of dermatophytes
and the need to avoid bacterial contamination
while culturing them was the driving force
behind the development of this medium.
Sabouraud also sought to provide a medium that
would yield reliable results for fungal
identification across laboratories. He recommended that all mycologists detail their exact
media formulations and sources of ingredients as
well as the temperatures and times of specimen
incubation, in order to standardize observations
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_15, © Springer Science+Business Media, LLC 2013
211
212
and reduce media-derived sources of differences
in appearance [3].
Ironically, given Sabouraud’s original desire
to standardize the construction of fungal media,
there are currently many sources of confusion
and variation in both the names and ingredients
associated with Sabouraud agar, also called
Sabouraud’s agar (abbreviated either SDA or
SAB). Because of the old-fashioned use of the
term “dextrose” to refer to d-glucose, the medium
has been referred to as Sabouraud dextrose agar
as well as Sabouraud glucose agar, the term most
appropriate and consistent with standard chemical nomenclature [3]. Finally, a more recent
modification of Sabouraud agar by Emmons is
called either Sabouraud agar (Modified), or
Sabouraud agar, Emmons [4]. Many of the historical details behind these names and ingredient
variations are described in Odds’ excellent review
article [3].
Theory
Sabouraud agar is a selective medium that is formulated to allow growth of fungi and inhibit the
growth of bacteria. The available means of inhibiting bacterial growth in Sabouraud’s pre-antibiotic era was an acidic medium (pH 5.6). Currently,
the addition of antibiotics or antimicrobials to the
acidic medium is used to inhibit bacterial growth
(and sometimes saprophytic fungi, depending on
the particular antimicrobial used).
Sabouraud agar medium is complex and
undefined, but contains few ingredients. Peptones,
as soluble protein digests, are sources of nitrogenous growth factors that can vary significantly
according to the particular protein source. The
most variation is present in the source and method
of these protein digests. Both Difco and BBL
brand Sabouraud agars use pancreatic digests of
casein as their peptone source, but they and other
vendors also use a combination of pancreatic
digest of casein and peptic digest of animal tissues. Sabouraud’s original formulation contained
a peptone termed “Granulée de Chassaing,”
which is no longer available. Mold morphology
can vary slightly based on the peptones used, but
J.M. Hare
pigmentation and sporulation can be consistent if
one uses a consistent method of medium prepared
with the ingredients from the same source each
time. Researchers should also explicitly describe
the commercial or laboratory-prepared ingredients used in their medium.
Sabouraud originally used the sugar maltose as
an energy source, and although this medium is
still commercially available, glucose (formerly
referred to as dextrose) is currently used most frequently. Glucose is present at the high level of 4%
in Sabouraud’s formulation to assist in vigorous
fermentation and acid production by any bacteria
present, inhibiting later bacterial growth [5].
In 1977, Emmons formulated an alternative
version of Sabouraud’s agar, which contains half
the amount of glucose (2%) and a neutral pH of
6.8–7.0. The neutral pH seems to enhance the
growth of some pathogenic fungi, such as dermatophytes. Agar concentrations ranging from
1.5% to 2.0% are found in commercial preparations of Sabouraud agar in both the original formula and Emmons modification, and serve to
solidify the medium in tube and plate medium.
Materials
Sabouraud agar can be either made from individual ingredients (Tables 15.1 and 15.2), purchased
either as dehydrated powder that must be dissolved
in water, autoclaved, and dispensed, or as prepared
medium that can be purchased in tube, plate, or
broth form from a variety of commercial sources
such as Becton Dickinson/Difco, Remel, or BBL.
Various antimicrobials can be added to either the
original recipe (Sabouraud agar), or Sabouraud
agar, Modified/Emmons (see Table 15.2).
1. Deionized, distilled water.
2. Autoclave.
3. Graduated cylinder, 1,000 mL.
4. Erlenmeyer flask (2 L if making 1 L of
medium).
5. Analytical balance (if using antimicrobial
agents).
6. Balance for weighing media ingredients.
7. Stir bar.
8. Stirring hotplate.
15
Sabouraud Agar for Fungal Growth
213
Table 15.1 Ingredients
for Sabouraud agar and
Sabouraud agar (Emmons)
Ingredient
Pancreatic digest of casein
Glucose
Agar
Table 15.2 Antimicrobial
and other additives to
Sabouraud agar
Ingredient
Chloramphenicola
Cycloheximidea
Gentamicin
Lecithin
Tween 80
Olive oil
Sabouraud agar (g/L)
10
40
15–20
Amount (per liter)
50 mg
0.5 g
50 mg
0.7 g
5g
Sabouraud agar (Emmons) (g/L)
10
20
15–20
Notes
Dissolve in 10 mL 95% ethanol
Dissolve in 2 mL acetone
Dissolve in 5 mL water
Add directly with other powdered
medium ingredients before autoclaving
Spread 0.1 mL sterile olive oil on
surface of plate
Add these to molten, autoclaved media once it has been tempered in water bath to
45–50 °C
a
9. Slant tube rack for holding media tubes after
autoclaving to solidify with a slanted
surface.
10. Pancreatic digest of casein.
11. Glucose.
12. Chloramphenicol.
13. Gentamicin sulfate.
14. Cycloheximide.
15. Tween 80 (polysorbate 80).
16. Lecithin.
17. Olive oil, sterilized by autoclaving.
18. Sterile glass test tubes with caps.
19. Sterile Petri dishes, 100 mm diameter.
hand). If the antimicrobials chloramphenicol
or cycloheximide are to be added, aseptically
add them at this point and swirl medium gently
(see Variations on Standard Sabouraud Agar).
6. Pour into Petri dishes or tubes and leave at
room temperature overnight to solidify and
dry. When pouring plates, fill each Petri dish
with at least 25 mL of medium to allow for
medium dehydration during the longer incubation period required for fungi. If preparing
tubes, slant the rack of covered tubes immediately after pouring in a slant tube rack, either
at a 5° or 20° slant.
7. Store all media at 4 °C, regardless of whether
they contain antimicrobials (see Table 15.1).
Method of Sabouraud Agar
Preparation
1. Combine all ingredients, except any antimicrobials to be used, in ~900 mL of deionized
water in a graduated cylinder while stirring
with a magnetic stir bar.
2. Adjust to pH 5.6 with hydrochloric acid and
adjust final volume to 1 L.
3. Transfer contents to a 2-L flask and boil on a
heating/stirring plate while stirring, for 1 min.
4. Cover opening of flask loosely with aluminum
foil and autoclave 15 min at 121 °C under
pressure of 15 lb/in2.
5. Cool to ~45–50 °C (roughly until one can support the flask underneath with an ungloved
Method of Sabouraud Agar, Emmons
Modification Preparation
1. Combine all ingredients, except any antimicrobials to be used, in ~900 mL of deionized
water in a graduated cylinder while stirring
with a magnetic stir bar.
2. Adjust to pH 6.8–7.0 with hydrochloric acid
and adjust final volume to 1 L.
3. Transfer contents to a 2-L flask and boil on a
heating/stirring plate while stirring, for 1 min.
4. Cover opening of flask loosely with aluminum
foil and autoclave 15 min at 121 °C under
pressure of 15 lb/in2.
214
5. Cool to ~45–50 °C (roughly until one can support the flask underneath with an ungloved
hand). If the antimicrobials chloramphenicol
or cycloheximide are to be added, aseptically
add them at this point and swirl medium gently
(see Variations on Standard Sabouraud Agar).
6. Pour into Petri dishes or tubes and leave at
room temperature overnight to solidify and
dry. Fill each Petri dish with at least 25 mL of
medium to allow for medium dehydration during the longer incubation period required for
fungi. If preparing tubes, slant the rack of covered tubes immediately after pouring in a slant
tube rack, either at a 5° or 20° slant.
7. Store all media at 4 °C, regardless of whether
they contain antimicrobials.
Variations on Standard Sabouraud Agar
Either Sabouraud agar or its Emmons version can
be made more selective by adding antimicrobials
(see Table 15.2). Antimicrobials commonly used
are the aminoglycoside gentamicin, which inhibits
gram-negative bacteria; chloramphenicol, which
inhibits a wide range of gram-positives and gramnegatives; and/or cycloheximide, which inhibits
primarily saprophytic fungi, but not dermatophytes
or yeasts [6, 7]. Chloramphenicol and gentamicin
are used at 50 mg/L (50 mg of chloramphenicol
dissolved in 10 mL of 95% ethanol before adding
to molten medium) and cycloheximide at 0.5 g/L
(0.5 g dissolved in 2 mL of acetone before adding
to molten medium) [8]. Chloramphenicol and
cycloheximide should only be added after media
has been autoclaved and then cooled to ~45–50 °C
(see step 5 in Method of Sabouraud Agar
Preparation). Gentamicin may be added to the
medium ingredients before autoclaving.
Lecithin and Tween 80 are added to Sabouraud
agar (see Table 15.2) that is used in monitoring
environmental surfaces that may have been
treated with antiseptics and quaternary ammonium compounds, as these additives neutralize
the cleaning compounds [9]. Sterile olive oil can
be spread on the surface of Sabouraud agar plates
to grow lipophilic Malassezia species [10] (see
Table 15.2).
J.M. Hare
Methods of Inoculation and Incubation
Sabouraud agar plates can be inoculated by streaking for isolation, as with standard bacteriological
media, by exposing the medium to ambient air, or
by tamping clinical sample material (hair, skin
scrapings, etc.) onto the surface of the agar
medium. When growing cultures in tubes, the
caps should be screwed on loosely to admit air, as
dermatophytes and most molds are obligate aerobes. Isolation of fungi is performed on plates,
while slants are primarily used for maintaining
pure, or stock, cultures once isolated. If using
selective Sabouraud media, a control plate/tube
without antimicrobials should also be inoculated
for comparison. Typically, molds are incubated at
room temperature or slightly warmer (25–30 °C),
yeasts are incubated at 28–30 °C or both 30 and
37 °C if suspected to be dimorphic fungi.
Incubation times will vary, from approximately two days for the growth of yeast colonies
such as Malassezia, to 2–4 weeks for growth of
dermatophytes or dimorphic fungi such as
Histoplasma capsulatum. Indeed, the incubation
time required to acquire fungal growth is one
diagnostic indicator used to identify or confirm
fungal species. Dermatophytes in particular show
characteristic incubation times ranging from 5 to
7 days (some Epidermophyton or Microsporum
species) to 3–4 weeks for some Trichophyton
species [11]. Cultures should be examined twice
weekly and be held for 4–6 weeks before being
reported as negative if infection by systemic
agents such as Histoplasma, Blastomyces, or
Coccidioides is suspected.
Results
Depending on the antimicrobials used, different
types of microorganisms and groups of fungi may
grow on Sabouraud agar (Table 15.3). Typically,
saprophytic fungi are inhibited by cycloheximide
and/or chloramphenicol, but yeasts and dermatophytes grow well in their presence. Conversely,
even Sabouraud agar is unable to support the
growth of a few dermatophytes in the absence
of additives. For example, some Trichophyton
15
Sabouraud Agar for Fungal Growth
Table 15.3 Expected
growth of various microbes
on Sabouraud agar
containing antimicrobials
Microbe
Candida albicans
Cryptococcus neoformans
Aspergillus niger
Trichophyton mentagrophytes
Microsporum audouinii
Blastomyces dermatitidis
Histoplasma capsulatum
Rhizopus spp.
Sporothrix schenckii
Penicillium roqueforti
Escherichia coli
215
Growth on SAB + CAMa
Yes
Yes
Yes
Yes
Yes
Yes (mold phase at 25 °C)
No (yeast phase at 37 °C)
Yes (mold phase at 25 °C)
No (yeast phase at 37 °C)
Yes
Yes
Yes
No
Growth on SAB + CHXb
Yes
No
No
Yes
Yes
Yes (mold phase at 25 °C)
No (yeast phase at 37 °C)
Yes (mold phase at 25 °C)
No (yeast phase at 37 °C)
No
Yes
No
No
SAB + CAM = Sabouraud agar plus chloramphenicol
SAB + CHX = Sabouraud agar plus cycloheximide
a
b
species require additional growth factors, such as
thiamine and inositol (T. verrucosum) or nicotinic
acid (T. equinum), and may not grow well, if at
all, on Sabouraud agar [12]. T. mentagrophytes
and T. rubrum, however, grow well on Sabouraud
agar. Similarly, the growth of Malassezia species
is significantly impaired without the addition of
olive oil overlaid on the surface of a Sabouraud
agar plate [10] (see Table 15.3).
Mold morphology should be observed on both
the top (obverse) and bottom (reverse) surfaces,
as differences can be seen on each surface.
Variation from lot to lot as well as between
commercial vendors of Sabouraud agar can
significantly impact the qualitative and quantitative growth of fungi. One study comparing five
different commercial preparations of Sabouraud
glucose agar observed significant differences in
the quantitation of yeasts as well as the color of
Aspergillus colonies; however, the dermatophytes
yielded reliably similar appearances on the five
media sources tested [13].
Notes
Fungi often produce spores that are easily dispersed into the laboratory upon opening of plates.
Plates should be incubated with the lid on the top
(as opposed to the typical practice of inverting
microbiological plates for incubation) to avoid
spreading spores when the plates are opened.
After growth, plates should be wrapped in
Parafilm to maintain them securely closed for
storage and/or transport. Plate or tube cultures
should be opened only within a class II biological
safety cabinet to avoid contamination of laboratory spaces with fungal spores, possible infection
of individuals by pathogenic fungi, or induction
of allergic responses.
Because the growth of large numbers of fungi
can pose a potential infection hazard, measures
must also be taken to prevent infection of laboratory researchers. Note that some fungi are biosafety level one (BSL-1), whereas most are
BSL-2 [14]. The American Society for
Microbiology strongly recommends that environmental enrichment experiments should only be
performed in BSL-2 laboratories. The following
precautions apply to the use of Sabouraud agar:
1. Direct environmental samples (e.g., soil,
water) that are known to contain infectious
organisms should be handled according to the
biosafety level of that infectious agent.
2. Cultures of enriched microorganisms derived
from environmental samples should be handled using BSL-2 precautions.
3. Mixed, enriched, or pure cultures of microorganisms from environmental samples with a
significant probability of containing infectious
agents should be manipulated in a class II biosafety cabinet, if available.
216
4. Researchers should be aware if they work in
regions with endemic fungi capable of causing
systemic infections, and should avoid environmental isolations. Some safe fungi for student
experimentation and handling include the
molds Penicillium camemberti and P. roqueforti (used in making cheeses), Rhizopus stolonifer (used in making tempeh), Aspergillus
species (except A. fumigatus and A. flavus), the
yeasts Saccharomyces cerevisiae, Rhodotorula
rubrum, and Neurospora crassa.
Acknowledgements I thank my colleague Ted Pass for
helpful comments on this chapter.
References
1. Sabouraud R (1896) La question des teignes. Ann
Dermatol Venereol (series 3) 7:87–135
2. Sabouraud R (1896) Recherche des milieux de culture
propres a la différenciation des espèces trichophytiques a grosse spore. In: Les trichophyties humaines.
Masson et Cie, Paris, pp 49–55
3. Odds FC (1991) Sabouraud(‘s) agar. J Med Vet Mycol
29:355–359
4. Emmons CW, Binford CH, Utz JP, Kwon-Chung KJ
(1977) Culture media. In: Medical mycology, 3rd edn.
Lea & Febiger, Philadelphia, p 535
J.M. Hare
5. Jarrett L, Sonnenwirth AC (1980) Gradwohl’s and
parasitic infections, 7th edn. American Public Health
Association, Washington, DC
6. McDonough ES, Ajello Georg LK, Brinkman S
(1960) In vitro effects of antibiotics on yeast phase of
Blastomyces dermatitidis and other fungi. J Lab Clin
Med 55:116
7. Lorian V (2005) Antibiotics in laboratory medicine.
Lippincott, Williams & Wilkins, Baltimore
8. Hungerford LL, Campbell CL, Smith AR (1998)
Veterinary mycology laboratory manual. Iowa State
University Press, Ames
9. Curry AS, Graf JG, McEwen GN Jr (1993) CTFA
microbiology guidelines. The Cosmetic, Toiletry and
Fragrance Association, Washington, DC
10. Kwon-Chung KJ, Bennett JE (1992) Infections caused
by Malassezia species. In: Medical mycology. Lea &
Febiger, Philadelphia, pp 70–182
11. Robert R, Pihet M (2008) Conventional methods for
the diagnosis of dermatophytosis. Mycopathologia
166:295–306
12. Georg LK, Camp LB (1957) Routine nutritional tests
for the identification of dermatophytes. J Bacteriol
74:113–121
13. Brun S, Bouchara JP, Bocquel A, Basile AM, ContetAudonneau N, Chabasse D (2001) Evaluation of five
commercial Sabouraud gentamicin-chloramphenicol
agar media. Eur J Clin Microbiol Infect Dis
20:718–723
14. Centers for Disease Control and Prevention. Biosafety
in microbiological and biomedical laboratories
(BMBL). 5th edn. Section VIII-B: Fungal agents, pp
170–181. Available from: http://www.cdc.gov/biosafety/publications/bmbl5/BMBL.pdf
A Method for the Formation
of Candida Biofilms in 96 Well
Microtiter Plates and Its Application
to Antifungal Susceptibility Testing
16
Christopher G. Pierce, Priya Uppuluri,
and Jose L. Lopez-Ribot
Abstract
Fungal infections are an increasing threat to an expanding population of
immunocompromised patients. Of these, candidiasis remains the most
common, now representing the third to fourth most prevalent infection in
US hospitals. Candida albicans remains the major causative agent of candidiasis. Most manifestations of candidiasis are associated with biofilm
formation on either host tissues or implanted biomaterials (i.e., catheters),
which carries important negative consequences, as cells within biofilms
show dramatically increased levels of antifungal drug resistance and protection from host defenses. Here we describe a rapid and robust model for
the formation of C. albicans biofilms in vitro using 96 well microtiter
plates, which can also be easily adapted for antifungal susceptibility testing. The read-out is colorimetric, based on the reduction of a tetrazolium
salt (XTT) by metabolically active cells. This method simplifies biofilm
formation, democratizes biofilm research, and provides a framework for
the standardization of antifungal susceptibility testing of fungal biofilms.
Keywords
Candida • Candidiasis • Biofilms • Antifungals • Fungi • Susceptibility
testing
Introduction
C.G. Pierce • P. Uppuluri • J.L. Lopez-Ribot (*)
Department of Biology, South Texas Center for
Emerging Infectious Diseases, The University of Texas
at San Antonio One UTSA Circle, San Antonio,
TX 78249, USA
e-mail: jose.lopezribot@utsa.edu
Since microorganisms can be diluted to a single
cell and studied in pure culture, most investigations
in the field of microbiology have typically involved
the use of free-living (planktonic) cells in liquid
cultures, leading to the “dogmatic” and almost universal consideration of microorganisms as unicellular life forms. However, during the last few
decades there has been an increasing recognition of
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_16, © Springer Science+Business Media, LLC 2013
217
218
the role that microbial biofilms play both in
nature and during disease [1, 2]. Biofilms are
defined as structured microbial communities that
are attached to a surface and encapsulated within
a self-produced matrix. Biofilm formation carries
important negative clinical consequences,
because cells in biofilms, in stark contrast with
their free-floating planktonic counterparts, are
notoriously resistant to most antibiotics [3].
Candidiasis remains the most common fungal
infection in hospitalized patients, now representing the third to fourth most frequent nosocomial
infection worldwide [4–8]. C. albicans remains
the main etiologic agent of candidiasis, although
other species are on the rise [9]. The increase in
the frequency of candidiasis in the last few
decades is associated with the increase in use of a
variety of medical implant devices in which
Candida can form biofilms. These include, among
others, different types of catheters (i.e., urinary,
intravascular), endotracheal tubes, intracardiac
devices, neurosurgical shunts, and prosthetic
joints [10, 11]. Other manifestations of candidiasis, such as denture stomatitis and oropharyngeal
candidiasis, also have a biofilm etiology [12].
From the clinical point of view, the most
salient feature of Candida biofilms is their high
levels of resistance against conventional antifungal agents, particularly azoles and polyenes
[13–15]. However, newer antifungal agents,
such as the echinocandins and liposomal formulations of amphotericin B, display increased
efficacy against fungal biofilms [16–18]. Also,
biofilms provide a safe haven where microorganisms are protected from host immune defenses
and from which they can disseminate to colonize
and infect distal sites [11]. In other instances,
biofilm formation is directly responsible for failure of a contaminated device, very often necessitating removal. It is important to note that the
commonly used Clinical and Laboratory
Standards Institute broth microdilution techniques for antifungal susceptibility testing are
based on the use of planktonic populations and
will not enable prediction of the drugs’ efficacy
against fungal biofilms, which underscores the
importance of developing standardized techniques for biofilm formation and to determine
C.G. Pierce et al.
the effectiveness of different antifungal agents
and regimens against fungal biofilms.
Traditionally, most models for the formation
of fungal biofilms are cumbersome, requiring the
use of specialized equipment, expert handling,
and long processing times. Moreover, since relatively few equivalent biofilms can be produced at
the same time, the majority of these methods do
not allow for high throughput screening [19].
Here, we describe a fast and highly reproducible
method for the formation of multiple equivalent
biofilms on the bottom of wells of microtiter
plates, coupled with a colorimetric method that
measures the metabolic activity of cells within
the biofilm based on the reduction of 2,3-bis(2methoxy-4-nitro-5-sulfo-phenyl)-2H-tetrazolium-5-carboxanilide (XTT). Although the
method was initially developed for C. albicans
and other Candida spp., it can also be easily
adapted for other biofilm-forming fungal species
of clinical interest, such as Cryptococcus neoformans and Aspergillus fumigatus [20]. The method
can be used to examine multiple parameters and
factors influencing biofilm formation, to estimate
the biofilm-forming ability of multiple fungal
isolates and mutant strains, and to perform antifungal susceptibility testing of fungal biofilms.
Materials
1. Clinical or laboratory strains of C. albicans
(or other fungal species).
2. Yeast peptone dextrose (YPD) (1% w/v yeast
extract, 2% w/v peptone, 2% w/v dextrose,
1.5% agar) or Sabouraud-dextrose agar plates
or slants for subculturing Candida isolates.
3. YPD liquid medium (1% w/v yeast extract,
2% w/v peptone, 2% w/v dextrose).
4. RPMI-1640 without sodium bicarbonate
supplemented with l-glutamine and buffered
with 165 mM morpholinepropanesulfonic
acid. From now on this medium will be
referred to simply as RPMI 1640.
5. Sterile phosphate buffered saline, PBS
(10 mM phosphate buffer, 2.7 mM potassium
chloride, 137 mM sodium chloride, pH 7.4)
(Sigma-Aldrich, St Louis, MO, USA).
16
A Method for the Formation of Candida Biofilms…
219
6. Haemocytometer
(Hausser
Scientific,
Horsham, PA, USA).
7. Polystyrene, flat-bottomed, 96 well microtiter plates.
8. 2,3-bis(2-methoxy-4-nitro-5-sulfo-phenyl)2H-tetrazolium-5-carboxanilide (XTT). The
XTT saturated solution is prepared at 0.5 g/L
in sterile Ringer’s lactate or PBS. This solution is light sensitive, so it should be covered
with aluminum foil during preparation. The
solution is filter-sterilized using a 0.22-mmpore size filter (the filtration step will leave
yellow residues on the filter, but this does not
constitute a problem). Aliquot into 10 mL
working volumes, and store at −70 °C,
wrapped in aluminum foil.
9. Menadione, prepared as a 10-mM stock solution in 100% acetone. Aliquot into smaller
volumes (about 50 mL) and store at −70 °C.
10. Vortex mixer.
11. Microtiter plate reader with 490-nm optical
filter.
12. Bright field inverted microscope.
13. Multichannel pipette.
14. Antifungal drugs. Most drugs are initially
solubilized in DMSO, but some antifungals
are also soluble in water. If needed, concentrated stock solutions (i.e., 1 mg/mL) of the
antifungals can be aliquoted into smaller volumes and stored at −70 °C until required.
2. Flasks containing YPD liquid medium
(typically 20 mL of medium in a 150-mL
flask) are inoculated with a loopful of cells
from the stock cultures and incubated overnight in an orbital shaker (150–180 rpm) at
30 °C. C. albicans should grow in the budding–yeast form under these conditions (check
under the microscope).
3. Centrifuge the liquid cultures (approximately
3,000 g for 5 min at 4 °C), remove supernatant, and wash twice in sterile PBS (by resuspending the pellet in approximately 20 mL of
ice-cold buffer, vortexing vigorously, followed
by centrifugation as described previously).
4. Resuspend the final pellet of cells in approximately 20 mL of RPMI 1640 medium that has
been prewarmed to 37 °C.
5. Prepare 1:100 and/or 1:1,000-fold dilutions in
the same medium and count using a haemocytometer on a bright field microscope with a
40× objective lens. Calculate the volumes
needed to prepare a suspension of cells at a
final density of 1.0 × 106 cells/mL in RPMI
1640. The total volume needed will depend on
the total number of wells (or plates) that need
to be seeded for biofilm formation.
6. Use as many 96 well microtiter plates as
needed according to the experimental design.
We recommend performing a minimum of
2–4 replicates (entire rows or columns of the
microtiter plate[s]) for each condition (i.e.,
isolate, strain, antifungal concentration) to be
tested. Pipette 100 mL of the standardized
inoculum into selected wells of the microtiter
plate(s). Ideally, leave wells in column 12 on
each plate empty, and use these wells as negative background controls during subsequent
analyses. If multiple rows in the same plate, or
the entire plate, or if multiple plates are to be
seeded with the same fungal isolate, the use of
a multichannel pipette is strongly recommended for this and successive steps.
7. After initial seeding, cover the microtiter plate
with its original lid, seal with parafilm, and
place inside an incubator. Incubate statically
for 24–48 h at 37 °C.
8. After biofilm formation, use a multichannel
pipette to aspirate the medium carefully so as
not to touch and disrupt the biofilm. Wash the
Methods
The methods described below summarize the formation of C. albicans biofilms and procedures
for antifungal susceptibility testing of cells within
biofilms.
Formation of Candida Biofilms on 96
Well Microtiter Plates (Fig. 16.1)
1. Candida isolates are typically stored as glycerol stocks. From these stocks (or from a fresh
culture if a recent clinical isolate) streak a
loopful of cells onto a plate containing YPD
agar or Sabouraud-dextrose agar and incubate
overnight at 37 °C.
220
C.G. Pierce et al.
Fig. 16.1 Schematic diagram of the
protocol for the formation of C. albicans
biofilms on 96 well microtiter plates
plates (using a multichannel pipette or an
automatic plate washer) three times in sterile
PBS (200–300 mL per well) to remove nonadherent cells that remain in the wells. After
each wash the microtiter plates should be
drained in an inverted position by blotting
with paper towels to remove any residual PBS.
Biofilms are now ready to be processed for
XTT (to determine the extent of biofilm formation) or alternatively to be treated with
antifungals for susceptibility testing.
At this point after biofilm formation, if the
main purpose of a particular set of experiments is
to estimate the biofilm-forming ability, the extent
of biofilm formation, or to determine biofilmforming kinetics, the XTT/menadione reagent
can be added and the resulting color read using a
microtiter plate reader.
Methods for Antifungal Susceptibility
Testing Against Candida Biofilms
on 96 Well Microtiter Plates
The following steps describe the preparation of
antifungal agents for testing and the challenging
of pre-formed biofilms with antifungal agents.
1. From stock solutions or powder prepare a final
“high” working solution in RPMI 1640
medium of each antifungal to be tested. Typical
high concentrations are 1,024 mg/mL for
fluconazole, and 16 mg/mL for both amphotericin B and caspofungin. Other concentrations can be used for different agents.
2. Using a multichannel pipette, add 200 mL of
the high working concentration of antifungal
to the corresponding wells on column 1 of each
microtiter plate containing the C. albicans
16
3.
4.
5.
6.
A Method for the Formation of Candida Biofilms…
221
biofilms. Be careful not to touch or otherwise
disrupt the biofilms.
Add 100 mL of RPMI 1640 to each well in
columns 2–11.
Remove 100 mL of antifungal agent from the
wells of column 1 and add to the adjacent
wells in column 2 (already containing 100 mL
of medium). Mix the contents well by pipetting
up and down and remove the tips.
Repeat moving right until the wells of column
10, after which the final 100 mL volume from
the wells of column 10 after mixing is discarded. In this way, a series of doubling dilutions of your agent(s) of interest have been
created; from most concentrated in wells of
column 1 to least concentrated in wells of column 10. Unchallenged biofilms in column 11
will serve as positive controls, and empty
wells in column 12 will serve as negative
controls.
Cover the plates with their lids, seal with
parafilm, and incubate for 24–48 h at 37 °C.
Other incubation times/conditions may be
used depending on the experimental design.
of the stock solution of menadione to achieve
a final menadione concentration of 1 mM (for
uniformity, if multiple plates are processed at
the same time, we recommend pooling all the
resulting XTT/menadione tubes into a single
solution in a clean sterile container).
Using a multichannel pipette add 100 mL of
XTT/menadione solution to each well containing a pre-washed biofilm as well as to
negative control wells for the measurement of
background XTT-colorimetric levels.
Cover the plates in aluminum foil and incubate in the dark for 1–2 h at 37 °C.
Uncover the plates. Using a multichannel
pipette remove 80 mL of the resulting colored
supernatant from each well and transfer into
the corresponding wells of a new microtiter
plate, and read the plate(s) in a microtiter plate
reader at 490 nm.
From the resulting colorimetric readings (and
after subtracting the corresponding values for
negative controls from wells in column 12
containing XTT only), calculate the sessile
minimum inhibitory concentrations SMIC50
and SMIC80, which are the antifungal concentrations at which a 50 or 80% decrease
in XTT readings are detected in comparison
to the control biofilms formed in the absence
of antifungal drug (in this case, values for
column 11).
Use of the XTT-reduction Assay to
Estimate Fungal Cell Viability After
Treatment with Antifungals
The XTT assay relies on the measurement of
metabolic activity of cells within the biofilm
based on the reduction of XTT, which yields a
water-soluble formazan-colored product that can
be measured using a microtiter plate reader. We
(and others) have previously shown that the XTTreduction assay shows excellent correlation
between cellular density and metabolic activity,
thus providing a semiquantitative measurement
of biofilm formation [19, 21]. The XTT assay is
nondestructive, and requires minimal post-processing of samples in stark contrast with other
methods (i.e., cell counts).
1. After the incubation period wash the plates 3
times with PBS.
2. Thaw as many tubes containing 10 mL of the
XTT solution as required for the experimental
design (one per plate). To each tube, add 1 mL
3.
4.
5.
6.
Notes
1. Always use proper microbiological handling
techniques and universal precautions to handle
microorganisms.
Follow
Institutional
Guidelines Regarding Biosafety.
2. Please note that the seeding densities provided
here have been optimized for the formation of
biofilms on wells of microtiter plates, as quorum-sensing mechanisms play an important
role in biofilm formation. Cell densities that
are too high or too low will likely result in
poor biofilms.
3. After initial seeding of the plates the time of
incubation for biofilm formation may be varied depending on the specific objectives of the
222
4.
5.
6.
7.
study: for example, if the main objective of the
study is examination of initial adherence, the
incubation time can be reduced to 2–4 h, other
investigators may want to study “fully mature”
biofilms after 72–96 h incubation, or perhaps
study the kinetics of biofilm formation for
which multiple plates can be seeded at the
same time and then individual plates can be
processed after different incubation times (i.e.,
0, 2, 4, 6, 8, 12, 24, and 48 h).
There are multiple washing steps during the
entire protocol. It is critical to preserve biofilm
integrity during these washing steps. Normally
the biofilms are strongly attached to the bottom of the wells, and these washing procedures should not disrupt the preformed
biofilms. If at the end of the washing procedures wells are observed with clearly disrupted
biofilm layer at the bottom (normally this is
visible by the naked eye), then these wells
should be excluded from the analyses. This is
one of the main reasons why we recommend
performing sufficient replicates for each condition tested.
If all the methods have been followed properly, biofilms formed on the bottom of the
wells should be visible by the naked eye, simply by looking at the underside of the microtiter plate. In addition, an inverted microscope
can be used to examine morphological details
of the formed biofilms. Images can be captured if such microscope is equipped with a
camera(s) that would allow for image
acquisition.
If testing experimental and/or new agents with
unknown activity against biofilms, we recommend starting with high concentrations of the
drug, normally up to 100–1,000 times higher
than one would use against planktonic
populations.
SMIC results are typically presented in a tabular fashion (i.e., multiple antifungals against
multiple strains or isolates) or, alternatively,
results for each individual strain against each
antifungal can be presented as a graph by plotting percent inhibition (or colorimetric readings) versus antifungal concentration.
C.G. Pierce et al.
Acknowledgments Biofilm-related work in the laboratory has been funded by Public Health Service grants
numbered R21DE017294 and R21AI080930 from the
National Institute of Dental & Craniofacial Research and
the National Institute for Allergy and Infectious Diseases
(to Lopez-Ribot). Pierce is supported by a predoctoral fellowship, 51PRE30004, from the American Heart
Association. Uppuluri is supported by a postdoctoral fellowship, 10POST4280033, from the American Heart
Association. The content is solely the responsibility of the
authors and does not necessarily represent the official
views of the NIDCR, the NIAID, the NIH, or the AHA.
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Screening for Compounds Exerting
Antifungal Activities
17
Jean-Paul Ouedraogo, Ellen L. Lagendijk,
Cees A.M.J.J. van den Hondel, Arthur F.J. Ram,
and Vera Meyer
Abstract
There is a strong demand for the discovery of new antifungal drugs. More
and more human and plant pathogenic fungi develop resistance against
currently used drugs and therefore do not respond to antifungal treatments.
As humans and fungi are both eukaryotic cells in which many molecular
processes are conserved, compounds that have antifungal activity are also
often toxic for humans. To circumvent this, it is important to develop
methods and screens for the identification of compounds that specifically
kill fungi but do not affect men and the environment. In this chapter, we
describe methods to screen compounds for their ability to prevent growth
of the filamentous fungus Aspergillus niger, and to monitor whether these
compounds are fungicidal and whether they switch on the agsA reporter
system, which is representative for cell wall or cell membrane stress.
Keywords
Antifungal • Fungicide • Aspergillus • Cell wall • Cell membrane •
Susceptibility assay • Azoles • Polyenes • Echinocandins
Introduction
J.-P. Ouedraogo • V. Meyer
Department Applied and Molecular Microbiology, Berlin
University of Technology, Institute of Biotechnology,
Gustav-Meyer-Allee 25, Berlin 13355, Germany
E.L. Lagendijk • C.A.M.J.J. van den Hondel
• A.F.J. Ram (*)
Department of Molecular Microbiology
and Biotechnology, Leiden University, Sylviusweg 72,
Leiden 2333 BE, The Netherlands
e-mail: a.f.j.ram@biology.leidenuniv.nl
Both the plasma membrane and the cell wall of
fungi contain components that are unique to the
fungal kingdom. Hence, drugs that interfere with
the biosynthesis of these components are likely
to be fungal-specific. Azoles, polyenes, and
echinocandins are three groups of drugs that are
used nowadays to treat fungal infections [1].
Azoles inhibit ergosterol synthesis in fungi,
which is the cholesterol equivalent of animal
membranes. The four currently used azoles
include fluconazole, itraconazole, voriconazole,
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_17, © Springer Science+Business Media, LLC 2013
225
226
and posaconazole; these drugs block ergosterol
biosynthesis by inhibiting the activity of the
cytochrome P450 lanosterol demethylase [2].
Fenpropimorph is a morpholine fungicide that
also inhibits ergosterol biosynthesis and protects
plants against pathogenic fungi [3]. Polyenes are
amphiphatic drugs that strongly bind to ergosterol and create channels, thereby disrupting the
integrity of fungal membranes. The most often
used polyene, amphotericin B, is effective against
several pathogenic fungi; however, its use is
restricted because of detrimental side effects on
mammalian cells [1].
In addition to the cell membrane, fungal cells
are surrounded by a cell wall, which is essential
for the fungus to withstand the internal turgor
pressure. The fungal cell wall is composed of chitin, beta-glucans, and mannosylated proteins.
Depending of the fungal species, polysaccharides
(e.g., alpha-glucans, galactomannans) can be
present as well [4, 5]. Some of these components
are covalently linked and connected to each other
to ensure the rigidity and strength of the fungal
cell wall. Echinocandins are currently the only
class of antifungals that target the biosynthesis of
the fungal cell wall. The tree echinocandins used
in medicine (caspofungin, micafungin, and anidulafungin) inhibit the function of the (1,3)-betad-glucan synthase, which is an essential enzyme
for fungal cell wall biosynthesis [6].
Whether a fungal cell is directly killed by a drug
is dependent on a variety of factors. Both the concentration of the drug as well as the intrinsic resistance of the specific fungus are important factors
that determine drug sensitivity. At nonlethal concentrations, the drugs can trigger stress responses
that can make the fungal cell more resistant towards
the drugs. It has been well established that the addition of drugs that interfere with cell membrane or
cell wall biosynthesis trigger the cell wall stress
response pathway [7–9]. The pathway is partially
conserved in fungi such as the yeasts Saccharomyces
cerevisiae and Candida albicans and the
filamentous fungi such as Aspergillus fumigatus
and Aspergillus niger [10–12]. We have previously
shown that the induction of the agsA gene from
A. niger, which encodes a putative (1,3)-alpha-dglucan synthase, is a very suitable and specific
reporter to monitor fungal cell wall stress [7–11].
J.-P. Ouedraogo et al.
In the following sections, we describe microtiter- and microscopic-based methods to identify
compounds that are fungicidal and, moreover,
induce the agsA reporter. These methods allow
the set up of high-throughput approaches to
identify potential drugs that very specifically
disrupt fungal-specific mechanisms essential for
survival.
Materials
1. Glucose (50 %): For 1 L: Boil 500 mL
Milli-Q (MQ) in a 1,000-mL beaker on a
heated magnetic stirrer. Slowly add 500 g of
d(+)-Glucose anhydrous. After glucose has
been dissolved, let the solution cool down to
RT, add MQ up to 1 L and autoclave.
2. ASPA + N (50×): For 1 L: Add 297.5 g
(3.5 M) NaNO3, 26.1 g (0.35 M) KCl, and
74.8 g (0.55 M) KH2PO4 to 600 mL MQ in a
1-L cylinder. When all salts are dissolved, set
pH to 5.5 with KOH (use 5 M KOH). Add
MQ up to 1 L and autoclave.
3. ASPA-N (50×): For 1 L: Add 26.1 g (0.35 M)
KCl and 74.8 g (0.55 M) KH2PO4 to 600 mL
MQ in a 1-L cylinder. When dissolved, set
pH to 5.5 with KOH. Add MQ up to 1 L and
autoclave.
4. MgSO4 (1 M): For 1 L: Add 246.5 g
MgSO4⋅7H2O to 600 mL MQ in a 1-L cylinder. When all salts are dissolved, add MQ up
to 1 L and autoclave.
5. Trace element solution (1,000×): For 1 L:
Add 10 g (26.9 mM) EDTA, 4.4 g (15.3 mM)
ZnSO4⋅7H2O, 1.01 g (5.1 mM) MnCl2⋅4H2O,
0.32 g (1.3 mM) CoCl2⋅6H2O, 0.315 g
(1.3 mM) CuSO4⋅5H2O, 0.22 g (0.18 mM)
(NH4)6 Mo7O24⋅4H2O, 1.11 g (10 mM) CaCl2
and 1.0 g (3.6 mM) FeSO4⋅7H2O to 600 mL
MQ. When dissolved, set pH to 4.0 with
NaOH (use 1 M NaOH; 40 g/l) and HCl (use
1 M HCl; 75 mL 37 % hydrochloric acid/l),
fill MQ up to 1 L and autoclave. (see Note 1)
6. Vitamin solution (1,000×): For 100 mL: Add
100 mg thiamin-HCl, 100 mg riboflavin,
100 mg nicotinamide, 50 mg pyridoxine,
10 mg pantotenic acid, 2 mg biotin to 50 mL
of warm MQ (about 50–60 ° C) in a 100-mL
17
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
Screening for Compounds Exerting Antifungal Activities
cylinder. When all vitamins are dissolved,
add MQ up to 100 mL, sterilize by filtration,
and store at 4 °C under dark conditions.
Minimal medium (MM): For 500 mL: Add
under sterile conditions to 480 mL of sterile
MQ: 10 mL of 50 % glucose, 10 mL of
50 × ASPA + N, 1 mL of 1 M MgSO4, and
500 mL of 1,000× trace element solution. For
MM + agar, autoclave 480 mL of MQ with
7.5 g of agar (Scharlau) and add all components after autoclaving under sterile
conditions.
2× Minimal medium (2× MM): 2 % glucose,
2× ASPA + N, 4 mM MgSO4, 2× trace element solution, 0.06 % yeast extract.
Complete medium (CM): For 500 mL: Add
0.5 g casamino acids, 2.5 g yeast extract and
if required, 7.5 g agar to 480 mL of MQ and
autoclave. Afterwards, add under sterile conditions: 10 mL of 50 % glucose, 10 mL of
50× ASPA + N, 1 mL of 1 M MgSO4, 500 ml
of 1,000× trace element solution.
Saline solution: For 1 L: Add 9 g (0.9 % w/v)
NaCl to 900 mL MQ in a 1-L cylinder. When
NaCl is dissolved, add MQ up to 1 L and
autoclave.
YPD medium: 0.3 % yeast extract, 1 % bactopeptone, 2 % glucose.
Myracloth (Calbiochem, La Jolla, CA,
USA).
Cotton sticks (Hecht).
Flat bottom 96 well plate (transparent,
Sarstedt, Newton, NC, USA).
V-bottom 96 well plate (Sarstedt, Newton,
NC, USA).
Polystyrol 96 well plate, black (Greiner,
Monroe, NC, USA).
Multichannel pipette (20–300 mL) (Rainin,
Woburn, MA, USA).
Microtiter plate reader (e.g., Victor 3,
PerkinElmer, Waltham, MA, USA).
Incubator (Heraeus, Thermo Scientific,
Waltham, MA, USA).
Inverted microscope (Leica ICC50).
Microscope counting chamber.
SYTOX-Green (Invitrogen, Paisley, UK).
Fluorescence microscope allowing both light
and fluorescence imaging (GFP).
227
Methods
Inhibitory Testing Using
Growth Assays
This method can be used to generally test growth
inhibitory effects of compounds towards fungi
like A. niger. To study growth inhibition, the optical density of the cultures is followed and visualized by microscopic means. Figure 17.1 depicts
growth inhibition of A. niger in a 96 well plate
incubated with different concentrations of the
antifungals caspofungin and fenpropimorph.
1. Prepare spore solution of A. niger wild-type
strain as follows: Streak spores from a single
colony on a CM agar plate and incubate until
the plate is abundantly covered with sporulated mycelium (3–6 days, 25–37°).
2. In order to harvest spores from CM agar
plate, add 10 mL of saline solution to the
plate and carefully release spores by scraping over the surface plate with a sterile cotton
stick.
3. Pipette spore solution from the plate into a
sterile 15-mL tube. If required, remove
mycelial debris (vegetative mycelium, conidiophores) by filtration through a sterile
myracloth filter.
4. Count spores using a microscope counting
chamber.
5. Prepare a spore solution with the final titer of
7.5 × 105 spores/mL (see Note 2).
6. Start the growth inhibition assay with letting
the spores germinate first: Fill each well of a
flat-bottom 96 well plate with 30 mL sterile
MQ, 50 mL 2× MM and 20 mL A. niger spore
solution. Close the plate with a lid and incubate for 7 h at 30 °C.
7. Prepare a compound stock plate for efficient
and fast addition of the compounds by using
a V-bottom 96 well plate: Prepare serial dilutions of the compounds and add each 40 mL
per well. Every compound should be tested
in triplicate. Pipette also 40 mL water or any
other solvent used as negative control to at
least three wells. Store the compound plate at
4 °C until germination is finished.
228
J.-P. Ouedraogo et al.
Fig. 17.1 Growth inhibition of A. niger incubated with
different concentrations of the antifungals caspofungin
(cas) and fenpropimorph (fen). Note that DMSO is the
solvent for both compounds. (a) Optical density (OD620)
measured by a plate reader. (b) Microscopic pictures taken
with an inverted microscope after 24 h of growth
8. Add 45 mL sterile MQ and 75 mL 2× MM to
each well of the germination plate using a
multichannel pipette.
9. Transfer 30 mL of the compounds (and controls) from the compound plate to the germination plate using a multichannel pipette.
10. Incubate the growth inhibition plate at 30 °C
and record the kinetics of growth every hour
by measuring the optical density at 620 nm
(see Fig. 17.1a). Incubate for a maximum of
24 h (see Note 3).
11. Visualize growth of A. niger in the 96 well
plate of each condition via microscopy (see
Fig. 17.1b).
Cell Membrane Susceptibility Testing
Using a Sytox Green Assay
The integrity of cell membrane can easily be carried out using the SYTOX-Green assay [13].
SYTOX Green is a high-affinity nucleic acid
17
Screening for Compounds Exerting Antifungal Activities
stain that can penetrate cells having compromised
cell membranes but does not cross intact membranes. The method given below describes the
procedure for testing the susceptibility of cell
membranes of filamentous fungi and yeast
towards potential antifungals [8].
1. Obtain fresh spores or cells of the fungal
strains under investigation and count them
using a microscope counting chamber.
2. Inoculate 102 spores or 105 yeast cells in black
polystyrol 96 well plates containing 150 mL
YPD medium and incubate at 28 °C for
20–40 h for filamentous fungi and for 12–16 h
for yeast cells (see Note 4).
3. Add 0.2 mM final concentration of SYTOXGreen for filamentous fungi and 1 mM for
yeast cells and place the plate into the dark
(see Note 5).
4. Add 25 mL of the antifungal under investigation using serial dilutions.
5. Continue cultivation in the dark at 28 °C.
Measure the kinetics of fluorescence formation in minute intervals up to 2 h using a
microtiter plate reader at an excitation wavelength of 480 nm and an emission wavelength
of 530 nm (see Note 6).
6. Calculate the relative fluorescence values by
subtracting the fluorescence values of a culture incubated only with SYTOX-Green without an antifungal compound.
229
cytoplasmatically (strain JvD1.1) or nuclear
(strain RD6.47) targeted gfp gene under the control of the agsA promoter, can be used.
1. Obtain fresh spore solutions from the reporter
strains JvD1.1 (expressing PagsA-GFP) and
RD6.47 (expressing PagsA-H2B-GFP) as
described previously.
2. Inoculate 2 × 104 conidia from the reporter
strains in flat-bottom 96 well plate (Sarstedt)
containing 100 mL 2× CM.
3. Incubate for 6 h at 37 °C.
4. After spore germination, add 100 mL of a twofold dilution series for each antifungal compound to individual wells. The effect of each
compound shall be tested for at least three to
four different concentrations. Include respective negative (water or other solvent) and positive (caspofungin) controls.
5. After adding the compound solution, place the
microtiter plates for 3 more hours at 30 °C
(see Note 7).
6. Discard the medium by inverting the microtiter plate and analyze germlings that are
adherent to the bottom of each well by
fluorescence microscopy (see Note 8).
Compounds which induce agsA expression
will induce a strong GFP fluorescence even if
germ tube elongation is inhibited. A wildtype A. niger strain shall always be used as a
negative control because agsA expression
will be naturally induced after prolonged
cultivation.
Cell Wall and Cell Membrane
Susceptibility Testing Using an
agsA::GFP Assay
Notes
The agsA gene coding for (1,3)-alpha-d-glucan
synthase is specifically induced in response to
compounds that interfere with cell wall or cell
membrane integrity of A. niger [7, 11]. The agsA
gene is therefore an excellent and fungal-specific
marker for detecting cell surface integrity. Note
that bacteria, yeasts (except Schizosaccharomyces
pombe), plants, and mammals do not have a
(1,3)-alpha-d-glucan synthase. To study the effect
of compounds on (1,3)-alpha-d-glucan synthesis,
two A. niger reporter strains, containing either a
1. The color of the 1,000× trace element solution
is green when freshly made. After autoclaving, the color changes from green to purple
within 2 weeks.
2. Spore solutions of A. niger can be stored at
4 °C. However, all assays described work best
if the spore solution used is not older than
2 weeks.
3. The maximum time for incubation is 24 h,
because the wells dry out owing to medium
evaporation.
230
4. YPD is a complete medium for both yeast
and filamentous fungi. Cultivate cells until
the culture reaches the mid-logarithmic
growth phase. The time required is straindependent.
5. It is very important that all experiments involving SYTOX Green are performed in the dark.
0.2 mM and 1 mM SYTOX Green are the optimal concentration for filamentous fungi and
yeast, respectively.
6. Permeabilized mycelia or cells respond with
increasing fluorescence already after a few
minutes of incubation with the antifungal [14].
7. It is important to cultivate at 30 °C as higher
temperatures negatively interfere with GFP
folding.
8. Use a 40× objective. For GFP images, use a
fixed exposure of, for example, 2 s. Process
images using Adobe Photoshop.
References
1. Ostrosky-Zeichner L, Casadevall A, Galgiani JN,
Odds FC, Rex JH (2010) An insight into the antifungal pipeline: selected new molecules and beyond. Nat
Rev Drug Discov 9:719–727
2. Odds FC, Brown AJ, Gow NA (2003) Antifungal
agents: mechanisms of action. Trends Microbiol
11:272–279
3. Marcireau C, Guilloton M, Karst F (1990) In vivo
effects of fenpropimorph on the yeast Saccharomyces
cerevisiae and determination of the molecular basis of
the antifungal property. Antimicrob Agents Chemother
34:989–993
4. Latge JP (2007) The cell wall: a carbohydrate armour
for the fungal cell. Mol Microbiol 66:279–290
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5. Klis FM, Ram AF, De Groot PWJ (2007) A Molecular
and genomic view of the fungal cell wall. In: Howard J,
Gow NAR (eds) The mycota: biology of the fungal cell
VIII. Springer-Verlag, Berlin, Heidelberg, pp 97–120
6. Kartsonis NA, Nielsen J, Douglas CM (2003)
Caspofungin: the first in a new class of antifungal
agents. Drug Resist Updat 6:197–218
7. Meyer V, Damveld RA, Arentshorst M, Stahl U, van
den Hondel CA, Ram AF (2007) Survival in the presence of antifungals: genome-wide expression profiling
of Aspergillus niger in response to sublethal concentrations of caspofungin and fenpropimorph. J Biol
Chem 282:32935–32948
8. Ouedraogo JP, Hagen S, Spielvogel A, Engelhardt S,
Meyer V (2011) Survival strategies of yeast and
filamentous fungi against the antifungal protein AFP.
J Biol Chem 286:13859–13868
9. Agarwal AK, Rogers PD, Baerson SR, Jacob MR,
Barker KS, Cleary JD et al (2003) Genome-wide
expression profiling of the response to polyene,
pyrimidine, azole, and echinocandin antifungal agents
in Saccharomyces cerevisiae. J Biol Chem
278:34998–35015
10. Levin DE (2005) Cell wall integrity signaling in
Saccharomyces cerevisiae. Microbiol Mol Biol Rev
69:262–291
11. Damveld RA, vanKuyk PA, Arentshorst M, Klis FM,
van den Hondel CA, Ram AF (2005) Expression of
agsA, one of five 1,3-alpha-d-glucan synthase-encoding genes in Aspergillus niger, is induced in response
to cell wall stress. Fungal Genet Biol 42:165–177
12. Valiante V, Jain R, Heinekamp T, Brakhage AA (2009)
The MpkA MAP kinase module regulates cell wall
integrity signaling and pyomelanin formation in
Aspergillus fumigatus. Fungal Genet Biol 46:909–918
13. Thevissen K, Terras FR, Broekaert WF (1999)
Permeabilization of fungal membranes by plant
defensins inhibits fungal growth. Appl Environ
Microbiol 65:5451–5458
14. Theis T, Marx F, Salvenmoser W, Stahl U, Meyer V
(2005) New insights into the target site and mode of
action of the antifungal protein of Aspergillus giganteus. Res Microbiol 156:47–56
Fluorescence In Situ Hybridization
of Uncultured Zoosporic Fungi
18
Télesphore Sime-Ngando, Marlène Jobard,
and Serena Rasconi
Abstract
Recently, molecular environmental surveys of the eukaryotic microbial
community in lakes have revealed a high diversity of sequences belonging
to uncultured zoosporic fungi. Although they are known as saprobes and
algal parasites in freshwater systems, zoosporic fungi have been neglected
in microbial food web studies. Recently, it has been suggested that zoosporic fungi, via the consumption of their zoospores by zooplankters,
could transfer energy from large inedible algae and particulate organic
material to higher trophic levels. However, because of their small size and
their lack of distinctive morphological features, traditional microscopy
does not allow the detection of zoosporic organisms such as chytrids in the
field. We have designed an oligonucleotidic probe specific to Chytridiales
(i.e., the largest group of the true-fungal division of Chytridiomycota) and
provide simplified step-by-step protocols for its application to natural
samples using both the classical monolabeled-FISH and the CARD-FISH
approaches, for the assessment of uncultured zoosporic fungi and other
zoosporic microbial eukaryotes in natural samples.
Keywords
Fungi • Zoosporic fungi • Sporangia • Spores • Fluorescence in situ hybridization (FISH) • Classical monolabeled-FISH • Catalyzed reporter deposition-FISH (CARD-FISH) • Environmental samples
T. Sime-Ngando (*)
UMR CNRS 6023, Université Blaise Pascal,
Clermont II, 24 Avenue des Landais, BP 80026,
Aubière 63171 Cedex, France
e-mail: telesphore.sime-ngando@univ-bpclermont.fr
M. Jobard
LMGE UMR CNRS, U.F.R. SCIENCES ET
TECHNOLOGIES, 24 Avenue des Landais, BP 80026,
Aubière, 63171 Cedex, France
S. Rasconi
Department of Biology, University of Oslo,
Blindernvn. 31, Oslo 0371, Norway
Introduction
Recent molecular surveys of microbial eukaryotes have revealed overlooked, uncultured
environmental fungi with novel putative functions [1–3], among which zoosporic forms (i.e.,
chytrids) are the most important in terms of
diversity, abundance, and functional roles, primarily as infective parasites of phytoplankton
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_18, © Springer Science+Business Media, LLC 2013
231
232
[4, 5] and as valuable food sources for zooplankton via massive zoospore production, particularly in freshwater lakes [6–8]. However,
owing to their small size (2–5 mm), their lack of
distinctive morphological features, and their
phylogenetic position, traditional microscopic
methods are not sensitive enough to detect fungal zoospores among a mixed assemblage of
microorganisms. Chytrids occupy the most
basal branch of the kingdom Fungi, a finding
consistent with choanoflagellate-like ancestors
[9]. These reasons may help explain why both
the infective (i.e., sporangia) and disseminating
(i.e., zoospores) life stages of chytrids have
been misidentified in previous studies as,
respectively, phagotrophic sessile flagellates
(e.g., choanoflagellates, bicosoecids) and as
“small undetermined” cells. These cells often
dominate the abundance of free-living heterotrophic nanoflagellates (HNFs) and are considered the main bacterivores in aquatic
microbial food webs [2, 10]. Their contribution
ranges from 10 to 90% of the total abundance
of HNFs in pelagic systems (see review in
ref. [11]). Preliminary data have shown that up
to 60% of these unidentified HNFs can correspond to fungal zoospores [12], establishing
the HNF compartment as a black box in the
context of microbial food web dynamics [4].
A recent simulation analysis based on a Lake
Biwa (Japan) inverse model indicated that the
presence of zoosporic fungi leads to (1) an
enhancement of the trophic efficiency index,
(2) a decrease of the ratio detritivory/herbivory,
(3) a decrease of the percentage of carbon
flowing in cyclic pathways, and (4) an increase
in the relative ascendency (indicates trophic
pathways more specialized and less redundant)
of the system [13]. Unfortunately, because
specific methodology for their detection is not
available, quantitative data on zoosporic fungi
are missing.
Sporangia and the associated rhizoidal system
are characterized by a chitinaceous wall (a common fungal structure element for many species)
that can be targeted by specific fluorochromes
such as calcofluor white [14]. In contrast, because
the chitinaceous wall springs out after zoospore
encystment, chytrid zoospores completely lack
T. Sime-Ngando et al.
cell wall and chitin, precluding any simple use of
fluorochromes for their quantitative assessment
in natural environments [12]. Molecular
approaches, primarily fluorescence in situ hybridization (FISH), offer an alternative for quantitatively probing both chytrid sporangia and
zoospores in nature. FISH method is based on the
detection of targeted nucleic acid sequences by
the use of oligonucleotide probes labeled by a
fluorochrome, usually Cy3 [15, 16]. One of the
major limitations of FISH-based methods for
natural samples is the autofluorescence interference from autotrophic organisms. During the past
few years, numerous efforts have been made to
improve the sensitivity of monolabeled probes
for FISH assay, including the use of brightener
fluorochromes [17] or of signal amplification
with reporter enzymes [18]. Of particular interest
is the hybridization method using horseradish
peroxidase (HRP)-labeled probes activated by
fluorescent tyramide (also known as catalyzed
reporter deposition, CARD-FISH), which is very
efficient in overcoming the interference from
natural fluorescence [19]. The method is based
on the fact that each HRP-labeled probe catalyzes
the deposition of many labeled tyramides, so that
numerous fluorescent molecules are introduced
at the hybridization site, resulting in net
fluorescence signal amplification, compared to
the classical Cy3-monolabeled FISH probes.
[20]
The main objective of this chapter is to provide, in a simplified step-by-step format, classical FISH and CARD-FISH protocols for the
identification and quantitative assessment of
uncultured zoosporic fungi and other zoosporic
microbial eukaryotes in natural aqueous environments (cf. 12), together with practical advices on
how to apply the methods.
Materials
1. Gloves (should be worn when manipulating
most of the following materials).
2. 0.6-mm pore size polycarbonate white filters
(e.g., catalog no. DTTP02500, Millipore,
Billerica, MA, USA).
18 Fluorescence In Situ Hybridization of Uncultured Zoosporic Fungi
3. Appropriate Cy3-labeled oligonucleotidic
probe and its reverse complement stored at
−20 °C (see Note 1).
4. Sodium dodecyl sulfate (SDS).
5. Formaldehyde 37%.
6. FISH hybridization buffer (0.9 M NaCl,
20 mM Tris–HCl (pH 7.2), 0.01% SDS) containing 30% formamide and 2.5 ng ml−1 of
Cy3-labeled probe (see Note 2).
7. Washing buffer—20 mM Tris–HCl (pH 7.2),
5 mM EDTA, 0.01% SDS, 112 mM
NaCl [21].
8. Appropriate filtration columns equipped with
a peristaltic pump.
9. DAPI—4.6-diamidino-2-phenylindole.
10. Glass slides and coverslips.
11. Nonfluorescent immersion oil.
12. Epifluorescence microscope equipped with
appropriate filter sets (blue and UV) and
neofluar objective lens (optional).
13. CARD-FISH hybridization buffer: 30%
deionized formamide, 0.9 M NaCl, 20 mM
Tris–HCl (pH 7.5), 0.01% SDS, and 10%
blocking reagent (e.g., Roche Diagnostics/
Boehringer).
14. Appropriate oligonucleotide probe labeled
with HRP (in our case, commercially synthesized by Biomers, Germany).
15. TNT buffer—0.1 M Tris–HCl (pH 7.5),
0.15 M NaCl, and 0.05% Tween 20.
16. TSA mixture—(1:1) of 40% dextran sulphate
(Sigma-Aldrich, St. Louis, MO, USA) and
1× amplification diluent (PerkinElmer LAS,
Waltham, MA, USA).
17. Fluorescein isothiocyanate coupled with tyramide (1×, Perkin-Elmer LAS).
Methods
Classical FISH Probing (see Note 3)
Fix experimental samples with 2% formaldehyde,
vol:vol final concentration. The fixation step is
facultative and can be avoided when observations
are made without delay.
1. Filter-collect appropriate volumes (×3 replicates) of cultures, enriched cultures, or natural
2.
3.
4.
5.
6.
7.
233
samples containing zoosporic organisms onto
0.6-mm pore size polycarbonate white filters
(see Note 4) by using gentle vacuum (<20 kPa).
In the dark, pour the filters with targeting fungal zoospores and sporangia and perform
hybridization in the standard FISH hybridization buffer (containing 30% formamide and
2.5 ng mL−1 of the Cy3-labeled oligonucleotide
probe) for 3 h at 46 °C (see Note 5).
Use the reverse complement probe in a negative control to check for the autofluorescence
interference from fungi and other natural
plankton present in natural samples.
After hybridization, thoroughly rinse the filters
in the washing buffer for 30 min at 48 °C.
Counterstain the filters in the dark at room
temperature for 5 min with DAPI 0.5 mg mL−1,
and repeat the washing step.
Mount the filters between glass slides and
coverslips using appropriate nonfluorescent
immersion oil (see Note 6). At this stage,
mounted filters can be conserved at −20 °C
until microscopic observation.
In a dark room, examine the filters under an
epifluorescence microscope equipped with
appropriate set of filters and objective lens.
Shift between blue and UV light to distinguish
between Cy3 stain and DAPI, use different
convenient magnifications for sporangia and
zoospores, and apply a standard procedure for
microscopic counting.
CARD-FISH Probing (see Note 7)
Perform steps 1 and 2 in Classical FISH
Probing.
1. In the dark, pour the filters with targeting fungal zoospores and sporangia and perform
hybridization in the CARD-FISH hybridization buffer (containing 30% formamide and
2.5 ng mL−1 of HRP labeled oligonucleotide
probe) for 3 h at 35 °C (see Note 5).
2. After hybridization, thoroughly rinse the filters
in the washing buffer for 2 × 20 min at 37 °C.
3. Equilibrate samples to increase enzyme activity in TNT buffer at room temperature for
15 min.
234
4. Perform signal amplification by 30-min incubation in TSA mixture, to which fluorescein
isothiocyanate coupled with tyramide was
added (1:50 vol/vol).
5. Transfer filters in two successive 5-ml TNT
buffer baths at 55 °C for 20 min, in order to
stop the enzymatic reaction and remove the
dextran sulphate.
6. Follow steps 6 and 7 in Classical FISH
Probing.
Notes
1. We propose to used a probe named Chyt1061
(sequence 5¢ > 3¢: CATAAGGTGCCGAACAA
GTC), because of the sequence position (1,061
base pairs) on Saccharomyces cerevisiae smallsubunit rDNA molecule (GenBank accession
no. J01353). According to Behrens and collaborators [22], this position provides a good
accessibility for FISH probing. There were
two mismatches in the middle of the probe
with sequences of chytridiales species (cf. 12),
which did not result in loss of positive signal.
Chyt1061 was designed in silico for targeting
fungal species in the order Chytridiales,
which is the largest order of the division
Chytridiomycota (chytrids), mainly represented
by phytoplanktonic parasites in aquatic environments [12]. The design was based on the
alignment of rDNA sequences of Chytridiales
obtained from GenBank (http://www.ncbi.nlm.
nih.gov/), together with 106 sequences derived
from 18S rDNA PCR surveys of freshwater
picoeukaryotes conducted in French Lakes
Pavin, Godivelle, and Aydat [12]. Distinct
rDNA sequence unique to target organisms was
localized and imported in Primer3 software
(http://fokker.wi.mit.edu/primer3/input.htm) in
order to design a probe with size between 18
and 27 bases, probe melting temperature (Tm)
between 57 and 63 °C, and GC percentage at
about 50%. The probe was analyzed for potential complementarities and no dimers or hairpins were found using Netprimer software
(http://www.premierbiosoft.com/netprimer/
netprlaunch/netprlaunch.html). The probe was
T. Sime-Ngando et al.
commercially synthesized by MWG-Biotech
Company (Germany) and labeled with the
fluorochrome Cy3 for classical FISH or application to environmental samples using the
CARD-FISH approach [12].
2. In case you design your own probe because of
the increasing availability of sequences in the
database, hybridization stringency should be
tested and validated using appropriate positive
and negative cultures, before application to
natural samples. In the absence of laboratory
cultures, our probe Chyt1061 was evaluated
from an adaptation of an alternative approach
called clone-FISH, known from prokaryotes
[23]. This approach is based on the genetic
modification of a clone of Escherichia coli by
inserting plasmid vector containing the target
rDNA sequence. In our adaptation of the
approach, cells of E. coli clone BL21 star were
genetically transformed by inserting plasmid
vector containing rDNA sequence from several different target fungal cells. Specific plasmid inserts came from freshwater lake surveys
of picoeukaryote 18S rDNA and fungal 18SITS rDNA as well (cf. 12).
3. The specificity of the designed probe should be
checked both in silico by using a basic local
alignment search tool (e.g., BLAST, [24]), and
in vivo by screening of clone libraries with
classical FISH (the clone-FISH approach can
be used here). In our case, clones containing
rRNA gene inserts from different eukaryotes
closely related to microorganisms of interest,
and negative controls as well, were FISHtargeted following the protocol described in
this chapter (i.e., with 30% formamide in the
hybridization buffer). In addition, the in vivo
transcription of the 18S rRNA gene insert was
induced with IPTG (1 mM) for 1 h. The
designed probe or its reverse complement
probe was used, depending on the orientation
of insertion into the vector, i.e. 3¢ → 5¢ or 5¢ → 3¢
way downstream the T7 promoter (cf. 12).
4. In vivo tests using the clone-FISH approach
(see Note 1) yield the best fluorescence signal
when hybridization was performed at 30%
formamide concentration in the hybridization
buffer. Assuming an increase of the effective
18 Fluorescence In Situ Hybridization of Uncultured Zoosporic Fungi
5.
6.
7.
8.
9.
hybridization temperature of 0.5 °C per 1% of
added formamide, the melting temperature
(Tm) of the probe Chyt1061 was experimentally calculated at 61 °C [12].
This protocol is suitable for cultures and
enriched cultures (i.e., concentrates of targeted
zoosporic organisms during host blooms) (cf.
12). However, the FISH resolution for fungal
images and species identification based on
sporangium features is poor, compared to the
calcofluor approach, which is more appropriate for the identification of zoosporic organisms [14]. In addition, The Cy3-monolabeled
FISH probing of natural samples clearly
showed that the fluorescence of targeted chytrid zoospores may be quite similar to the
autofluorescence from natural picoautotrophs.
That is why CARD-FISH is more appropriate
for environmental samples.
For natural waters, the appropriate volume
depends on the trophic status of the natural
waters, whether the sampling period corresponds to a bloom period, and on the nature of
the phytoplankton species. Zoosporic parasites
are much more abundant when large-size phytoplankton hosts such as diatoms or filamentous
cyanobacteria develop [5, 14]. In oligotrophic
waters, concentration of natural samples could
be required before harvesting targeted organisms onto polycarbonate filters [15].
One filter corresponding to one sampling time
point can be cut in pieces before hybridization
when several probes are used.
The mounting medium should not be
fluorescent and will minimize the fading of
fluorochromes. An example of mountant is a
solution composed of 50% glycerol, 50%
phosphate buffered saline (0.05 M Na2HPO4,
0.85% NaCl, pH 7.5), and 0.1% p-phenylenediamine (made fresh daily from a frozen 10%
aqueous stock solution; Sigma-Aldrich, St.
Louis, MO, USA).
The CARD-FISH protocol is suitable for natural samples, primarily in oligotrophic waters
(see Note 3), where its application improves
the detection and the recognition of chytrid,
because of the enhanced signal conferred by
HRP-labeled probes, compared to monolabeled
235
oligonucleotides. In addition, the choice of
fluorescein as stain (emission in the green spectrum at 520 nm) significantly reduces the interference from natural fluorescence of autotrophic
organisms, thereby preventing the use of the
deductive approach based on a double counting
of the same sample (i.e., with and without
hybridization) [25]. However, similar to the
simple FISH approach, the CARD-FISH
resolution for fungal images and species
identification based on sporangium features is
poor, compared to the calcofluor approach,
which is more appropriate for the identification
of zoosporic organisms [14].
Acknowledgements M. Jobard and S. Rasconi were
supported by Ph.D. Fellowships from the Grand Duché du
Luxembourg (Ministry of Culture, High School, and
Research) and from the French Ministère de la Recherche
et de la Technologie (MRT), respectively. This study
receives grant-aided support from the French ANR
Programme Blanc # ANR 07 BLAN 0370 titled DREP:
Diversity and Roles of Eumycetes in the Pelagos.
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Staining Techniques
and Biochemical Methods
for the Identification of Fungi
19
Jeyabalan Sangeetha and Devarajan Thangadurai
Abstract
In the past, conventional identification of fungi relied on the combination of
morphological and physiological properties. In recent years, morphological
studies, supplemented with staining techniques and biochemical methods,
still play an important role in the overall identification of fungi in the molecular era. In most instances, these tools are widely used to determine the
correct identity of yeasts and molds at the genus and species levels.
Keywords
Identification • Molds • Yeasts • Staining techniques • Biochemical
methods
Introduction
In general, fungal identification requires greater
visual acuity than bacteria. Unlike other important microorganisms such as bacteria and viruses,
the identification of fungi heavily relies on morphological criteria. The characteristics of fungal
structure are identified by observing colonial
growth both macroscopically and microscopically. These morphological features and other
classical methods that are routinely used in
J. Sangeetha (*)
Department of Zoology, Karnataka University,
580003, Dharwad, Karnataka, India
e-mail: drsangeethajayabalan@gmail.com
D. Thangadurai
Department of Botany, Karnataka University,
580003, Dharwad, Karnataka, India
classification are also useful in fungal
identification. The correct identity of fungal taxa
is of great practical relevance in clinical mycology,
plant pathology, biodeterioration, and biotechnology. A recent review illustrates the inability to
identify fungi at the species, or even at the genus,
level in many cases [1]. Many species of fungi,
ascomycetes, basidiomycetes, and zygomycetes
in particular, have different microscopic and
macroscopic characteristics in each stage of their
life cycle. Moreover, they are synonymous to
each other with many names used to describe the
same organism [2].
The ever-increasing number of yeasts and
molds that are frequently impossible to identify
using morphological criteria due to lack of sporulation has driven the need for the design and
development of rapid and robust biochemical and
molecular identification tools [3–7]. In recent
years, the ability to accurately and reproducibly
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_19, © Springer Science+Business Media, LLC 2013
237
238
identify fungi has been greatly enhanced through
automated biochemical methods and comparative
DNA analysis [8–14]. The procedures for the
identification of yeasts and molds are different.
Identification of molds to the species level has
been difficult, because of the amount of experience required to accurately identify these
filamentous fungi. In many cases, molds are
identified based upon colony and microscopic
characteristics. The microscopic structures and
morphological features like type, size, shape, and
arrangement of spores, and size, color, and septation of hyphae usually provide definitive
identification for molds. However, species-level
mold identification has not been held to the same
standards as bacterial and yeast identification. In
yeast, morphological criteria and biochemical
tests are generally used to determine genus and
species level, respectively.
Direct microscopic examination of the fungal
specimen provides a clear view and valuable
information about the fungal structure. Yeasts
and molds can be identified based on a combination of macroscopic, microscopic, and biochemical analysis. Macroscopic evaluation of fungi
provides information about the probable region
of the presence of fungi in a specimen, whereas
microscopic examination reveals the important
features like type and color of hyphae, conidia,
septae, spores, and also the concentration of
fungi. A definitive fungal stain is an important
tool that is needed to begin the identification of
fungal specimens. Detection of fungi using direct
microscopy with various stains is quick, simple,
and can be optimized through the use of ready-touse staining solutions. The selection of staining
method is primarily based on the sample used. A
number of stains are routinely available to visualize fungi; some of these are special fungal stains
and others are more general in use [15]. Achieving
a successful identification comes from the use of
appropriate stains and further microscopic examination. The major growth forms of the fungi that
help in identification after staining are the yeast
cells, hyphae, pseudohyphae, arthroconidia, chlamydoconidia, and endosporulating spherules [16,
17]. On the other hand, visualization of fungi in
smear often helps in identification; in some cir-
J. Sangeetha and D. Thangadurai
cumstances, it is essential to establish further
study on fungi. Even more, the evaluation of
staining is useful when multiple organisms are
cultured [18].
Additionally, fungal identification requires
biochemical tests that will distinguish genera
among families and species among genera.
Further, strains within a single species are usually
distinguished by genetic or immunological criteria. The identity of certain fungi depends on their
ability or inability to grow in the absence of nutritional substances such as carbon and nitrogen
sources. In addition, several routine biochemical
tests, including urease production and proteolysis, are available for the identification of many
molds. Moreover, the presence of various
enzymes as determined by the biochemical tests
is also useful in identifying fungi. When grown in
selective liquid or solid media, fungi ferment carbohydrates and produce acids, alcohols, gases,
and metabolic and enzymatic products in patterns
characteristic of their genus and/or species. These
fermentation products are commonly used in the
differential identification of fungi [19].
Most frequently, these instant and incubated
biochemical tests and other “expert systems”
monitor the aptitude of the isolate to assimilate
and ferment various sugar and nitrogen sources.
In addition, the identification of yeasts has now
been regularized by a variety of commercially
available strips and kits that can be used to examine rapidly the absorption of carbon as well as
nitrogen. The most reliable commercially available yeast identification kits are API 20C AUX,
ATB 32C, MicroScan, and Vitek systems. In general, biochemical tests are not important in identifying molds as they are specific for yeasts and
dermatophytes [20]. Many yeast-like fungi such
as the genera Geotricum and Trichosporon, which
form arthrospores, require a series of biochemical tests for their definitive identification. Most
recently, fungal DNA analysis has been considered a powerful identification tool that requires
specialized equipment and is impractical in much
of routine laboratory work. Although there is an
increasing move towards molecular diagnostic
approaches, often use of the more easily available
and still fundamental staining techniques and
19
Staining Techniques and Biochemical Methods for the Identification of Fungi
other biochemical tests are the choice of mycologists for much of the day-to-day fungal
identification. The goal of this chapter is to
acquaint mycologists with various staining techniques and differential biochemical tests available for detecting fungal specimens.
Materials
See Note 1.
1. A pure culture of 24- to 48-h old yeast cells
(growing on Sabouraud or other nonselective agar)
2. Microscopic slides
3. Sterile saline or water
4. Flame source
5. Fume hood
6. A straight nichrome wire with a long handle
for stabbing inoculation
7. A bent nichrome wire with a long handle for
handling the mycelia types of fungal
cultures
8. Pair of short, stiff teasing needles helpful in
pulling apart dense masses of mycelium on
the slide for better microscopic examination
9. Scalpel with blades
10. Pair of forceps
11. Light microscope, phase contrast microscope, and fluorescence microscope with
filters
12. 20% KOH-Glycerol solution—20 g KOH,
20 mL glycerol, 80 mL distilled water (see
Note 2)
13. Lactophenol Cotton Blue stain—20 mL lactic acid, 20 mL phenol, 40 mL glycerol,
0.05 g cotton blue or aniline blue. Dissolve
phenol in lactic acid, glycerol, and distilled
water, finally add cotton blue and mix well
14. India ink
15. Crystal violet
16. Gram’s iodine
17. Ethanol
18. Absolute alcohol
19. Safranin
20. Giemsa stain
21. Wright stain—9.0 g (0.3% w/v) powdered
Wright’s stain, 1.0 g (0.033% w/v) powdered
22.
23.
24.
25.
26.
27.
28.
29.
30.
31.
32.
33.
34.
35.
36.
37.
38.
39.
239
Giemsa stain, 90 mL glycerin, and 2,910 mL
absolute acetone-free methanol (see Note 3)
Weigert’s solution A—0.6 g ferric chloride,
100 mL distilled water, 0.75 mL hydrochloric acid
Weigert’s solution B—1.0 g hematoxylin,
100 mL 95% ethanol; combine equal parts
of solution A and B
Weigert’s iron hematoxylin solution
A—1.0 g of hematoxylin, 100 mL 95%
ethyl alcohol
Weigert’s iron hematoxylin solution
B—4.0 mL 29% aqueous ferric chloride,
95 mL distilled water, 1.0 mL concentrated
hydrochloric acid
Weigert’s iron working solution—mix equal
parts of Weigert’s iron solution A and B
Acridine orange
Schiff’s reagent—0.025 g pararosaniline,
100 mL distilled water, sulfur dioxide gas
(see Note 4)
Aldehyde fuchsin—2 g basic fuchsin, 1 mL
paraldehyde, 1 mL concentrated hydrochloric acid, 200 mL 70% ethanol; ripen at room
temperature for 48–72 h
Chromic acid—5 g chromic trioxide,
500 mL distilled water (see Note 5)
Metanil yellow—0.25 g metanil yellow,
100 mL distilled water, 0.25 mL glacial
acetic acid
15% Potassium hydroxide solution—15 g
potassium hydroxide, 20 mL glycerol,
80 mL distilled water; store at 25 °C and
discard if precipitation occur
0.1% Calcoflour White (CFW) solution
Mucicarmine stain—1.0 g carmine, 0.5 g
aluminum chloride (anhydrous), 2 mL distilled water (see Note 6)
Formal–ethanol mixture—10 mL 40%
formaldehyde, 90 mL absolute alcohol
5% Periodic acid (see Note 7)
Basic fuchsin solution
Sodium metabisulphate solution—1.0 g
sodium metasulphite, 10 mL hydrochloric
acid, 190 mL distilled water (see Note 8)
0.2% Light green solution—0.2 g light
green, 100 mL distilled water, 0.2 mL glacial acetic acid
240
40. Xylene (see Note 5)
41. 1% Sodium metabisulphite
42. Hexamine, preheated in a water bath to
56 °C for 1 h
43. Ferric chloride
44. 5% Sodium thiosulphate solution
45. 5% Silver nitrate, store in dark-colored bottle at 4 °C (see Note 9)
46. 3% Aqueous methenamine (hexamethylene
tetramine) (see Note 9)
47. Aniline–acetic acid (see Note 5)
48. Thiosemicarbazide
49. Schmorl’s solution—30 mL ferric chloride
(1% aqueous), 4 mL potassium ferric cyanide, 6 mL distilled water; make immediately before use and do not reuse
50. Mellor bleach solution A—1% potassium
permanganate
51. Mellor bleach solution B—1% sulphuric
acid
52. Mellor bleach working solution—mix
Mellor bleach solution A and B
53. Mellor bleach solution C—1% oxalic acid
54. Ammonium silver stock solution—25 mL
10% silver nitrate, add ammonium hydroxide drop by drop, until solution precipitates
and clears again
55. Ammonium
silver
working
solution—12.5 mL ammonium silver stock solution, 37.5 mL distilled water (see Note 10)
56. 0.1% Gold chloride
57. Neutral red stain
58. Sulfation reagent—45 mL glacial acetic
acid, 15 mL concentrated sulphuric acid
(see Note 11)
59. Toluidine blue O—3 g toluidine blue O,
60 mL distilled water, 2 mL concentrated
hydrochloric acid, 140 mL absolute ethyl
alcohol
60. Basal salt medium
61. 0.1% Carbohydrate
62. Bromocresol purple
63. Sugar disc
64. Whatman filter paper
65. Sugars
66. Hot-air oven
67. Yeast Nitrogen Base (YNB) medium
68. Molten agar
69. Yeast Carbon Base (YCB) medium
J. Sangeetha and D. Thangadurai
70.
71.
72.
73.
74.
75.
76.
77.
78.
79.
80.
81.
82.
83.
84.
85.
86.
87.
88.
89.
90.
91.
92.
93.
94.
95.
96.
97.
98.
99.
100.
101.
102.
103.
Peptone disc
Potassium nitrate
Durham’s tube
Skim milk agar
Casein agar
Microgranular cellulose
Inoculation chamber
Autoclave
Petriplates
Incubator
Mycosel agar with cyclohexamide plate
10% Tween solution
Cellulolysis Basal Medium (CBM)—5 g
C4H12N2O6, 1 g KH2PO4, 0.5 g MgSO4·7H2O,
0.1 g yeast extract, 0.001 g CaCl2.2H2O,
1000 mL distilled water
0.5% Esculin
0.5% Arbutin
2% Ferric sulfate
Sabouraud agar plates
Christensen’s urea agar
API 20C Yeast Identification Kit
Biomerieux Vitek Yeast Biochemical Card
Abbott Yeast Identification Kit
Colorimeter (Vitek colorimeter, 52–1210)
Filling stand (e.g., Vitek 52–0700)
Sterile wooden applicator sticks
Sterile Pasteur or plastic pipettes (5 mL)
Squeeze bottle
Water bath (48–50 °C)
0.45–0.50% Saline
Sterile tubes
Fine-tip markers
0.05% Noble agar
Sterile inoculating loop
Vortex mixer
Polyester film
Methods
Wet Mount Techniques
Fungal specimens can be visualized using wet
mount techniques through suspension of culture
in either water or saline, mixed with alkali to dissolve background material [21] or mixed with a
combination of alkali and contrasting dye (e.g.,
lactophenol cotton blue or India ink) [22, 23].
19
Staining Techniques and Biochemical Methods for the Identification of Fungi
The dyes nonspecifically stain the fungal material, which increases contrast with the background
and permits examination of the detailed structures. A variation is the India ink method, in
which the ink darkens the background rather than
the fungi.
Potassium Hydroxide Wet Mount
Potassium hydroxide (KOH) is used to dissolve
proteinaceous material and facilitate detection of
fungal elements that are not affected by strong
alkali solution. It is a strong alkali used as a clearing agent to observe fungi in a wet mount preparation. The concentration of KOH is usually based
on the specimen that is being used. Normally,
10–20% KOH is used; occasionally, 40% is used
when the specimen is not cleared by 10–20% of
KOH. In this method, the fungal structures, such
as hyphae, large yeasts (Blastomyces), spherules,
and sporangia, are well distinguished. In unstained
preparations (KOH without stain), the fungal
structures may be enhanced by using a phase contrast microscope [18, 24]. The clearing effect
throughout the specimen can be accelerated by
gently heating the KOH preparation.
Visualization of fungi can be further enhanced
by the addition of dyes to the preparation. This
method is quick, simple, and inexpensive [25, 26].
1. Place a large drop of KOH solution with a
Pasteur pipette.
2. Transfer small quantity of the culture with a
loop or the tip of a scalpel into the KOH drop.
3. Put a clean coverslip over the drop gently so
that no air bubble is trapped.
4. Clearing can be hastened by gentle heating of
the slide, but it is best avoided.
5. Observe under 20× and 40× objective of light
or phase contrast microscope.
6. Look for budding yeast cells; branching
hyphae; type of branching; and the color, separation, and thickness of hyphae (see Note 12).
Lactophenol Cotton Blue Wet Mount
Lactophenol cotton blue (LCB) is a mounting
medium commonly used in microbiology laboratories for preparing mounts of fungal cultures.
LCB is used as both mounting fluid and stain. In
this method, phenol will kill the organisms, and
241
the lactic acid preserves fungal structures; chitin
in the fungal cell wall is stained by the cotton
blue. It can be used alone or in conjunction with
KOH. Library slides may be made by allowing
the mount to dry for 3 weeks and then sealing
with collodion [24, 27, 28].
1. Put a large drop of LCB with a Pasteur
pipette.
2. Transfer a small quantity of the culture to the
drop.
3. Tease the culture well with teasing needles, so
as to get a uniform spread.
4. Put on a coverslip gently to avoid entrapment
of air bubbles.
5. Examine under the 20× and 40× objectives of
light microscope.
6. Observe the morphological features carefully.
7. Fungal elements will stain deep blue against a
clear pale-blue background.
India Ink Wet Mount
India ink can be added to specimens to provide
dark background that will highlight hyaline yeast
cells and capsular material. This method is used
to detect microorganisms that are surrounded by
capsules. The dye is excluded by the capsule, creating a clear halo around the yeast cell. It is a
rapid method for the preliminary detection and
identification of specimens containing species of
Cryptococcus [29].
1. Add a small drop of India ink on a smear.
2. Place a coverslip over the smear and press it
gently to obtain a thin mount.
3. If India ink is too thick (dark), dilute it by 50%
with saline.
4. Allow the preparation to stand for few minutes
to settle.
5. Scan under low power in reduced light; switch
to high power, if necessary.
6. Organisms possessing a capsule appear highly
refractile, surrounded by a clear zone against a
dark background.
Staining Techniques
Direct microscopic examination without stain
lacks sensitivity, especially when hyphae are
242
sparse in the specimen. A variety of differential
stains are commonly used like Gram, Giemsa,
Wright stain, toluidine blue O, and Weigert’s
iron hematoxylin to stain fungi [30]. The sensitivity of microscopic examination is improved
when fungus-enhancing stains like Mayer’s
mucicarmine, periodic acid Schiff, Gomori’s
methenamine silver, acridine orange fluorescent,
calcoflour white, thiosemicarbazide, FontanaMasson, and Gridley’s stains are used. Since the
stain is immediately taken up by the fungal cell
wall in the scraping, the staining usually becomes
much brighter after 5–10 min [31]. Some specimens need alkali pretreatment. In that case, it is
important to make sure that they do not react for
a long period of time; otherwise, a gelatinous
consistency will form, and the specimen should
be neutralized with 10% lactic acid before staining and adjusted to pH 3.0–5.0 [32]. The method
of preparation of smear for staining is as
follows:
1. Take a clean grease-free glass slide.
2. Place a large drop of saline solution.
3. Transfer a small quantity of the culture with a
loop or the tip of a scalpel into the saline drop.
4. Make a smear over the surface of the slide.
5. Fix by heat, if necessary.
Gram Staining
Gram stain is a key starting point to identify
microbial species. The stain differentiates membrane structures between gram-positive and
gram-negative microorganisms. Gram-positive
microbes have a thick cell wall made up of peptidoglycan (50–90%), which are stained purple
by crystal violet, whereas gram-negative
microbes have a thinner layer (10% of cell wall),
which are stained pink by the counter-stain
safranin [24, 30, 33–36].
1. Apply two drops of crystal violet on smear
for 30 s.
2. Wash with tap water.
3. Add two drops of Gram’s iodine for 30 s.
4. Repeat step 2.
5. Add 95% ethanol.
6. Repeat step 2.
7. Add two drops of safranin.
8. Repeat step 2.
J. Sangeetha and D. Thangadurai
9. Observe in microscope under oil immersion.
10. Yeasts are gram-positive, but poorly stained;
Cryptococcus neoformans is a notable exception (gram-negative).
Giemsa Staining
A variety of “Romanowsky-type” stains with
mixtures of methylene blue and azure eosin compounds have been used successfully for many
years on diverse fungi with various procedures
and modifications. Giemsa stain is a member of
the Romanowsky group of stains, which are
defined as being the black precipitate formed
from the addition of methanol [37]. In this stain,
eosin ions are negatively charged and stain basic
components of cells orange to pink. It was also
originally designed to incorporate cytoplasmic
(pink) staining with nuclear (blue) staining and
fixation as a single step for smears and thin films.
This stain has widely been used to examine
Pneumocystis jiroveci, Rhinosporidium seeberi,
and Histoplasma capsulatum [38–40].
1. Flood the smear with methyl alcohol and leave
for 3–5 min for fixation.
2. Add prepared Giemsa stain and leave for
45 min.
3. Wash slide thoroughly with running tap
water.
4. Blot dry with absorbent paper.
5. Observe under oil immersion.
6. Look for intracellular budding yeasts; fungi
stain with purplish-blue.
Wright Staining
The Wright stain is an alcoholic solution of methylene blue, azure A, thionin, and eosin Y. Methyl
groups are activated and react with charged components of the cell to produce coloration. It is
used to detect blood parasites, viral and chlamydial inclusion bodies, yeast cells, and species of
Pneumocystis. Eosin ions are negatively charged
and stain basic components of cells orange to
pink, whereas other dyes stain acidic cell structures to various shades of blue to purple [41, 42].
1. Cover the smear with freshly filtered Wright
stain and leave for 1–3 min.
2. Without removing the stain, pour on buffer
solution (pH 6.4).
19
Staining Techniques and Biochemical Methods for the Identification of Fungi
3. Gently mix buffer and stain; upon proper mixing, metallic green sheen (green scum) rises to
the surface of the fluid.
4. Leave for 3 min or longer.
5. Wash the slide gently with flowing tap water
and wipe the bottom of the slide with a clean
filter paper.
6. Air-dry the slide and observe under the
microscope.
7. Intracellular yeast cells are typically stain blue
and species of Pneumocystis stain purple.
Weigert’s Iron Hematoxylin Staining
This stain can be used with fixatives that include
polyvinyl alcohol, sodium acetate, and formalin.
The staining method involves application of haemalum, which is a complex formed from aluminum ions and oxidized hematoxylin. This stains
nuclei of cells blue. Counterstain eosin Y may
also be used to color other structures in various
shades of red, pink, and orange [28, 43–47].
1. Add staining solution on a smear and leave for
1–2 h.
2. Rinse with tap water.
3. Add 1% HCl.
4. Add 70% ethanol.
5. Repeat step 2.
6. Counterstain with eosin Y, if necessary.
7. Dehydrate with ethanol.
8. Clear with xylene and observe under
microscope.
9. Yeast cells stain blue–gray to black.
Acridine Orange Staining
Acridine orange is a fluorochromatic dye that
binds to nucleic acids of fungi. Under UV light,
acridine orange stains RNA and single-stranded
DNA orange, while double-stranded DNA
appears green. At neutral pH, fungi and cellular
materials stain reddish orange. At acid pH, fungi
remain reddish orange but background material
stains greenish yellow [31, 48–53].
1. Add Weigert’s iron hematoxylin on smear for
5 min.
2. Wash well with tap water.
3. Place few drops of acridine orange solution
for 2 min.
4. Repeat step 2.
243
5. Observe smear on the fluorescence
microscope.
6. Fungi stain bright orange and the background
appears greenish yellow.
Gridley Staining
Gridley staining method is used to identify fungi,
based on Bauer chromic acid leucofuchsin stain
with the addition of Gomori’s aldehyde fuchsin
stain and metanil yellow as counterstains. Against
a yellow background, hyphae, conidia, yeast capsules, elastin, and mucin appear in different
shades of blue to purple. It can be used to identify
Rhiosporidium seeberi and Histoplasma capsulatum [28, 47, 51, 54, 55].
1. Place chromic acid on smear for 1 h.
2. Wash well with tap water.
3. Treat with sodium metabisulphite bleach for
1 min.
4. Repeat step 2.
5. Rinse with distilled water.
6. Place in Schiff’s reagent for 20 min.
7. Repeat step 2.
8. Rinse with 70% ethanol.
9. Place in aldehyde fuchsin for 30 min.
10. Rinse off excess with 95% ethanol.
11. Repeat step 2.
12. Counterstain with metanil yellow for 1 min.
13. Rinse well with distilled water.
14. Dehydrate and observe under fluorescence
microscope.
15. Fungi show purple color with yellow background (see Note 13).
Calcoflour White Staining
Calcoflour White (CFW) stain is used to detect
fungal elements, particularly Pneumocystis species. The fluorophore shows a high affinity for
chitin-forming hydrogen bonds with free hydroxyl
groups and stains fungal cell walls blue. The use
of CFW staining requires the addition of KOH,
which helps to dissolve keratinized particles and
emulsify solid, viscous material and enhance the
visualization of fungal elements in microscopic
examination. Positive results are indicated by a
bright green to blue fluorescence using a
fluorescence microscope (see Note 14) [56]. A
bright yellow-green fluorescence is observed
244
when collagen or elastin is present. KOH-CFW
preparations may be preserved for several days at
4 °C. This stain can be used to identify Fusarium
solani, Aspergillus fumigatus, and Candida
albicans [24, 56–59].
1. On smear add a drop of 15% KOH and a drop
of the CFW solution or mix in equal volumes
before processing.
2. Mix and place a coverslip over the material.
3. If necessary, allow the KOH preparation to
remain at room temperature (25 °C) for a few
minutes until the material has been cleared;
the slide may be warmed to speed up the clearing process.
4. Observe the slide by UV microscopy.
5. Fungal cell walls fluorescence apple green to
blue.
Mayer’s Mucicarmine Staining
Mucicarmine is a red stain that contains aluminum chloride and carmine. Aluminum is believed
to form a chelation complex with the carmine and
change the molecule to a positive charge, allowing it to bind with the acid substrates of low density, such as mucins. It is used to detect
mucin-secreting fungi and capsules of
Cryptococcus neoformans and Rhinosporidium
seeberi. It will also stain the walls of the spores
and the inner surface of the sporangia. However,
the cell walls of yeasts and Blastomyces dermatitis may stain weakly with mucicarmine
[55, 60–64].
1. Stain the smear with a working solution of
Weigert’s hematoxylin for 7 min.
2. Wash well in tap water.
3. Add metanil yellow for 1 min.
4. Repeat step 2.
5. Place in mucicarmine stain for 45 min.
6. Rinse quickly in distilled water.
7. Dehydrate in 95% ethanol and absolute alcohol (two changes of each).
8. Clear with two changes of xylene.
9. Mount in DPX and view under the
microscope.
10. Mucopolysaccharide capsule stain deep rose
to red, nuclei are black, and the other debris
stain yellow.
J. Sangeetha and D. Thangadurai
Periodic Acid–Schiff Staining
Periodic acid–Schiff (PAS) reactions are effective
stains for demonstrating fungal elements of essentially all fungi. Periodic acid attacks some carbohydrates containing 1,2-glycol or OH group with
the conversion of this group to 1,2-aldehydes,
which then react with the fuchsin-sulfurous acid
to form the magenta color [25, 32]. Identification
of fungal elements can be enhanced if a counterstain such as light green is used. Species of
Coccidioides,
Cryptococcus,
Histoplasma,
Candida, Malassezia, and Aspergillus can be
stained with this stain [65].
1. Immerse the smear in ethanol for 1 min.
2. Place 5% periodic acid for 5 min.
3. Wash gently in running tap water.
4. Place basic fuchsin for 2 min.
5. Repeat step 3.
6. Add sodium metabisulphite (0.5%) for
3–5 min.
7. Repeat step 3.
8. Counterstain with dilute aqueous light green
(0.2%) for 2 min.
9. Dehydrate with 70%, 80%, 95%, 100% ethanol and xylene, each for 2 min.
10. Observe under microscope.
11. Fungi stain bright pink-magenta or purple
against green background when light green
is used as a counterstain.
Grocott-Gomori Methenamine Silver
Staining
Grocott-Gomori methenamine silver (GMS)
staining is preferred for screening degenerated
and nonviable fungi because it provides better
contrast. The fungal cell wall contains mucopolysaccharides that are oxidized by GMS to release
aldehyde groups, which later react with silver
nitrate. Silver nitrate is converted to metallic silver, which becomes visible in the silver stains;
this is useful in detecting fungal elements. Fungi
stain in black against a pale-green background.
Pneumocystis jiroveci, Cryptococcus neoformans, Coccidiodes immitis, Histoplasma capsulatum, Aspergillus fumigatus, and Candida
albicans can be detected by this staining technique [18, 24, 55, 64, 66, 67].
19
Staining Techniques and Biochemical Methods for the Identification of Fungi
1. Add two drops of absolute ethanol for 5 min.
2. Wash in distilled water.
3. Flood the smear with 4% chromic acid for
45 min.
4. Repeat step 2.
5. Add 1% sodium metabisulphite for 1–2 min.
6. Repeat step 2.
7. Add working solution of hexamine (smear
becomes dark brown).
8. Wash with distilled water or if smear turns
black, wash with 0.1% ferric chloride.
9. Add 5% sodium thiosulphate for 2 min.
10. Repeat step 2.
11. Wash with 1% light green solution for 1 min.
12. Dry and view under oil immersion.
13. The slide with fungal elements stains black;
inner part of micelle or hyphae stains pink
with background in pale green.
Double-Oxidation Thiosemicarbazide,
Schmorl
The hydrazine group (H2NNH-) combines with
any aldehyde generated by periodic acid oxidation. The thiocarbamyl group (−CSNH2) is a
more powerful reducing agent than aldehydes
and rapidly reduces ferricyanide to ferrocyanide,
which immediately forms a prussian blue deposit
at the site. The mallory bleach lightens background staining and improves contrast. It may
also produce some aldehyde, which is removed in
step 6. This method is widely used to identify
fungal colonies in tissues [39, 68].
1. Do mallory bleach (place mellory working
solution for 10 min and rinse with water;
then place mallory solution C for 2 min until
the tissue is bleached).
2. Wash with a running tap water.
3. Oxidize in periodic acid for 10–20 min.
4. Repeat step 2.
5. Place aniline-acetic for 30 min.
6. Repeat step 2.
7. Repeat step 3.
8. Repeat step 2.
9. Place into thiosemicarbazide for 10 min.
10. Repeat step 2 to remove all traces of
thiosemicarbazide.
11. Place into freshly made Schmorl’s solution
for 10 min.
245
12.
13.
14.
15.
Repeat step 2.
Optionally, counterstain with eosin.
Repeat step 2.
Dehydrate with ethanol, clear with xylene,
and mount with a resinous medium.
16. Fungi stain with blue, nuclei with red, and
background is pink.
Fontana-Masson Staining
The Fontana-Masson (FM) stain can be used to
detect the presence of melanin in cell walls of
dematiaceous fungi such as species of Bipolaris,
Curvularia, Exophiala, and Phialophora. FM
stain is often believed to be a diagnostic tool to
differentiate
dematiaceous
fungi
from
Aspergillus sp. and some Zygomycetes. Also, it
is particularly useful for distinguishing capsuledeficient Cryptococcus neoforamans from
Histoplasma capsulatum and Blastomyces dermatitis. Melanin has the ability to reduce solutions of ammonical silver nitrate to metallic
silver without the use of an external reducing
agent. The intensity and amount of staining may
reflect differences in melanin deposition owing
to growth rate, age, availability of precursors, or
loss of pigment staining associated with hyphal
death and destruction. Extent of stain intensity
and its distribution in fungal elements in tissue
were evaluated by means of intensity; for example, dark brown (strong intensity), medium
brown (moderate intensity), and pale brown
(weak intensity) [69].
1. Treat smear with ammonical silver nitrate
solution for 20 min at 60 °C.
2. Check microscopically after 15 min and
repeat step 1 if necessary.
3. Wash well in distilled water.
4. Tone with 0.1% gold chloride for 2 min.
5. Repeat step 3.
6. Fix in 2% aqueous sodium thiosulphate for
2 min.
7. Repeat step 3.
8. Counterstain with neutral red stain for
1 min.
9. Repeat step 3.
10. Rapidly dehydrate well in absolute alcohol,
clear, and mount.
11. Observe under microscope.
246
12. All dematiaceous fungi show strong intensity (black); species such as Bipolaris,
Exophiala, Fonsecaea, and Phialophora are
darkly pigmented because of melanin.
Toluidine Blue O Staining
This stain is primarily used for the detection of
Candida albicans, Rhinosporidium seeberi, and
Pneumocystis carinii. Background staining is
removed by sulfation reagent. Yeast cells get
stained differentially and are difficult to distinguish from Pneumocystis cells. The stain can be
replaced with specific fluorescent stains.
Toluidine blue O gives polychromatic staining
for all the fungal structures (such as conidia,
germ tubes, haustoria, and hyphae) as well as
cells [55, 64, 70–73].
1. Add sulfation reagent for 10 min.
2. Wash with tap water.
3. Add toluidine blue O for 3 min.
4. Add 95% ethyl alcohol, absolute ethyl alcohol,
and xylene, each for 10 s for decolorizing.
5. Place a coverslip on the slide.
6. Observe with 20× and 40× objectives.
7. Fungi stain reddish blue to dark purple on
light-blue background.
Manual Biochemical Methods
Biochemical tests have been used to classify and
identify various groups of fungi. Because fungi
grow rapidly in pure culture, it is possible to use
biochemical methods to identify and classify
them [74]. The classical broth methods were
originally developed by Wickerham for utilization and fermentation testing of yeasts [75–77].
Biochemical methods like utilization of carbon
and nitrogen, fermentation of carbon, and enzyme
activity like caseinase, cellulase, gelatinase, glucosidase, fatty acid esterase, lipase, urease, and
so forth are currently in use to assist in the differentiation of fungi.
Utilization of Carbon Sources
Assimilation tests are extremely important in the
taxonomy of yeasts. They should be performed
before tests involving chemical analysis of the
J. Sangeetha and D. Thangadurai
fungus for the very simple reason that they are
easier to perform and generally require no specialized apparatus. An assimilation test is performed
based on the fact that nutritional factors are capable of differentiating fungi. Storage of carbohydrates fulfills multiple functions in fungi; they not
only constitute a source of carbon and energy but
also protect fungi against a variety of environmental stresses, such as desiccation and frost. Fungal
metabolism dominates the assimilation of exogenous carbohydrates into tissues. Several sugars
can be used as a carbon source. In physiologic
characterization, tests for the ability of a fungi to
utilize various carbon substrates (see Note 15) as
the sole source of carbon by employing a basal
medium such as yeast nitrogen base that contains
ammonium sulfate (universal nitrogen source),
vitamins, amino acids, and trace elements are
required for growth of yeasts. Assimilation tests
are read by growth and turbidity. For each test or
organism, negative controls without a carbon
source should be maintained [78–82].
Liquid Auxanographic Method
1. Prepare basal medium with bromocresol purple (0.5 g/L).
2. Adjust pH to 5.4 by adding NaOH or HCl.
3. Sterilize at 121 °C for 20 min.
4. Add filter-sterilized 1% (w/v) selected
carbohydrate.
5. Pour the medium into test tubes.
6. Inoculate with fungal mycelium.
7. Incubate at 20 °C for 14 days.
8. Change in color of the medium to orange or
yellow is taken as positive, whereas a change
to pink or purple is negative.
Pour Plate Auxanographic Method
1. Sugar discs can be obtained commercially or
prepared manually (steps 2–5).
2. Punch 6-mm diameter discs from Whatman
no. 1 filter paper.
3. Sterilize the discs by placing them in a hotair oven for 1 h.
4. Allow to cool, and then add one drop of 10%
filter-sterilized sugar solution to each disc.
5. Dry the disc at 30 °C in incubator and store
at 0 °C.
19
Staining Techniques and Biochemical Methods for the Identification of Fungi
6. Prepare yeast nitrogen base (YNB) medium.
7. Prepare a yeast suspension from 24- to 48-h
old culture in 2 mL of YNB by adding heavy
inoculum.
8. Add this suspension to the 18 mL sterilized
Molten agar mix well.
9. Pour the entire medium into Petri dish.
10. Allow the media to solidify at room
temperature.
11. Now place the various carbohydrate-impregnated discs onto the surface of the agar
plate.
12. Incubate at 30 °C for 4–7 days.
13. Positive reactions can be noted by growth
and color change around the disc.
Utilization of Nitrogen Sources
Simple basal media to which single nutrients
(vitamins, amino acids) could be added will be
used as the bases of these tests. For testing various nitrogen substrates (see Note 16) as the sole
source of nitrogen, one can use yeast carbon base
that contains glucose (universal carbon source)
and the vitamins, amino acids, and trace elements
required for growth. In general, if yeast can utilize nitrate, it can also use nitrate as a nitrogen
source. For each test or organism, negative controls without nitrogen source should be maintained [83–89].
Liquid Auxanographic Method
1. Prepare basal medium along with bromocresol purple (0.5 g/L).
2. Adjust pH to 4.5 with HCl or NaOH.
3. Sterilize the medium at 121 °C for 20 min.
4. Add filter-sterilized nitrogen compound
(2 g/L).
5. Inoculate with fungal mycelium.
6. Incubate at 20 °C for 4–7 days.
7. Hyphal mat in liquid media is the positive
result.
Pour Plate Auxanographic Method
1. Peptone discs can be obtained commercially
or prepared manually (steps 2–5).
2. Punch 6-mm diameter discs from Whatman
no. 1 filter paper.
247
3. Sterilize the discs by placing them in a hotair oven for 1 h.
4. Allow to cool, and then add one drop of 3%
filter-sterilized potassium nitrate or peptone
solution to each disc.
5. Dry the disc in 30 °C in incubator and store
at 0 °C.
6. Prepare yeast carbon base (YCB) medium.
7. Prepare a yeast suspension from 24- to 48-h
old culture in 2 mL of YCB by adding heavy
inoculum.
8. Add this suspension to the 18 mL sterilized
molten agar and mix well.
9. Pour the entire medium into Petri dish.
10. Allow the media to solidify at room
temperature.
11. Now place the various nitrate-impregnated
discs onto the surface of the agar plate.
12. Incubate at 30 °C for 4–7 days.
13. Positive reactions can be noted by growth
and color change around the disc.
Carbohydrate Fermentation
This method is a powerful tool for definitive
characterization and taxonomy of yeasts.
Carbohydrate fermentation tests whether a certain microbe has the capability to ferment different carbohydrates. Fungi able to ferment a
particular sugar are also able to assimilate the
same sugar; however, the reverse is not always
true. To test fermentative abilities, a different
basal medium is employed. Normally, 2% sugar
solution is added to the basal medium in a test
tube that also contains an inverted Durham tube
in order to observe production of CO2 and ethanol as the by-products of sugar fermentation.
Because most yeasts are also able to assimilate
ethanol as the sole source of carbon, it is necessary to incubate assimilation tests separately from
fermentation tests, as ethanol vapor produced by
fermentation can dissolve in assimilation tests
and cause false-positive results [90–98].
1. Prepare basal medium and sterilize at 121 °C
for 20 min.
2. Add filter-sterilized sugar at the concentration
of 2% (w/v) to the medium aseptically.
3. Pour the medium to the test tubes.
248
4. Insert inverted single sterile Durham’s tube in
each and close the lid.
5. Incubate at 20 °C for 7 days.
6. Gas accumulation in Durham’s tube is indicative of a positive result.
Casein Hydrolysis
Caseinase is an exoenzyme that is secreted outside
of the cells into the surrounding media. It has the
ability to break down milk protein, called casein,
into small peptides and individual amino acids for
their energy use or as building material. The
hydrolytic reaction creates a clear zone around the
cell as the casein protein is converted to soluble
and transparent end products, like small chains of
amino acids, dipeptides, and polypeptides. This
test can be used to identify some species of yeast
and fungi like Citeromyces matritensis, Aspergillus
dimorphicus, A. ochraceus, Fusarium illudens, F.
moniliforme, F. solani, Penicillium citrinum, P.
brevicompactum, P. chrysogenum, P. fellutanum,
and P. waksmanii [99, 100].
1. Prepare Petri plates with autoclaved skim milk
agar or casein agar in sterile conditions.
2. Inoculate fungal mycelia onto the center of the
plate and incubate at 20 °C for 14 days.
3. Examine for the presence of a clear zone.
4. The appearance of a clear zone around the
fungal colony is the positive result.
Cellulose Hydrolysis
Cellulase is produced chiefly by fungi, bacteria,
and protozoans that catalyze cellulolytic activity.
In the most familiar case of cellulose activity, the
enzyme complex breaks down cellulose to bglucose. This type of cellulase is produced by
symbiotic bacteria and fungi in the ruminating
chambers of herbivores. Three hydrolytic
enzymes, such as five endo-1,4-b-glucanases,
one exo-1,4-b-glucanase, and one or several
1,4-b-glucosidases, are involved in cellulolysis.
This can be used to identify Sporotrichum poulverulentum, Trichoderma viride, Aspergillus
niger, Peziza sp., Fusarium sp., and Penicillium
sp [101–103].
1. Prepare basal salt medium with the addition of
1% (w/v) microgranular cellulose (sigma) and
1.2% (w/v) agar.
J. Sangeetha and D. Thangadurai
2. Autoclave at 121 °C for 30 min, disperse into
Petri dishes.
3. Inoculate fungi onto the center of the agar
plates.
4. Incubate at 20 °C for 14 days.
5. Appearance of a clear zone around the fungal
colony is a positive result.
Cyclohexamide Resistance
This technique is to confirm the presence of a
possible dimorphic fungus or dermatophytes.
Determination of the resistance of isolates to
cyclohexamide is useful when screening cultures
for Blastomyces dermatitidis, Coccidioides immitis, Epidermophyton floccosum, Histoplasma
capsulatum, Microsporum sp., Paracoccidioides
brasiliensis,
Sporothrix
schenckii,
and
Trichophyton sp. All these fungi will grow in the
presence of cycloheximide at 30 °C or less, while
fungal species such as Absidia, Aspergillus,
Mucor, Rhizopus, Scedosporium and many more
are inhibited by cyclohexamide [104–106].
1. Inoculate a small portion of mold colony onto
mycosel agar with cyclohexamide and
Sabouraud agar plates.
2. Incubate at 30 °C for 7–10 days.
3. Observe for the growth of the colonies on
plate.
4. Growth on Sabouraud agar and mycosel agar
indicates resistant, and growth on Sabouraud
agar but no growth on mycosel agar indicates
sensitive. Repeat the test if there is no growth
on Sabouraud agar or mycosel agar.
Fatty Acid Esterase Activity
Fatty acid esters are cleaved by enzymes including esterase, cutinase, and lipases, which can
release free fatty acids from several sources,
including lipids, phospholipids, sterol esters,
waxes, cutin, and suberin. In this method,
Rhizopus circinans, R. microspores, Fusarium
oxysporum, R. boreas, R. thermosus, R. usamii,
R. stolonifer, R. fusiformis, and Pseudomonas
cepacia are screened by its enzyme activity
[107–112].
1. Prepare basal medium with bromocresol purple as indicator at pH 5.4.
2. Prepare 10% tween solution.
19
Staining Techniques and Biochemical Methods for the Identification of Fungi
3. Autoclave the medium and tween solution
separately at 121 °C for 30 min.
4. Add the tween solution in a ratio of 1:9 by volume to the cooled medium.
5. Pour the medium into petri dishes.
6. Inoculate the fungal mycelia onto the center of
the medium.
7. Incubate at 20 °C for 14 days.
8. Change in the color of the medium to purple is
a positive result.
Gelatin Hydrolysis
Gelatin, a protein derived from collagen, is too
large to enter the cell as a whole and, hence, the
exoenzyme gelatinase cleaves gelatin to polypeptides and then further degrades polypeptides
to amino acids, which are taken up and utilized
by the fungi. This test can be performed to differentiate between fungi that produce gelatinase
and those that do not produce the enzyme
[113–116].
1. Inoculate culture into a nutrient gelatin tubes
with a straight needle.
2. Incubate for 48 h at 22 °C.
3. Observe for liquification.
4. Liquification of the gelatin is a positive test for
gelatin hydrolysis.
b-Glucosidase Activity
The hydrolysis of cellobiose to glucose is
achieved by b-glucosidase. b-glucosidase is predominantly a cell associated or intracellular
enzyme in many fungi. This enzyme is ubiquitous among cellulolytic fungi producing hydrolytic endonucleases or cellobiohydrolases.
Activity of b-glucosidase can be detected by
growth of the test fungi on agar containing esculin (6,7-dihydroxycoumarin 6-glucosidase) or
arbutin (hydroquinone-b-d-glucopyranoside) as
the sole carbon source. Splitting of the substrate
by the enzyme yields glucose and a coumarin
product that react with iron sulfate to produce a
black color in the growth medium [117–120].
1. Sterilize cellulolysis basal medium (CBM)
supplemented with 0.5% (w/v) esculin or arbutin and 1.6% (w/v) agar.
2. Add 1 mL of sterile 2% (w/v) aqueous ferric
sulfate solution per 100 mL of CBM agar.
3.
4.
5.
6.
249
Pour the medium to the Petri dishes.
Inoculate the fungal culture into the medium.
Incubate at 25 °C in darkness for 5 days.
Development of black color in the medium
indicates the production of b-glucosidase.
Lipase Activity
Lipase catalyzes the hydrolysis of the ester bonds
of triacylglycerols, thereby releasing free fatty
acids. Lipolytic activity has been associated with
survival and pathogenicity of several fungal
species. It can be used to detect species of
Malassezia and Candida [121–124].
1. Prepare Sabouraud agar medium and sterilize
at 121 °C for 30 min.
2. Add 0.1% (v/v) n-tributylin to the medium
and pour into the Petri dishes.
3. Inoculate fungal mycelia onto the surface of
the medium and incubate at 20 °C for 7
days.
4. Occurrence of clearance in the medium is a
positive result.
Urease Activity
Urea may be hydrolyzed by some fungi with the
help of urease so that the ammonia that is liberated can be used as a nitrogen source. Phenol red
indicator is added to the broth or solid medium at
pH 8.4, and the media will turn from red color to
pink color owing to ammonia production. Urea
hydrolysis is primarily used to distinguish
Trichophyton mentagrophytes, T. sulphureum, T.
tonsurans, and T. verrucosum from T. gallianae
and T. rubrum and to identify Cryptococcus neoformans [125–128].
1. Prepare Christensen’s urea agar medium and
sterilize at 121 °C for 30 min.
2. Pour the medium into Petri dishes and inoculate fungal mycelium onto the center of the
medium.
3. Incubate the inoculated tubes at 25–30 °C for
8 days.
4. Examine the slants every 2–3 days for color
change.
5. Change in the color of the medium to pink or
purple can be taken as a positive result, while
a change to orange or yellow is considered as
a negative result.
250
Automated Biochemical Methods
Conventional methods of identifying fungi
require as long as 14–28 days for the completion
of all the biochemical tests. In recent years, it has
become important to develop rapid, automated,
and modern methods for species identification
and strain differentiation in fungi [129–138]. In
this context, the newer miniaturized fungal
identification systems such as the API 20C Yeast
Identification System (API Analytab Products,
Plainview, NY), Biomerieux Vitek Yeast
Biochemical Card (Hazelwood, MO), and Abbott
Yeast Identification System (Abbott Laboratories,
Irving, Texas) provide biochemical testing in 3–7
days with an acceptable degree of reliability [98,
139]. The biochemical tests designed in such
automated systems are those routinely followed
in conventional identification systems, except the
test for assimilation of mixture of r-hydroxybenzoic acid and protocatechuic acid, as in the Abbott
Yeast Identification System, which helps in the
identification of Candida parapsilosis and certain other yeasts [140]. These three fungal
identification systems are based on modifications
of the classic auxanographic technique of carbohydrate assimilation [141, 142]. They are easy to
use, require less preparation of reagents, and
offer a significant saving of time compared to
conventional tube tests and other currently available biochemical approaches. These commercially available kits have been widely used to
identify filamentous fungi and yeasts [143–145].
API 20C Yeast Identification System
The commercially available API 20C Yeast
Identification System is easy to use, requires less
preparation of reagents, and has been widely used
to identify both yeast and filamentous fungi. The
kit consists of 20 microtubes containing dehydrated substrates in which 19 assimilation tests
are performed. After inoculation and incubation,
the reactions are interpreted by comparison to
growth controls and use of a reference
identification table that is provided with each kit.
The system is based on modifications of the classic auxanographic technique of carbohydrate
J. Sangeetha and D. Thangadurai
assimilation. When the yeast is able to assimilate
a particular carbohydrate accompanied with a
color change, the system must also be supplemented with morphological studies. Germ tube
tests should be done in conjunction with them as
a means of obtaining a more complete profile of
the yeast cells being identified (see Note 17)
[143–145].
1. Melt the basal medium in the ampoules by
placing them in an autoclave for 2 min.
2. Place the ampules in a water bath at 50 °C and
allow to cool.
3. Dispense 20 mL of water into the incubation
tray.
4. Place the strips into incubation tray.
5. Open the ampules according to manufacturer’s
instructions and inoculate the molten medium
with an applicator stick that has touched one
or two colonies of >2 mm diameter.
6. Inoculate the strip (20 cupules of approximately 200 ml each) using a Pasteur Pipette
and then place the lid on the tray.
7. Incubate the trays at 28–30 °C for 24–72 h and
check for growth.
8. Record and compare the results with the
identification table to identify yeast (see
Note 18).
Biomerieux Vitek Yeast Biochemical Card
The disposable plastic Vitek 30-well Yeast
Biochemical Card contains 26 conventional biochemical tests and 4 negative controls. The card
is one of several testing packages used with the
Vitek System, which includes a programmed
computer, a reader-incubator unit, a filling module, a sealer module, and a printer. Identification
is generally based on conversion of biochemical
test results into nine-digit biocodes that will be
analyzed by the Vitek computer. This automated
system has been used in the identification of clinically significant yeasts, particularly Candida
albicans [146–151].
1. Use 1–3 colonies to prepare yeast suspension
in 1.8 mL saline tubes (see Note 19).
2. Adjust the suspension to a McFarland no. 2
standard using colorimeter (46–56% transmittance, 450 nm).
19
Staining Techniques and Biochemical Methods for the Identification of Fungi
3. Label the yeast cards with a marker and place
in the filling stand with a transfer tube kept in
the yeast suspension.
4. Inoculate the cards via the filling module.
5. Seal the cards via the sealer module and incubate at 30 °C for 24–48 h, depending on the
readings provided by the instrument.
6. Vitek computer converts biochemical test
results into nine-digit biocodes in printed form.
7. Most of the results are obtained after 24 h and a
few isolates may require additional incubation.
8. Identification can be accepted if the printed
result had a reliability of more than 85%.
If the reliability is less than 85%, an API
20C Yeast Identification System or any
manual biochemical tests in addition to morphological features can be used to identify
the yeast (see Note 18).
Abbott Yeast Identification System
The Abbott Yeast Identification System is an
instrumental method based on matrix analysis of
19 biochemical reactions in addition to the germ
tube test for identifying yeast. It is a disposable
plastic unit of 20 chambers containing lyophilized
biochemical substrates such as arabinose, cellobiose, dulcitol, erythritol, galactose, glucose,
inositol, lactose, maltose, melibiose, melizitose,
nitrate, r-hydroxybenzoic acid and protocatechuic acid, raffinose, rhamnose, sucrose, trehalose, urea, xylose, and negative control. This
rapid and automated system has now been widely
in use for the identification of yeasts within
20–24 h after test initiation and offers significant
saving in time compared to conventional biochemical and other currently available manual
systems [152–157].
1. Isolate yeast pure colonies from Sabouraud
agar plates.
2. Prepare the inocula and incubate for 24–48 h
at 30 °C.
3. Select colonies of a test organism and suspend in 0.05% Noble agar using a sterile
inoculating loop or cotton-tipped applicator
(see Note 20).
4. Mix the suspension on a vortex mixer and
deliver 0.2 mL into each chamber of the
Abbott IDS cartridge.
251
5. Seal the cartridge with a polyester film (see
Note 21).
6. Take the initial optical density of each reaction chamber with the MS-2, which will be
automatically stored in the computer
memory.
7. Incubate the cartridges off-line at 30 ° C for
22–25 h.
8. Reinsert the cartridge into the MS-2 for final
reading.
9. Enter the results of several other morphological and additional tests (see Note 22).
10. Take the MS-2 data printout with a record of
positive and negative biochemical tests.
11. Identify the yeast with up to five species
listed in the printout in descending order of
likelihood based on the percent likelihood
value for each species.
Notes
1. Wear gloves, goggles, and lab coat. Nitrile
gloves are suggested when working with solvents and acids. Avoid contact and inhalation
of dyes and chemicals. Reagents should be
prepared in a fume hood [72].
2. Addition of glycerol to KOH solution will
prevent crystallization from occurring in the
solution, thus enhancing the shelf life of this
reagent. It will also help in preserving KOH
preparation for a couple of days.
3. The components have to be mixed in a brown
bottle and allowed to stand for 1 month
before use. The stain must be stored at 4 °C;
otherwise the components may degrade.
4. Under fume hood, dissolve pararosaniline
with distilled water and slowly bubble sulfur
dioxide gas through the solution until solution
color begins to change. Stopper well and store
in the dark for one or two days. Add activated
charcoal and shake for about a minute if solution is not clear; filter and store at 0–5 °C.
5. Chromic acid is corrosive to skin and mucous
membranes, highly toxic, and carcinogen.
Aniline and xylene are moderate skin and
severe eye irritants, possible carcinogens,
and combustible liquids.
252
6. Mix stain in test tube and heat it in a water
bath for 2 min. Liquid becomes almost black
and syrupy. Dilute with 100 mL of 50% alcohol and let stand for 24 h and filter. Again
dilute to 1:4 with tap water for use.
7. Periodic acid solution and the stock of periodic acid (a white powder) should be kept in
dark bottles.
8. Add HCl to distilled water in a brown bottle
before adding sodium metabisulphite, cool at
5 °C, and filter. Then add 20 mL diluted HCl
(83 mL concentrated HCl/l,000 mL distilled
water) and cool to 25 °C. Add 1 g sodium
bisulphite and store in screw-top bottle in
dark for 2 days. Add 0.5 g activated charcoal,
shake intermittently for 1 h, and filter. Store
in dark-colored, tightly closed bottle in
refrigerator (expiration 5 years); pour into a
Coplin jar for further use. Solution may be
reused until it turns pink, at which time it
must be discarded.
9. Silver nitrate is toxic, and skin contact should
be avoided; methenamine is a flammable
solid and an irritant. Any spill should be
mopped up immediately with water.
10. Neutralize the ammonium silver solution
immediately after use, as it can be explosive
when allowed to dry.
11. To avoid splashing while mixing sulphuric
acid with glacial acetic acid, place the jar in
a plastic tub filled with cold water.
12. For more precision, stains like methylene
blue or Parker blue-black fountain ink or
chlorazol may be used along with KOH.
13. If background fluorescence is too bright for
fungi to be distinguished, it may be quenched
with alum hematoxylin for 1 min or potassium permanganate for 1 min. Quench immediately before the final dehydration step. This
should be done with caution because it may
reduce fungal fluorescence.
14. An epifluorescent microscope equipped with
a mercury vapor lamp and either an ultraviolet (UV) or a blue-violet (BV) excitation
filter to achieve radiation on the slide below
412 nm should be used, because the maximum absorbance of CFW is 347 nm. A
microscope with selective filters that will
J. Sangeetha and D. Thangadurai
15.
16.
17.
18.
19.
20.
21.
22.
prevent radiation below 490 nm should not
be used for CFW [56].
Glucose, sucrose, lactose, arabinose, galactose, xylose, mannose, dulcitol, ethanol, etc.
Potassium nitrate, sodium nitrate, amino
acids, urea, glycine, ammonium sulfate,
asparagine, peptone, aliphatic amines, etc.
Germ tube tests and morphological studies
should be included, as API 20C Yeast
Identification System does not include rhamnose and urea. The API yeast profiles sometimes give many different yeast identifications
for an individual isolate; this warrants supplemental tests.
For quality control, include known isolates
of Torulopsis glabrata, Candida albicans,
and Cryptococcus laurentii.
Heavy encapsulated yeasts and isolates with
extensive mycelial growth are difficult to
suspend [146].
Prepare a slightly turbid suspension that
matches with the turbidity of 0.5 McFarland
standard.
To eliminate the adverse effects of volatile
metabolic end products of adjacent reactions,
if any.
To get a comprehensive record of all the
results, including morphological observations of hyphae, chlamydospores, arthroconidia, germ tube formation, capsules, and
phenol oxidase activity [156].
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853–856
Protocol for the In Vivo
Quantification of Superoxide
Radical in Fungi
20
Konstantinos Grintzalis, Ioannis Papapostolou,
and Christos Georgiou
Abstract
The presented protocol for superoxide radical in vivo quantification
in fungal tissues is based on the quantification of 2-hydroxyethidium
(2-OH-E+) after its isolation and fluorometric quantitation. The protocol is
ultrasensitive (i.e., <1 pmol) and applicable to any kind of fungal tissue; it
can also be used for in vitro studies.
Keywords
Superoxide radical • In vivo • Fungi • 2-Hydroxyethidium (2-OH-E+)
Introduction
Superoxide radical has been implicated in many
physiological conditions, such as aging, differentiation, development, reproduction, cell cycle,
and apoptosis, as well as in pathological conditions, such as cancer, atherosclerosis, hypertension, diabetes, ischemia, and epilepsia [1]. Several
methods have been employed for its detection
(e.g., superoxide dismutase-inhibited cytochrome
c reduction, reduction of nitroblue tetrazolium,
and chemiluminescence, or even much more
cumbersome assays such as EPR). The presented
protocol has been used for the in vivo
quantification of superoxide radical in fungi [2].
It is based on the quantification of 2-hydroxyethidium (2-OH-E+), resulting from the reaction
of superoxide radical with hydroethidine (HE).
2-OH-E+ is isolated by alkaline-acetone extraction, microcolumn cation exchange and hydrophobic chromatographies, and quantified by
fluorescence (after its enzymatic destruction by
horseradish peroxidase/H2O2 system). It is ultrasensitive (i.e., <1 pmol) and applicable to any
kind of fungal tissue for the in vivo detection of
superoxide radical. It can also be used for in vitro
studies (e.g., superoxide radical production in
culturing/growth mediums) [3].
Materials
K. Grintzalis • I. Papapostolou • C. Georgiou (*)
Department of Biology, University of Patras,
University Campus, 25600 Patras, Greece
e-mail: c.georgiou@upatras.gr
1.
2.
3.
4.
Acetone
Acetonitrile (ACN)
Albumin from bovine serum (BSA)
Balance (Kern, 770/65/6J)
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_20, © Springer Science+Business Media, LLC 2013
259
260
5. Cation exchanger Dowex 50WX-8 (400)
resin
6. Centrifugal vacuum concentrator (Savant,
model SPD111V), connected to a vacuum
pump (N 820.3 FT.18, KNF)
7. Centrifuge tubes, 15 mL (ISC BioExpress,
cat. no. C-3394-1)
8. Chloroform (CHCl3)
9. Coomassie Brilliant Blue G-250 (CBB
G-250)
10. Cuvette for absorbance measurements
(quartz 45 × 10 × 10 mm, 3.5 mL) (Starna,
England)
11. Dimethyl sulfoxide (DMSO)
12. DNA from salmon testes
13. Glass tubes, 15 mL
14. Glass Pasteur pipette (internal diameter
0.5 cm, 22 cm long, by Hirschmann
Laborgeräte GmbH & Co, Germany)
15. HLB hydrophobic cartridges 30 mM, 30 mg
(Waters, Oasis cat. no. WAT 094225)
16. Horseradish peroxidase (HRP)
17. Hydrochloric acid, HCl, ³37%
18. Hydroethidine (HE or dihydroethidium)
19. Hydrogen peroxide (H2O2)
20. Methanol (MeOH)
21. Microcentrifuge clear tubes, 2 mL (VWR,
cat. no. 89000-028)
22. Micropipettes (adjustable volume pipettes)
2.5 mL, 10 mL, 20 mL, 100 mL, 200 mL, 1 mL,
and tips (Eppendorf Research)
23. Microcuvette for fluorescence measurements
(quartz 45 × 4 × 4 mm, 0.5 mL) with its FCA4
adaptor (Starna, England)
24. Nitrogen gas, 99.999%
25. pH meter (Metrohm, 827 pHlab)
26. Phosphate buffer (Na2HPO4)
27. Potassium nitrosodisulfonate (or Fremy’s
salt)
28. Refrigerated microcentrifuge (Eppendorf,
model 5417R)
29. Sodium chloride (NaCl)
30. Sodium hydroxide (NaOH)
31. Spectrofluorometer
(Shimadzu,
model
RF-1501)
32. Spectrophotometer
(Shimadzu,
model
UV-1200)
33. SPE cartridges-Extract Clean SPE Prevail
C18 1000 (Alltech, cat. no. 605430)
K. Grintzalis et al.
34. Tris
35. Trifluoroacetic acid (TFA)
36. Water (ddH2O) was purified by a Milli-Q
system (Millipore, Billerica, MA)
Methods
Solutions and Standard Curves
For applying the present protocol for the accurate
quantification of 2-OH-E+, the following stock
solutions and standard curves need to be
prepared.
Preparation of 20 mM HE Stock Solution
Prepare fresh by weighing 7.9 mg HE in a 1.5mL microcentrifuge tube and dissolving with
1 mL of DMSO. Keep the solution protected
from light and use within 1 h of its preparation.
Quantification of the HE Stock Solution by
Fluorescence
For measuring superoxide radical accurately via
the quantification of 2-OH-E+, it is crucial to
ensure that the superoxide radical trap HE is in
excess in the biological sample. This can be done
by isolating HE from the analyzed tissues (see
step 14 in the section “2-OH-E+ and HE
Purification by Microcolumn Hydrophobic
Chromatography”) and converting it into HE
concentration via an HE standard curve. The
HE-standard curve is made from a series of stock
solutions of HE (0–0.5 mM, in 60% ACN) against
their fluorescence at ex/em 370/420 nm.
Synthesis and Fluorescence Extinction
Coefficient of 2-OH-E+
2-OH-E+ is not commercially available; therefore, a standard for 2-OH-E+ is synthesized by
the reaction of HE with nitrosodisulfonate radical
dianion (NDS, Fremy’s salt) and purified by an
Alltech Prevail SPE C18 cartridge [4] as
follows:
Nds Stock Solution Preparation
1. Prepare fresh an aqueous solution of 1 mM
NDS in 50 mM phosphate buffer, pH 7.4, and
100 mM DTPA by dissolving 3.6 mg NDS in
20
Protocol for the In Vivo Quantification of Superoxide Radical in Fungi
10 mL of 50 mM phosphate buffer, pH 7.4,
containing 100 mM DTPA.
Note: Excess of NDS should be avoided because
it reacts with 2-OH-E+ decreasing its yield.
Note: NDS should be stored at 4 °C and used
within an hour of its preparation.
2. Using a quartz microcuvette (1.4 mL), adjust
baseline of the UV–Vis absorption spectrum
(from 200 to 950 nm) with 50 mM aqueous
solution of phosphate buffer, pH 7.4, containing 100 mM DTPA and run the spectrum of the
prepared NDS solution. Record the absorbance values of NDS solution at 248, 545, and
900 nm.
3. Subtract the absorbance value measured at
900 nm from the values measured at 248 and
545 nm, and with the corrected absorbance
values at 248 and 545 nm, calculate the concentration of NDS using the corresponding
extinction coefficients 1.6 × 103 and 20.8 M/
cm at 248 and 545 nm. Then, the average concentration of the NDS solution is calculated
and adjusted to 1 mM.
Synthesis of 2-OH-E+
Mix 24 mL ddH2O with 4 mL 0.5 M phosphate
buffer, pH 7.4, 4 mL 1 mM aqueous solution of
DTPA, and 200 mL 20 mM solution of HE in
DMSO. Add slowly 8.0 mL 1.0 mM NDS while
slowly stirring the solution. Incubate the solution
at room temperature for 2 h and purify the synthesized 2-OH-E+ by an Alltech Prevail SPE C18
cartridge as follows:
1. Activate the cartridge by passing 6 mL ddH2O,
followed by 3 mL ddH2O/MeOH (50/50) mixture, 3 mL of pure MeOH, and 6 mL ddH2O.
2. Load the reaction mixture for 2-OH-E+ synthesis onto the cartridge.
3. Wash the cartridge with 4 × 3 mL ddH2O, then
with 2 × 3 mL ddH2O/MeOH (50/50) mixture,
and finally with 3 × 3 mL ddH2O/MeOH
(20/80) mixture. 2-OH-E+ should elute in this
last wash step. If the eluate is still orange
(indicating incomplete elution) use more
ddH2O/MeOH (20/80).
4. Wash the cartridge with 2 × 3 mL of pure MeOH
and continue washing the cartridge until the
second band (containing E+) is eluted.
261
5. Split the fraction containing pure 2-OH-E+ in
1.5 mL-microcentrifuge clear tubes and place
them in a centrifugal vacuum concentrator,
dissolve the red pellets in 0.05 mL 0.1 M HCl,
combine them and quantify the stock solution
of 2-OH-E+ (ca. 4 mM).
Determination of 2-OH-E+ Standard Solution
Concentration
The concentration of the standard 2-OH-E+ stock
solution is determined as follows:
1. Mix 0.5 mL of aqueous solution of 100 mM
phosphate buffer pH 7.4 containing 0.2 mM
DTPA with 0.5 mL from a 100-fold dilution
(ca. 40 mM) of the standard stock solution and
transfer mixture to in a quartz microcuvette
(1.4 mL)
2. Record the UV–Vis spectrum in the range
200–800 nm (subtracting the spectrum with
the appropriate blank using 0.5 mL 1 mM HCl
instead of standard stock solution).
3. Read the absorbance values at 470 and
800 nm.
4. Correct the measured absorbance value for the
background by subtraction of the absorbance
value measured at 800 nm, calculate the concentration of standard in the cuvette using the
corrected absorbance values and the corresponding extinction coefficient 1.2 × 104 M/
cm, and then calculate the concentration of the
standard in stock solution.
Note: 2-OH-E+ can be prepared and stored for
months at 4 °C in 1 mM HCl.
Fluorescence Extinction Coefficient
of 2-OH-E+
For quantifying the 2-OH-E+ isolated from tissues,
a fluorescent standard curve is performed (using
the 2-OH-E+ standard stock solution), in the
absence and presence (for higher sensitivity) of
DNA. Specifically, to determine the fluorescence
extinction coefficient (FEC) in the absence of
DNA, prepare a series of dilutions (0–8 mM) of
2-OH-E+ in 50 mM phosphate buffer, pH 7.4 (in
1% DMSO) and measure fluorescence at ex/em
480/583 nm in a quartz cuvette. For estimating
the FEC in the presence of DNA prepare a series
of dilutions (0–100 nM) of 2-OH-E+ in 50 mM
262
phosphate buffer, pH 7.4 (in 1% DMSO) to which
55 mL/mL 2 mg/mL DNA stock solution is added,
and measure their fluorescence at ex/em
515/567 nm in a quartz cuvette. In this case, where
the FEC of 2-OH-E+ in the absence and presence
of DNA is calculated for the Shimadzu RF-1501
spectrofluorometer (with 10 nm excitation/emission slit width, at high sensitivity, and with a
quartz microcuvette of internal dimensions
45 × 4 × 4 mm, placed in its appropriate holder),
the following steps are needed:
1. FEC without DNA: Prepare various dilutions
(0–8 mM) of the 2-OH-E+ standard stock solution in 50 mM phosphate buffer, pH 7.4 (in
1% DMSO) and place 0.3 mL of each in a
quartz cuvette and measure their fluorescence
units (FU) at ex/em 480/583 nm.
For the Shimadzu, model RF-1501, the
FEC (-DNA) = 115 FU/1 mM 2-OH-E+ [5].
2. FEC with DNA: Prepare various dilutions
(0–100 nM final concentration) of the 2-OHE+ standard stock solution in 50 mM phosphate
buffer, pH 7.4 (in 1% DMSO) and add to each
one 20 mL from 2 mg/mL DNA stock solution
(prepared in ddH2O after dissolving it over
night at 0–4 °C in an ice-water bath) (DNA
final concentration 0.15 mg/mL) and place in a
quartz cuvette and measure their fluorescence
units (FU) at ex/em 515/567 nm. For example,
with the Shimadzu, model RF-1501, FEC
(+DNA) = 2,265 FU/1 mM 2-OH-E+ [5].
Note: FEC either with or without DNA needs
to be recalculated each time a different
fluorometer is used.
Protocol for 2-OH-E+ Quantification
The following steps describe the in vivo
quantification of 2-OH-E+ in fungal tissues:
Sample Treatment
1. Place 0.2 g (wet weight) fungal tissue sample
in 2 mL of its culture medium and incubate for
30–60 min with 20 mL 5 mM HE stock (final
50 mM). During incubation, HE will enter the
fungal tissue and will react with superoxide
radical in vivo, forming and 2-OH-E+.
K. Grintzalis et al.
2. After incubation, remove any bound HE and
2-OH-E+ (either externally formed in the
growth medium or contaminating the commercial HE) on the fungal tissue, by washing
it with an equal (1 g = 1 mL) to tissue wet
weight volume 10 N HCl, followed by three
times washes with equal volume ddH2O.
Discard the supernatant after centrifugation at
3,000 × g for 5 min.
Note: For yeast cells collect cells by centrifugation and wash them with 1 mL culture medium.
Discard the supernatant after centrifugation at
3,000 × g for 5 min and resuspend cells in
0.2 mL 50 mM phosphate buffer, pH 7.4.
3. Homogenize the above washed tissue (e.g.,
with liquid nitrogen) and 50 mM phosphate
buffer, pH 7.4 containing 5 mM KCN or 5
units catalase/mL (to inhibit destruction 2-OHE+ of nonspecific peroxidases and H2O2), keeping the homogenate volume as minimum as
possible (e.g., in a final volume 0.3 mL).
Note: For yeast and cell pellets resuspended in
50 mM phosphate buffer, pH 7.4 homogenization is not required.
Note: Samples should be analyzed immediately. However, the homogenates can be frozen and stored at −80 °C for 2 weeks at most.
4. Collect a small portion of the homogenate for
protein determination.
Sample Protein Determination
5. Quantify total protein amount in homogenate
by an ultrasensitive modification of the
Bradford assay.[6] Prepare a CBB-HCl reagent
by dissolving 60 mg CBB G-250 in 100 mL
1 N HCl, stirring for 30 min and filtering
through a Whatman no.1 filter paper by water
pump aspiration. This solution is stable for
months (kept light protected at 4 °C). Prepare
fresh the hydrophobic CBB-TCA reagent by
bringing 20 mL of the above CBB-HCl reagent
to 1% ethanol and 2% TCA (0.4 g solid TCA)
and adjusting its pH to 0.4 by Na3PO4. Remove
any particulates by centrifugation and use
within 1 h of its preparation. Prepare a series
of 50 mL BSA standards (100–3,000 ng BSA
made in ddH2O) or sample appropriately
diluted with ddH2O and add to it 0.95 mL
20
Protocol for the In Vivo Quantification of Superoxide Radical in Fungi
hydrophobic CBB-TCA reagent. Measure
absorbance at 610 nm against a reagent blank
containing 50 mL ddH2O in place of BSA standard. From the BSA standard curve, calculate
the protein concentration in the homogenates
using appropriate homogenate dilutions.
+
Alkaline Acetone Extraction of 2-OH-E
from Homogenates
6. To 1 volume homogenate add 9 volumes 100%
acetone and 1/100 volume 10 N NaOH (final
concentration 0.1 N NaOH). Vortex mixtures
vigorously and centrifuge for 5 min at
15,000 × g and collect supernatant.
7. Dilute the supernatant-acetone mixture to
60% acetone with ddH2O and add 1/50 volume 2.5 M Tris–HCl, pH 7.0. Adjust the pH of
the diluted mixtures to 7.0 and remove cloudiness (due to tissue material) by incubating it
on ice for 5 min and centrifuging it at 25,000 × g
and 4 °C for 5 min. Collect the clear
supernatant.
2-OH-E+ Isolation by Microcolumn Cation
Exchange Chromatography
8. Activate Dowex resin cation exchange
microcolumn by washing resin with regenerating it with 1 mL 0.1 N HCl, 5 mL ddH2O,
1 mL 0.1 N NaOH, and 5 mL ddH2O.
Note: In order to prepare the Dowex microcolumn wash (by swirling) 4 g cation
exchanger Dowex 50WX-8 (400) resin with
40 mL 100% ACN and 40 mL ddH2O.
Equilibrate (by stirring) the resin with 15 mL
0.1 N HCl for 30 min, discard the supernatant and wash (by stirring) the resin with
15 mL ddH2O for 30 min. Discard the supernatant and equilibrate (by stirring) resin with
15 mL 0.1 N NaOH for 30 min
Prepare microcolumns by packing 0.25 mL
bed volume (approx. 0.25 g activated resin)
in a glass Pasteur pipette plugged with glass
wool. After first use, the microcolumn can be
stored at RT for at least 3 months and reused
for up to ten times after regeneration.
9. Pass the 2-OH-E+ clear alkaline acetone
extract through the Dowex microcolumn at a
free flow rate. 2-OH-E+ (as well as HE) is
bound to the Dowex resin.
263
10. Wash the microcolumn in sequence with
1 mL 4 M NaCl, 1 mL ddH2O, 2 mL 100%
ACN, and 2 mL ddH2O and elute 2-OH-E+
from the microcolumn with 1 mL 10 N HCl
and dilute to 3 N HCl with ddH2O.
2-OH-E+ and HE Purification
by Microcolumn Hydrophobic
Chromatography
11. Activate and equilibrate HLB hydrophobic
cartridge by passing 1 mL MetOH and 1 mL
ddH2O at a flow rate 2 mL/min.
12. Pass the eluted 2-OH-E+ through the activated HLB microcolumn at a flow rate 2 mL/
min. Wash off impurities by passing through
1 mL 17% ACN-phosphate and elute 2-OHE+ (as well as HE and other oxidation products of HE) by passing through 1.5 mL 25%
ACN-phosphate.
13. Extract 2-OH-E+ by adding 1.5 mL chloroform, vortex and centrifuge to collect organic
bottom layer and evaporate it (e.g., using a
vacuum centrifuge evaporator).
Note: The red dry residue of 2-OH-E+ is contaminated with other oxidation products of
HE such as ethidium (E+).
Note: Samples should be analyzed immediately. However, the evaporated residues can
be frozen and stored at −80 °C for 2 weeks
at most.
14. Elute unreacted excess HE by passing
through 1 mL 60% ACN-phosphate and collect the eluate to estimate the amount of HE
present in sample by measuring its
fluorescence at ex/em 370/420 nm. Convert
the fluorescence to HE moles using the
fluorescent standard curve for HE in 60%
ACN (see Reagent setup).
Note: For accurate determination of 2-OH-E+
quantification, HE excess is required and
should be established by testing various incubation time and HE concentration conditions.
15. Clear the HLB column from any other hydrophobic interferences by washing with 1 mL
100% ACN (containing 0.1% trifluoroacetic
acid) and 1 mL ddH2O with a flow rate 1 mL/
min, and then equilibrate it by passing
through 1 mL MetOH and 1 mL ddH2O at a
flow rate 2 mL/min.
264
K. Grintzalis et al.
Note: After first use, the HLB microcolumn
can be stored at RT for at least 3 months
and reused for up to ten times after
regeneration.
Fluorometric Quantification of 2-OH-E+
16. The fluorometric quantification of 2-OH-E+
is based on measuring the total fluorescence
of concentrated 2-OH-E+ in the presence of
other contaminating oxidation products of
HE (e.g., E+) and measuring fluorescence
again after the enzymic destruction (by HRP/
hydrogen peroxide treatment) of 2-OH-E+.
Resuspend the red 2-OH-E+ residue from the
sample by dissolving it in 50 mL 50 mM
phosphate buffer, pH 7.4 containing 6%
DMSO. Add 250 mL 50 mM phosphate buffer, pH 7.4 (final DMSO 1%).
17. Measure fluorescence at ex/em 480/583 nm
(of 2-OH-E+ and other oxidation products,
designated as Total Fluorescent Units or
TFU) of the 0.3 mL 2-OH-E+ solution.
Note: If fluorescence is limiting, it can be
enhanced 25-fold at ex/em 515/567 nm by
addition of 20 mL from 2 mg/mL DNA
stock solution (DNA final concentration
0.15 mg/mL).
Record the FU in the absence/presence of
DNA which are designated as total FU
(TFU±DNA) since they result from 2-OH-E+
possibly mixed with other HE oxidation
products like E+.
18. To the above mixture (±DNA) add 25 mL
0.003% H2O2 and 10 mL 100 units/mL HRP
and incubate for 5 min at RT in order to
destroy 2-OH-E+. Measure fluorescence
resulting from all other oxidation products
except of 2-OH-E+ and correct it by multiplying with the factor 0.335/0.300 = 1.11
(to account for the dilution of the 0.30 mL
solution from HRP and H2O2).
Note: If DNA was added the correction
factor is the same (0.350/0.320 = 1.11).
This fluorescence is designated as FU.
19. Calculate the fluorescence of 2-OH-E+ which
is equal to TFU±DNA—FU±DNA, convert it to
2-OH-E+ moles using the fluorescent standard curve for 2-OH-E+ (see Reagent setup),
and express it per mg of protein.
References
1. Halliwell B, Gutteridge CMJ (1999) Free radicals in
biology and medicine, 3rd edn. Oxford University
Press, Oxford, UK
2. Papapostolou I, Georgiou DC (2010) Superoxide radical induces sclerotial differentiation in filamentous
phytopathogenic fungi: a SOD mimetics study.
Microbiology 156:960–966
3. Georgiou CD, Papapostolou I, Grintzalis K (2008)
Superoxide radical detection in cells, tissues, organisms (animals, plants, insects, microorganisms) and
soils. Nat Protoc 3:1679–1692
4. Zielonka J, Vasquez-Vivar J, Kalyanaraman B (2008)
Detection of 2-hydroxyethidium in cellular systems: a
unique marker product of superoxide and hydroethidine. Nat Protoc 3:8–21
5. Georgiou DC, Papapostolou I, Patsoukis N, Tsegenidis
T, Sideris T (2005) An ultrasensitive fluorescent assay
for the in vivo quantification of superoxide radical in
organisms. Anal Biochem 347:144–151
6. Georgiou CD, Grintzalis K, Zervoudaki G,
Papapostolou I (2008) Mechanism of Coomassie brilliant blue G-250 binding to proteins: a hydrophobic
assay for ng quantities of proteins. Anal Bioanal Chem
391:391–403
Isolation of Intact RNA from Sorted
S. cerevisiae Cells for Differential
Gene Expression Analysis
21
Jeannette Vogt, Frank Stahl, Thomas Scheper,
and Susann Müller
Abstract
Individuals within natural populations are highly diverse. They can vary
regarding their physiological states and thus contribute differently to the
performance of the whole community [1]. Hence, the characterization of
microbial physiology on the single cell level is crucial for optimization
and understanding of biotechnological processes.
Here a protocol is presented for detecting and monitoring the functional
heterogeneity in a given yeast population, on both the cellular and molecular levels, using flow cytometry and microarray analysis (first published in
Nature Protocols by Achilles et al. [2]). The protocol includes staining of
living S. cerevisiae cells with three different fluorescent dyes, flow cytometric analysis, sorting of live cells and stabilization of their status, the
subsequent isolation of RNA from small amounts of separated cells, and
finally the quantification and integrity check of this RNA. Owing to the
high quality and quantity of the isolated RNA from sorted cells, gene
expression analysis can be performed by using microarrays, for example.
Keywords
Multiparametric flow cytometry • Cell sorting • Population dynamics
• RNA isolation • Saccharomyces cerevisiae
Introduction
J. Vogt • S. Müller (*)
Department of Environmental Microbiology,
Helmholtz Centre for Environmental Research—UFZ,
Permoserstr. 15, Leipzig, Saxony 04318, Germany
e-mail: susann.mueller@ufz.de
F. Stahl • T. Scheper
Institute of Technical Chemistry, Callinstr. 5,
30559 Hanover, Lower Saxony, Germany
Microorganisms are widely used for biotechnological production processes. For example, yeasts
are employed not only to produce beer, wine, and
bread, which has been done for thousands of
years, but also for the production of vaccines [3–
5]. pharmaceuticals [6, 7], and chemicals [8–10].
The optimization of such biotechnological
processes is focused on the receipt of high
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_21, © Springer Science+Business Media, LLC 2013
265
266
amounts of desired target products with minimal
consumption of raw substrates. Commonly, the
analyses of biomass, temperature, pH value, consumption of oxygen, and formation of carbon
dioxide, for example, are used for monitoring of
the processes. However, these bulk-scale measurement methods do not afford information
about individual variations—for example, regarding size, shape, age, vitality, enzyme activity,
content of storage material, and proliferation
activity. Since the performance of a whole population is a sum of the individuals’ performances,
knowledge of specific characteristics of the individuals within the population is a valuable tool
for optimizing and understanding of biotechnological processes.
Flow cytometry is a technique that allows
analysis of the heterogeneity of cells within
microbial populations or communities quantitatively [11–14]. Other techniques, like image
analysis, fluorescence microscopy, confocal
laser scanning microscopy (CLSM), confocal
RAMAN-microscopy (CRM), or slide-based
laser scanning cytometry (LSC), afford single
cells analyses, too, but not in similar quantity
and/or rapidness. By using flow cytometry thousands of cells within a suspension are analyzed
within a few seconds and can be characterized
regarding their functional and structural parameters, such as size, shape, granularity, proliferation
activity, enzyme activity, membrane potential,
and amount of storage material like neutral lipids
and hydroxysterols (for review see Müller and
Nebe-von-Caron [14]). Hence, this technique is
suitable for studying microbial communities and
has often been used to get insights into the physiology and morphology of single cells of S. cerevisiae populations [15–26]. In addition, flow
cytometers can be equipped with cell sorters,
which separate subpopulations from each other,
showing differences in their characteristics. The
sorted cells can be used for further cultivation
and/or analysis, for example, regarding their gene
expression profile [27–30], their proteome [31–
33], or phylogenetic composition [34–37].
Consequently, flow cytometry is a useful tool to
understand biotechnological processes not only
on the single-cell level, but also on the level of
J. Vogt et al.
transcriptome and proteome by using cell sorting
and further analysis.
The yield of biotechnological production processes is dependent on the viability, growth, proliferation activity, and metabolic activity of each
single cell within the population influenced by
the availability and utilization of carbon and
energy sources. Therefore, there is a great interest in analyzing physiological states of single
cells. Of particular importance is knowledge
about the consumption of the substrate, including
the uptake and metabolization and the regulation
of these processes on the cellular and molecular
levels, even under different microenvironmental
conditions. For investigation of S. cerevisiae cell
populations well-established flow cytometric
methods are already available for certain cell
parameters [38, 39] and were used for development of control strategies in industrial processes.
However, these techniques relied mainly on measurement of proliferation activity or storage product accumulation.
In order to get an idea about the physiology of
cells and their potential to synthesize the desired
target and other products independent of the kind
and amount of the carbon source, it is advantageous to analyze the carbon consumption of the
cells. By using batch cultivation it is possible to
measure the heterogeneous substrate consumption pattern in detail and to find the best conditions for product synthesis in a simple, fast, and
low-cost way. Here, cellular glucose consumption can be followed in vital cells using fluorescent
2-NBDglucose [40]. In Fig. 21.1 a batch cultivation of S. cerevisiae H155 using a defined yeast
nitrogen base medium (2 % glucose, Difco) is
shown [40]. A number of physiological parameters like proliferation activity and cell size
were flow-cytometrically analyzed in addition to
the measurement of glucose and ethanol within
the medium as well as the optical density (see
Fig. 21.1a). As expected, during the first hours of
the batch cultivation the glucose concentration
decreased, whereas the concentration of ethanol
in the medium and the biomass measured by optical density increased. The batch cultivation
started with a high number of cells being in the
G2 phase of the cell cycle. This was caused by
21
Isolation of Intact RNA from Sorted S. cerevisiae Cells for Differential Gene Expression Analysis
267
Fig. 21.1 Batch cultivation of S. cerevisiae H155. (a)
Concentration of glucose in the medium (g/L; filled diamond), concentration of ethanol in the medium (g/L; open
triangle), optical density (700 nm; open square), cells in
the G2 phase of the cell cycle (%; filled square), and
mean values of cell size distributions (FSC; shaded circle).
(b) Cellular neutral lipid content (rel. FI; filled triangle)
and cells’ affinity to 2-NBD-glucose (molecules / cell
(x 108); shaded circle). Reprinted with permission from
Achilles J, Müller S, Bley T, Babel W. Affinity of single S.
cerevisiae cells to 2-NBDglucose under changing substrate concentrations. Cytometry A. 2004; 61: 88–98 [40]
using cells from a stationary phase, as the new
inoculum. S. cerevisiae cells taken from this
phase commonly arrest the process of cell division owing to limited substrate concentration.
Within the first 8 hours of the batch cultivation,
the cells started to divide. Consequently, the
amount of cells containing the double chromosome content as well as the size of the cells
decreased. At the ninth hour the number of G2
phase cells increased again, as a result of
replication, until the beginning of glucose
limitation. The cell size, analyzed by forward
scatter (FSC) signal of the cells, increased at the
same time owing to bud formation and growth.
The results of flow-cytometrical measurements
of the cells’ content of neutral lipids stained by
nile red as well as affinity of the cells to glucose
using the fluorescent glucose analogue
2-NBDglucose is shown in Fig. 21.1b. At the
beginning of the batch cultivation the neutral
268
lipid content within the cells declined rapidly,
obviously caused by cell division activity and the
associated high carbon turnover. Afterward the
amount of neutral lipids increased with glucose
consumption and finally glucose limitation. Yeast
cells are known for synthesizing neutral lipids
under limiting conditions to ensure their survival
[20]. These results of neutral lipid analyses were
confirmed by the side scatter (SSC) signal, which
is a measure for the granularity of the cells (not
shown). The analyses of the cells’ affinity to glucose were performed by measuring the
2-NBDglucose fluorescence intensities of the
cells. This fluorescent glucose analogue was previously used by Natarajan and Srienc [41, 42] to
analyze the glucose uptake by single Escherichia
coli cells. The higher the 2-NBDglucose
fluorescence, the higher is the cells’ affinity to
glucose. At the beginning of the batch cultivation, characterized by high extracellular glucose
concentration, the cells’ affinity to 2-NBDglucose
was very low. The cells’ affinity to glucose was
maximal at about 15 hours, when glucose was
almost exhausted, probably caused by expression
of high affinity glucose transport systems. Indeed,
during batch cultivation, subpopulations could be
determined differing in their affinities to the substrate and not correlated with the size and/or the
budding status of the cells. During batch cultivation, however, there is usually a very short time
period in which the rate of product synthesis
reaches nearly its maximum. Additionally, batch
cultivations are generally characterized by permanent concentration changes of the provided
substrates, formed metabolites and end products,
usually accompanied by changes of the pH-value,
the redox potential and the number of cells within
subpopulations differing in their age, vitality, proliferation activity or answer to stress conditions.
To study the behavior and physiological states
of the cells as well as product formation under
more constant and defined conditions, cells must
be cultivated in a chemostat under steady state or
transient state conditions. During chemostat
cultivation cellular regulations can be detected
(e.g., regulations depending on the extracellular
substrate concentration). The concentration of
sugar, especially glucose as the preferred carbon
J. Vogt et al.
and energy source of S. cerevisiae, mainly
determines the used metabolic pathways and
therefore the product formation [43]. Under oxic
conditions and extracellular glucose concentrations below 2 gl-1 glucose is exclusively metabolized by respiration via the tricarboxylic acid
cycle. Here, two different substrate flux conditions called “subcritical” and “critical” can be
distinguished according to the limited respiratory
capacity hypothesis of Sonnleitner and Käppli
[44]. Under “subcritical” substrate flux conditions the respiratory capacity is not utilized; thus,
ethanol can be used as second substrate in addition to glucose. If the oxygen demand equates to
the respiratory capacity the substrate flux is “critical.” Higher substrate concentrations result in
“supracritical” substrate flux. Under these conditions the substrate cannot be completely oxidized
and thus ethanol is produced as an overflow product by reductive metabolism. As long as the
respiratory capacity is exceeded, the overflow
products accumulate in the medium. However,
during continuous cultivation of S. cerevisiae the
yield of biomass per gram of glucose is at least
fivefold lower under respiro-fermentative conditions in comparison to complete respirative conditions [43–46]. Hence, in biotechnological
processes, yeasts should be incubated at first
under respirative conditions to generate sufficient
biomass for producing the desired target product
under respiro-fermentative conditions in the second step of the process.
In Fig. 21.2 results of flow-cytometrical analyses of transient state cultivated live cells of S.
cerevisiae stained with three different colors are
shown [47]. The three-color technique enables
the simultaneous analyses of the cell affinity to
glucose by 2-NBDglucose, the cell proliferation
activity by Hoechst 33342, as well as the dead
cell amount within the population by using propidium iodide. The physiology of S. cerevisiae
H155 cells independent of changing substrate
concentrations using transient state cultivation
with different and increasing glucose concentration is presented elsewhere [40]. In Fig. 21.2 it is
shown that during the transient state cultivation,
the affinities of cells to 2-NBDglucose were high
at low substrate concentration and low when the
21
Isolation of Intact RNA from Sorted S. cerevisiae Cells for Differential Gene Expression Analysis
269
Fig. 21.2 Transient state cultivated live cells of S. cerevisiae, stained with the three-color assay. The dead cell
amount is given in percent within the left lower quadrant
of each dot plot. For comparison, the DNA of the identical
but fixed samples were presented within the insets,
analyzed at linear scale. Data of at least 20,000 events are
displayed. Reprinted with permission from Achilles J,
Harms H, Müller S. Analysis of living S. cerevisiae cell
states—a three color approach. Cytometry A. 2006; 69:
173–7 [47]
glucose concentration started to rise in the
bioreactor (from a dilution rate of D = 0.25 h-1
onward). In addition, it was observed that the
cells’ affinities to 2-NBDglucose varied at identical extracellular glucose concentrations. This
phenomenon was observed at different substrate
concentrations. Thus, the number of cells in the
different subpopulations showing differences in
their affinities to the substrate varied at different
extracellular glucose concentrations. The results
of the flow cytometric analysis suggest that the
cell affinities to 2-NBDglucose varied to a high
degree independent of the budding status of the
cell. For comparison, the insets present the DNA
distributions of fixed cells of the same samples.
The dead cell amount within the population was
also determined over the course of the transient
state cultivation and is given in percent within
each dot plot. The knowledge of the dead cell
amount is important to exclude these cells from
further analysis, like gene expression analysis by
microarrays. In order to understand the phenomena of different cell affinities at identical substrate concentrations at the molecular level, a
method had to be established allowing the
isolation of intact RNA out of small subpopulations of living S. cerevisiae cells, stained and then
separated by a cell sorter for gene expression
analysis. The general flow chart of the established
protocol for the isolation of intact RNA from
cytometrically sorted 5 × 107 cells of S. cerevisiae
for the analysis of intrapopulation diversity of
gene expression is shown in Fig. 21.3. This protocol includes the fluorescence staining, the flow
cytometric analysis, the sorting of live yeast cells,
the stabilization of the sorted yeast cells, subsequently the isolation of RNA from the resulting
subpopulations, and finally the quantification and
the integrity check of the isolated RNA. This protocol is a slightly modified version of the protocol previously published in Nature Protocols [2].
The use of this protocol enables one to compare gene expression profiles of small numbers of
live microbial cells showing different properties.
Consequently, the protocol is useful for the examination of functional heterogeneities within
microbial populations on both cellular and molecular levels.
270
Fig. 21.3 Work flow of the isolation of intact RNA from
cytometrically sorted S. cerevisiae for the analysis of
intra-population diversity of gene expression. Reprinted with
permission from Achilles J, Stahl F, Harms H, Müller S.
J. Vogt et al.
Isolation of intact RNA from cytometrically sorted
Saccharomyces cerevisiae for the analysis of intrapopulation diversity of gene expression. Nat Protoc. 2007; 2:
2203–11 [2]
21
Isolation of Intact RNA from Sorted S. cerevisiae Cells for Differential Gene Expression Analysis
Materials
1. Schatzmann medium
2. 2-L Biostat®MD laboratory stirring bioreactor (Braun) equipped with a pH-electrode
(model 405-DPAS-SC-K8S/200; MettlerToledo GmbH) and an oxygen electrode
(model InPro 6000; Mettler-Toledo GmbH)
3. Sterile HEPES buffer (2-(4-(2-hydroxyethyl)1-piperazinyl)-ethanesulfonic acid; Merck;
10 mM; pH 7.2)
4. 2-NBDglucose (2-(N-(7-nitrobenz-2-oxa1,3-diazol-4yl)amino)-2-deoxyglucose;
Invitrogen; 20 mM in double distilled water)
5. Hoechst 33342 (2,5’-Bi-1 H-benzimidazole;
Invitrogen; 0.325 mM in HEPES buffer)
6. PI (propidium iodide; Sigma-Aldrich; 1 mM
in phosphate buffered saline pH 7.0)
7. Verapamil hydrochloride (Sigma-Aldrich;
1 mM in double distilled water)
8. Sodium azide (Merck; 10 % in distilled water
(wt/vol))
9. Glass tubes (Duran, 10 mL; treated with
Hellmanex®II (Hellma GmbH Co. KG))
10. UV-visible spectrophotometer
11. Water bath
12. Water jet pump
13. Centrifuge
14. Vortex
15. RNAseZap® (Ambion)
16. Flow-Check™ Fluorospheres (Beckman
Coulter)
17. Dry ice (dry ice to be used in direct contact
with cells must be of highest purity)
18. Sheath fluid
The sheath fluid contained 1.9 mM KH2PO4,
3.8 mM KCl, 16.6 mM Na2HPO4, and
139 mM NaCl, pH 7.0. A 10 × stock solution
of sheath fluid was used that was autoclaved
20 min at 121°C and afterward diluted with
bidistilled water filtered using a glass frit
(0.2 mm). Alternatively, a ready-to-use sheath
fluid can be purchased from Beckman
Coulter.
19. Sterile 50-mL plastic tubes
271
20. MoFlo Cell sorter (Beckman Coulter)
The flow cytometer was equipped with
two lasers for excitations at 488 nm and
multi-line UV (333–365 nm; Innova 90C and
Innova 70C from Coherent). FSC and SSC
were analyzed after excitation of 60 mW at
488 nm. For 2-NBDglucose and PI
fluorescence measurements, 530/40 and
630/30 band-pass filters were used, respectively. Detection of Hoechst 33342
fluorescence involved excitation by 40 mW
multi-line UV and a 450/65 band pass filter.
The trigger signal was FSC. The flow cytometer was equipped with a sort unit, allowing
separating subpopulations. The size of the
flow tip was 70 mm. The Summit®V.3.1 software (Dako) or updated versions (Beckman
Coulter) were used to evaluate the data.
Before analyzing and sorting the target population, the flow cytometer was calibrated
with calibration beads (Flow-Check
Fluorospheres) and a S. cerevisiae standard
with known subpopulations (e.g., ethanol
fixed and Hoechst 33342 or DAPI-stained
cells after 11 h batch cultivation showing
distinct subpopulations in the G1 and G2
phase of the cell cycle, respectively). In addition, the flow cytometer was prepared for
stable sorting by the appropriate settings (for
example the determination of the right drop
delay).
21. RNeasy Mini Kit (Qiagen)
22. Rnase-free DNAse Set (Qiagen)
23. Lyticase (from Arthrobacter luteus; SigmaAldrich)
24. b-Mercaptoethanol (Sigma-Aldrich)
25. Ethanol (³ 99.9 %)
26. Sterile RNAse-free water (diethyl pyrocarbonate (DEPC)-treated; from SigmaAldrich)
Sterile RNAse-free water was prepared
by adding 1 mL DEPC to 1 liter distilled
water and shaken until DEPC had completely
dissolved. The DEPC-water was incubated
12 h at 37 °C and then autoclaved 20 min at
121 °C to eliminate DEPC.
272
27. SG buffer (1 M sorbitol; 0.1 M ethylenediaminetetraacetate (EDTA) pH 7.4; in DEPCwater)
28. 2-mL microcentrifuge tubes
29. Microcentrifuge
30. Agarose
31. 0.5 × TAE (40 mM Tris-acetate, 1 mM EDTA,
pH 8.0)
32. Ethidium bromide (13 mM in 0.5 × TAEbuffer)
33. NanoDrop®ND-1000
spectrophotometer
V.3.1.0 (NanoDrop Technologies)
34. Gel electrophoresis system (Biometra®)
35. Gel image analysis system (software
GeneSnap V.6.08; Synoptics Ltd.).
Cultivation Conditions
Saccharomyces cerevisiae H155 was continuously cultivated in a bioreactor (e.g., 2-L
BiostatMD laboratory stirring bioreactor [Braun])
in 1 L Schatzmann medium (see later) at 30 °C.
The pH was constantly maintained at 5.4 by
adding 1 M NaOH as required controlled by a
pH-electrode (model 405-DPAS-SC-K8S/200;
Mettler-Toledo GmbH). The aeration rate was
constantly kept at 3 L of air/L of medium/min
controlled by an oxygen electrode (model InPro
6000; Mettler-Toledo GmbH) and the stirrer
velocity at 600 rpm.
Schatzmann medium was made as described
by Schatzmann [48] containing 167 mM glucose,
22 mM sodium citrate, 34 mM (NH4)2SO4,
14 mM (NH4)2HPO4, 12 mM KCl, 1.4 mM
MgSO4, 2.9 mM CaCl2, 56 mM FeCl3, 31 mM
ZnSO4, 56 mM MnSO4, 10 mM CuSO4,
333 mM myo-inositol, 126 mM Ca-pantothenate,
18 mM thiamine × HCl, 7 mM pyridoxine × HCl,
12 mM biotin; pH 5.4.
Method
Staining of the Cells
1. The cells were harvested, washed in
HEPES buffer, and diluted to 5 × 107 cells
using a calibration curve adjusted at a
J. Vogt et al.
2.
3.
4.
5.
6.
7.
8.
9.
10.
spectrophotometer (OD of 700 nm). The
staining was carried out in glass tubes since
staining in plastic tubes (e.g., microcentrifuge tubes) may influence the staining process because of dye and/or cell adherences to
the plastic surface.
5 × 107 cells were centrifuged for 5 min at
3,200 × g and 5 °C. Afterward the supernatant was carefully discarded.
25 mL of Hoechst 33342 solution (stock solution: 0.325 mM in HEPES buffer) was added
to the cell pellet for DNA staining.
25 mL of verapamil (stock solution: 1 mM in
double distilled water) was added to stabilize
the Hoechst 33342 staining, as verapamil
prevents the efflux of the Hoechst dye by
multidrug membrane transporters.
20 mL of 2-NBDglucose (stock solution:
20 mM in double distilled water) was added
to visualize the cell’s affinity to glucose.
The assay was gently mixed by pipetting up
and down three times.
The assay was incubated in a water bath at
30 °C for 20 min.
50 mL PI solution (stock solution: 1 mM in
phosphate buffered saline) was added followed by vortexing to stain dead cells.
100 mL of sodium azide (stock solution: 10 %
in distilled water (wt/vol)) was added followed by vortexing to stop the staining
reaction.
780 mL of HEPES buffer was added and the
cells were analyzed immediately by flow
cytometry. (see Note 1)
Cell Sorting
1. A ribonuclease-free working place was prepared by using for example RNAseZap.
Particular attention was paid to the sample
unit and the nozzle of the flow cytometer.
Additionally, gloves were worn to avoid ribonuclease contamination.
2. Subpopulations for sorting were selected
using the Summit V.3.1. software (or updated
versions) by appropriate gating (e.g.,
regarding the DNA content or the substrate
affinity).
21
Isolation of Intact RNA from Sorted S. cerevisiae Cells for Differential Gene Expression Analysis
3. The cells were sorted into 50-mL plastic tubes
standing in a container filled with dry ice. The
sorting was realized in the sort mode “purify
one” with a sample pressure resulting in a
sorting rate of approximately 30,000 cells/s
(coincidence rate: 4,700–5,000). This procedure was repeated until 5 × 107 cells per subpopulation were sorted. Always freshly
harvested cells were used. (see Note 2)
Isolation of RNA
For the extraction of RNA, the RNeasy Mini Kit
(Qiagen) and the protocol “Yeast II” were used
with some modifications.
1. The cells were thawed in a water bath at
60 °C for 7 min. The tube was vigorously
shaken every 1.5 min to maintain the temperature inside the tube below 12 °C.
2. The cells were centrifuged for 7 min at
6,300 × g at 5 °C.
3. The supernatant was carefully removed by
using a water jet pump with a glass Pasteur
pipette submerged in the supernatant.
4. The cell pellet was resuspended in 1 mL of
sterile DEPC-water and spun down at
1,500 × g for 4 min at 4 °C.
5. The supernatant was carefully removed and
discarded. As much liquid as possible should
be removed from the yeast pellet.
6. The cells were resuspended in 100 mL SG
buffer containing 100 Units (U) lyticase and
incubated 20 min at room temperature. The
cell suspension was gently swirled every
5 min to generate spheroplasts. (see Note 3)
7. 350 mL RLT buffer with b-mercaptoethanol
was added and followed by vigorously vortexing for 2 min to lyse the cells.
8. 250 mL of 100 % ethanol was added to provide appropriate binding conditions.
9. The sample was briefly vortexed and applied
to an RNeasy mini column. The tube was
gently closed and centrifuged for 15 s at
8,000 × g at 21 °C. The flow through tube
was discarded and the RNeasy mini column
was placed in a new tube.
273
10. The RNeasy mini column was washed by
adding 350 mL RW1 buffer provided by the
manufacturer and centrifuged for 15 s at
8,000 × g at 21 °C.
11. A DNase treatment was used to avoid contamination with genomic DNA. A DNaseI
incubation mix (10 mL DNaseI stock solution
plus 70 mL Buffer RDD provided by the manufacturer [Qiagen]) was added onto the
RNeasy mini column and incubated for 15 min
at room temperature. Afterward, the column
was spun for 15 s at 8,000 × g at 21 °C.
12. The RNeasy mini column was washed again
by adding 350 mL RW1 buffer and centrifuged for 15 s at 8,000 × g at 21 °C.
13. The RNeasy mini column was washed two
times by adding 500 mL of RPE buffer provided by the manufacturer and centrifuged
for 15 s at 8,000 × g at 21 °C, as described in
the manufacturer’s protocol.
14. To recover the RNA, the RNeasy mini column
was transferred to a new RNAse-free microcentrifuge tube. 30 mL of RNAse-free water
was added directly onto the center of the silica
gel membrane and allowed to sit for 1 min.
Afterward, the RNeasy mini column was centrifuged for 1 min at 8,000 × g at 21 °C to elute
the RNA. The elution step was repeated by
adding 20 mL RNAse free water onto the silica-gel membrane and the RNA was eluted
into the same collection tube. RNA samples
can be stored at −20 °C up to 2 months, or at
−80 °C for more than 2 months (see Note 4).
Estimation of the Quantity and Integrity
of the Isolated RNA
1. The isolated RNA was spectrophotometrically
quantified by using the NanoDrop spectrophotometer at 260 nm. The RNA purity was
assessed by measuring the 260/280 ratio and
the 260/230 ratio. A 260/280 ratio of 2.1 ± 0.15
indicated that the samples were free of protein. If the 260/230 ratio was at least 0.15
higher than the 260/280 ratio, the samples
were free of salts (see Note 5).
274
2. To perform the RNA integrity check using a
standard agarose gel, the RNA was loaded
onto a standard 1 % agarose/ 0.5 × TAE gel
and the gel was run for 1.5 h at 70 V.
Subsequently, the gel was stained with ethidium bromide for 20 min. The stained gel was
analyzed by a gel image analysis system (see
Note 6).
To check the integrity of the RNA using a
Bioanalyzer 2100 (Agilent Technologies, Santa
Clara, CA, USA) the manufacturer’s instructions
were followed.
J. Vogt et al.
3.
4.
Notes
1. The staining protocol was optimized for sort
rates of 30,000 cells/s to obtain RNA of high
quantity and integrity. Lower sort rates
required more time to obtain the 5 × 107 cells
necessary for RNA isolation resulting in lower
RNA quantitity and integrity.
The use of ethanol, methanol, formaldehyde, or formalin for fixation should be
avoided, because these substances are known
to hamper the analysis of mRNA [49, 50].
All staining solutions and equipment need
to be RNAse free.
2. The cells were always sorted in plastic tubes.
Using glass would have been resulted in tube
rupture when subjected to large temperature
changes. Sorted cells were kept frozen during
the whole sorting and collection procedure. It
was advantageous to add some splits of pure
dry ice directly into the collection tubes every
2–3 min. The tubes were changed when nearly
half-filled, to avoid splashing onto deflection
plates. Thinner collection tubes assure more
efficient freezing but will not hold the required
sample volume. Additionally, during centrifugation of multiple tubes, more cells can be
lost. The stable run of the flow cytometer was
checked by analyzing the yeast standard and
the calibration beads at regular intervals. After
every cytometer check with standards or other
interruption of sorting it was necessary to
repeat RNase activity removing step. Frozen
samples for RNA isolation can be stored at
5.
6.
−20 °C up to 2 months, or at −80 °C for longer
than 2 months.
The cell wall composition can strongly depend
on microenvironmental conditions [51] and/or
the growth phase. Concentration of lyticase
and exposure time might thus influence the
success of cell wall lysis. Therefore, the exposure time and the concentration of lyticase
need an appropriate adjustment.
Low yield of RNA can be caused by RNA
damage and degradation in the preceding
steps. Therefore, the use of RNAsecontaminated solutions and materials should
be avoided. Additionally, the sheath fluid for
flow cytometry should be prepared with
DEPC- water and RNAse should be removed
from the flow cytometer by extensively rinsing with DEPC-treated distilled and filtered
water. The solution used to elute RNA from
the silica-gel membrane (e.g., RNAse-free
water) needs a pH of approximately 7. Lower
pH could cause incomplete elution of RNA.
The manufacturer’s (Qiagen) instructions provide further help.
As RNA dissolved in unbuffered water might
have a high variance of the 260/280 and
260/230 ratios, RNA should be dissolved in
10 mM Tris–HCl, pH 7.5 for determining
these ratios. For low RNA concentrations, the
NanoDrop spectrophotometer can display
erroneous 260/280 and 260/230 ratios, due to
the detection limit of the spectrophotometer.
The detection limit regarding the RNA concentration is 2 ng/mL according to the
manufacturer.
As all nucleic acids absorb at 260 nm, a
high value does not guarantee a high quantity
and quality of the isolated RNA. The integrity
of the isolated RNA was checked by agarose
gel electrophoresis or by using a Bioanalyzer
2100. Excellent RNA integrity was indicated
by clear bands of 26S rRNA and 18S rRNA
with a 26S rRNA/18S rRNA band intensity
ratio of around 2.
In order to avoid RNA degradation during gel
electrophoresis, the loading dye solution needs
to be RNAse free. It was helpful to prepare
several tubes containing small volumes of this
21
Isolation of Intact RNA from Sorted S. cerevisiae Cells for Differential Gene Expression Analysis
solution in order to avoid contamination
during the handling. Additionally, RNAse
contamination of the gel electrophoresis system should be avoided by exhaustive cleaning
and rinsing the gel chamber and the comb with
H2O2 (1 %) and RNAseZap. Fresh 0.5xTAE
buffer should always be used.
Analyzing the gene expression of life cells is
highly difficult, especially when the cells have
to be stained, flow cytometrically analyzed, and
sorted beforehand. On the one hand, this is due
to the small RNA amounts of cells from the separated subpopulations, making any RNA extraction very difficult. Additionally, mRNA is
permanently and rapidly degraded, thus precluding long time periods for cell sorting. Hence,
a fast and well-designed RNA isolation procedure was required and several control steps were
essential. It was recommended to check the
influence of various preparation steps on the
RNA yield: cell staining, fixation, storing in
buffer, cell sorting, freezing, and recovery by
thawing.
The cellular amounts of RNA varied a lot
independent of the physiological states of the
cells. When S. cerevisiae was grown in a chemostat, the RNA content was proportional to the
growth rate: the faster the growth, the higher the
RNA content [52–54]. In addition, the yield of
isolated RNA was affected by the cell wall composition, which varied depending on both
microenvironmental conditions [51] and the
growth phase. Consequently, the concentration of
lyticase, an enzyme that breaks the yeast cell
wall, and the exposition time of cells to lyticase
was optimized for cells with different cultivation
histories.
Notably, samples must be quickly prepared,
optimally preserved, and stored to guarantee a
successful application of the protocol. The
described protocol enables one to isolate totalRNA from S. cerevisiae, which can be used for
additional purposes—for example, microarraybased analyses of gene expression profiles of
individual S. cerevisiae cells independent of the
different cells’ affinities to 2-NBDglucose.
Consequently, functional heterogeneities within
yeast populations can be examined on both the
275
cellular and the molecular level. With regard to
bioprocess optimization this is useful, assuming
that glucose—as an often-used substrate in biotechnological processes with yeast as whole cell
biocatalyst can be metabolized by different pathways, resulting in heterogeneities within a pure
yeast cell population and therefore different cell
abilities for product synthesis.
Acknowledgments We thank H. Engewald and C.
Süring for technical assistance. We thank T. Hübschmann
for technical assistance as well as for helpful discussions.
Additionally, we want to thank M. Pähler from the working group “Chip Technology” at the Institute of Technical
Chemistry of the University of Hannover. This work was
supported by the Deutsche Forschungsgemeinschaft (MU
1089/5-3).
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Quantitative PCR Analysis
of Double-Stranded RNA-Mediated
Gene Silencing in Fungi
22
José J. de Vega-Bartol, Vega Tello, Jonathan Niño,
Virginia Casado, and José M. Díaz-Mínguez
Abstract
Gene silencing in fungi produces a range of phenotypes based on the different amounts of target mRNA that are degraded by the RNAi machinery
in each transformed strain. Detection of this range of variation when analyzing groups of transformants requires a fast and sensitive method.
Quantitative or real-time PCR of reverse-transcribed target mRNA is particularly well suited for this analysis.
Keywords
Silencing • Transformation • Quantitative polymerase chain reaction •
Reverse transcription • PCR amplification
Introduction
Since its discovery in Caenorhabditis elegans
[1], dsRNA-mediated or RNAi silencing has been
widely used in many organisms, including fungi,
for gene functional analysis [2]. A main feature
of RNAi in fungi is the range of phenotypes that
can be observed in silenced or knock-down transformants [3–5] as a consequence of the variable
reduction in the amount of target RNA. The transcript abundance has to be quantified to demonstrate that gene silencing is the underlying
J.J. de Vega-Bartol • V. Tello • J. Niño • V. Casado •
J.M. Díaz-Mínguez (*)
Department of Microbiologia y Genetica-CIALE,
Universidad de Salamanca, C/Río Duero, 12, Campus de
Villamayor, Villamayor, Salamanca, 37185, Spain
e-mail: josediaz@usal.es
mechanism leading to the phenotypic effect.
Real-time quantitative polymerase chain reaction
(RT-qPCR) is the method of choice for the expression analysis of a limited number of different
samples. Among its advantages are the low template input required, due to the high sensitivity of
the method, and the high resolution, as small differences in expression between different transformants and the control can be measured. In
addition, it is less time-consuming and cumbersome than other methods, such as Northern analysis, and the cost per sample is relatively low.
RT-qPCR is a combination of two steps: (1)
reverse transcription from RNA to cDNA, followed by (2) PCR amplification of the cDNA and
quantification of the amplification products in
real time. There are different commercially available procedures to obtain a fluorescent signal
from the synthesis of product that could be measured by real-time PCR instruments [6].
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_22, © Springer Science+Business Media, LLC 2013
279
280
Quantitative detection of cDNA transcribed from
the RNA template involved in gene silencing can
be obtained using a fluorescent nucleic acid dye
as SYBR Green or EvaGreen, which undergo a
conformational change after binding to doublestranded DNA that results in an increase in
fluorescence.
Individual qPCR reactions are characterized
by the PCR cycle at which fluorescence, which is
proportional to the amount of DNA produced in
each PCR cycle, rises above a defined threshold,
a parameter known as the threshold cycle (Ct) or
crossing point (Cp). The more the target there is
in the starting material, the lower the Ct. Measured
variation is caused by both true biological variation and technical factors resulting in non-specific
variation [6, 7]. Therefore, Ct values must be normalized against the initial concentration in each
sample to correct for variability associated with
the various steps of the experimental procedure,
such as differences in the template input quantity
and quality, yields of the extraction process and
enzymatic reactions, and differences in the overall transcriptional activity of the cells analyzed.
Once the Ct is measured there are two methods to
quantify the amount of target DNA: the absolute
quantification method calculates the amount of
target DNA in the reaction by interpolation in a
calibration curve that relates Ct to known amounts
of template DNA; the relative quantification
method compares the Ct of the target DNA with
that of an endogenous control, which should be
cDNA obtained from a steadily transcribed gene.
Absolute quantification is required when a precise determination of the amount of amplicon is
desired, for example for the calculation of fungal
biomass in a host, but relative quantification is
simpler and informative enough to characterize
expression of silenced genes in fungi. Among the
several normalization methods proposed [8], the
use of expression of reference genes is currently
preferred, because they are internal controls that
are affected by all sources of variation during the
experimental workflow in the same way as the
genes of interest. However, a major problem is
that silencing may produce unexpected alterations of important pathways involving down or
upregulation of commonly used endogenous
J.J. de Vega-Bartol et al.
genes. Therefore, the assessment of the expression stability of the gene(s) to be used as internal
controls, under the experimental conditions
employed, prior to its use for normalization is of
paramount importance [9]. Some authors strongly
recommend using several endogenous genes in
parallel [10] to avoid the problems generated by
RNAi side effects in gene expression.
One of the relevant aspects to take into account
when designing RT-qPCR analysis of putatively
silenced genes is primer design. During RNAi the
RNA-induced silencing complex (RISC) cleaves
the target mRNA sequence in the region complementary to the dsRNA [11]. Complete nucleolytic
degradation of the resulting fragments is not
always guaranteed, which might result in variations of the measured expression depending on
the primer binding positions [12]. Thus,
quantification of target mRNA may lead to different results depending on the pair of primers
selected.
Software tools1 and mathematical models have
been developed to improve the accuracy and precision of RT-qPCR. A de facto standard is the DDCt
method [13, 14] based on a mathematical method
[15] dependent on cycle threshold (Ct) values and
amplification efficiencies, and lately improved to
include multiple reference genes [10]. Efforts have
focused on the improvement of the determination
of amplification efficiency because it is a known
source of errors [16], and considering a fixed
efficiency value is not acceptable. PCR efficiency
could be achieved by standard curves, but novel
quicker methods based on regression analysis of
the PCR reaction kinetics after qPCR [17] lead to
reproducible efficiency values [18].
Each step of the experimental workflow should
be meticulously standardized to avoid introducing undesirable variation in the results that cannot be eliminated by applying the final
normalization. Reverse transcription is awkward.
Comparative results demonstrate that different
RNA quantification methods produce different
data and it is prudent to measure all samples
Check www.gene-quantification.info for examples and
download links.
1
22
Quantitative PCR Analysis of Double-Stranded RNA-Mediated Gene Silencing in Fungi
using the same technique [19]. Absence of proteins, DNA contamination and inhibitors, and
RNA integrity have to be determined. Moderately
degraded RNA samples can be reliably analyzed
and quantitated, as long as amplicons are kept
short (<250 bp) and expression is normalized
against a reference gene [20, 21]. DNA contamination within the RNA and cDNA samples can be
respectively checked by the absence of
amplification and the right size product after PCR
amplification of a known gene containing an
intron. Reverse transcription yields depend on
the target, the reverse transcriptase enzyme, priming strategy, and experimental conditions. Also,
the use of random primers, oligo(dT) or genespecific primers has to be studied in each case
[6]. Each approach has advantages and no strategy always works better [22, 23].
The PCR amplification step is remarkably
reproducible under optimal conditions. As stated
before, primer design is quite important but recent
trends toward high throughput have resulted in a
reduction of the need to optimize primer concentration [6]. However, optimization of primer concentration can significantly improve sensitivity.
Melting curve analysis is a single step after
amplification, and consists in a slow decrease of
temperature that causes the melting of amplicons.
The melting temperature (Tm) is characteristic of
the size and nucleotide composition of the PCR
product: those longer and richer in G/C content
melt at higher temperatures. Melting causes a loss
of fluorescence that is quantified and represented
as a melting peak by calculating the first negative
derivate of the fluorescence. These peaks provide
the same information as DNA band visualization
in an electrophoresis gel, such as the number of
different amplicons obtained by reaction.
However, routine visualization of RT-qPCR products in agarose gels is still recommended.
Finally, we advise reading and adhering to the
recommendations proposed in the Minimum
Information for Publication of Quantitative RealTime PCR Experiments (MIQE) [24, 25], which
is a set of guidelines that describes the minimum
information necessary for evaluating RT-qPCR
experiments and ensuring the integrity of the
scientific literature.
281
Materials
Isolation of RNA
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
RNase-free 1.5-mL tubes and barrier tips.
Liquid nitrogen.
Mortar and pestle.
Commercial tri-reagent, such as Invitrogen
Trizol Reagent.
DEPC-treated water.
Chloroform (trichloromethane).
Isopropanol (2-propanol).
Absolute ethanol.
3 M Sodium acetate (NaOAc or
CH3COONa).
1.5-mL tubes centrifuge.
Speed vac or desiccator connected to
vacuum.
Determination of RNA Concentration
and Quality
1. Nanodrop spectrophotometer.
2. DEPC-treated water and distilled water.
3. Agarose gel electrophoresis reactives (agarose, TAE buffer, etc.) and equipment. Check
specific protocols.
DNase Treatment
1.
2.
3.
4.
5.
6.
RNase-free DNase I.
DNase kit buffer.
RNase-free water.
DNase inactivator.
0.2-mL RNase-free tubes.
Thermocycler or water bath.
cDNA Synthesis
1.
2.
3.
4.
5.
Random hexamers or oligo(dT)n primers.
RNase-free water.
cDNA synthesis kit buffer.
DTT.
dNTP mix.
282
6. Reverse transcriptase.
7. RNase
activity
inhibitor
(Invitrogen
RNaseOUT or Roche Protector RNase
Inhibitor).
8. Thermocycler.
qPCR Quantification and Analysis
1.
2.
3.
4.
5.
Gene-specific oligonucleotides.
SYBR Green reagent.
Molecular biology grade water.
RT-qPCR thermocycler.
Programs LinRegPCR, geNorm, and Microsoft
Excel.
Methods
Reverse Transcription from RNA
to cDNA
Isolation of RNA
In fungi, Tri-reagent2-based protocols generally
provide a better yield than column-based methods. All the handling steps have to be done placing the tubes in ice. Use RNase-free consumables
and barrier tips.
1. Grow the untransformed wild-type strain and
several transformed strains in appropriate
liquid medium. Previously, verify plasmid
insertion into the transformant genome by
PCR amplification of the promoter and a part
of the target gene. Centrifuge the culture,
wash the tissue (mycelia, spores, etc.) with
distilled water, and centrifuge to eliminate
the remaining liquid before freezing the samples under in liquid nitrogen. Samples may
be stored at −80 ºC until processing.
2. Remove fungal samples from the −80 ºC
freezer and grind 100 mg of mycelia with
mortar and pestle. Add liquid nitrogen and
grind to obtain a fine dust.
For example Invitrogen Trizol Reagent. Cat. Number:
15596-026.
http://products.invitrogen.com/ivgn/
product/15596026.
2
J.J. de Vega-Bartol et al.
3. Place the ground material in a 1.5-mL tube
and add 1 mL of Tri-reagent.
4. Repeatedly pipette to obtain a homogeneous
suspension. Place the tubes on ice.
5. Add 0.2 mL of chloroform, shake gently several times and incubate for 5–10 min.
6. Centrifuge at 4 ºC for 15 min at 12,000×g,
and transfer the supernatant to a fresh
tube.
7. Add 0.5 mL of cold isopropanol and precipitate RNA for 10 min at room temperature.
8. Centrifuge at 4 ºC for 8 min at 12,000×g.
9. Remove the liquid phase with a micropipette,
being careful not to disturb the pellet. Allow
the pellet to dry for 5–10 min in a fume
hood.
10. Dissolve the pellet in 0.4 mL of DEPCtreated water pipetting up and down.
11. Add 0.8 mL of cold phenol to precipitate
remaining contaminations, shake by hand
several times, and incubate 5–10 min in ice.
12. Centrifuge at 4 ºC for 15 min at 8,000×g and
transfer the supernatant to a fresh 1.5-mL
tube.
13. Add 2.2 volumes of cold absolute ethanol
and 0.1 vol of 3 M NaOAc to precipitate the
nucleic acids. Incubate at −20 ºC (the longer
the better). This is a good stopping point.
14. Centrifuge at 4 ºC for 20 min at 12,000×g.
15. Remove the remaining liquid with a micropipette, being careful not to disturb the pellet It
is not necessary to remove all of the
supernatants.
16. Wash the pellet with 70% ethanol.
17. Centrifuge being careful not to disturb the
pellet for 5 min at 8,000×g.
18. Remove the remaining liquid with a micropipette. Dry the pellet in speed vac for 5–10 min
until dry.
19. Dissolve the RNA pellet in 50 mL of DEPCtreated water.
Determination of RNA Concentration
and Quality
It is very important to assess the quality of the
RNA samples and accurately measure the RNA
concentration. For both purposes convenient
absorbance readings may be obtained from very
22
Quantitative PCR Analysis of Double-Stranded RNA-Mediated Gene Silencing in Fungi
small volumes by using a Nanodrop
spectrophotometer.
1. For an accurate measurement heat the
RNA for 5–10 min at 55 ºC to completely
resuspend it.
2. Set the Nanodrop to zero with 1 mL of distilled
water. Configure the system for RNA measurements and measure 1 mL of DEPC-treated
water as blank. Use 1–2 mL of each undiluted
RNA sample. Clean the pedestal after measuring each sample. Measurement of blank samples between RNA samples is not needed. In
nanodrop measurements, a ratio of absorbances at 260/280 nm around 2 is accepted as
pure RNA. Lower values indicate the presence of proteins, phenol, or other contaminants. The ratio at 260/230 nm has to be higher
than the previous 260/280 nm one for pure
RNA. Values over 2.2 or less than 1.8 indicate
the presence of contaminants. In agarose gels,
intact RNA is observed as two bands around 2
and 4 kb corresponding to 18S and 28S ribosomal RNA, respectively. The degraded RNA
appears as a lower molecular weight smear.
3. Visualize your RNA sample by agarose gel
electrophoresis in 1% gels. Dilute 1 g of agarose in 100 mL of TAE or TBE buffer.3 Heat at
70 ºC for 10–15 min a sample containing1 mg
of RNA (Optional: mix it with 0.5 vol of formaldehyde loading dye before loading on the
gel well).
DNase Treatment
DNA traces that may be present in the RNA samples should be removed by means of a DNase
treatment. It is best to use a procedure that does
not require phenol/chloroform extraction, heating or addition of EDTA. The quantities given
allow the amplification of ten target/endogenous
genes with three replicates per run (30rx).
1. Transfer a volume of sample containing 2 mg
of RNA to a fresh 0.2-mL PCR tube. Add
water for a final volume of 44 mL. Use the
same amount of total RNA in each reaction.
Optional: the addition of formaldehyde denatures the
high secondary structure of the RNA molecule for a clear
visualization.
3
283
2. Prepare a master mix with 5 mL of 10× Buffer
and 1 mL of DNase per sample.
3. Add 6 mL from the master mix to each sample
tube to sum up 50 mL.
4. Incubate at 37 ºC in a thermocycler for
30 min.
5. Add 5 mL of DNase inactivator reagent to each
tube and incubate during 5 min at room
temperature.
6. Centrifuge for 1.5 min at 10,000×g and room
temperature. Proceed immediately with the
cDNA synthesis.
cDNA Synthesis
The following information is for Roche
Transcription High Fidelity cDNA Synthesis kit4
(kit A) and Random hexamers. Differences when
using Oligo(dT)n or other common kits such as
Invitrogen Supercript III Reverse Transcriptase5
(kit B) are indicated in each step.
1. Transfer 37.6 mL from the aqueous phase
obtained in step 1.3.6 to a fresh 0.2-mL PCR
tube.
2. Add 8 mL of Random hexamers and denature
the mixture by heating the tube for 10 min at
65 ºC in a thermocycler. Also, set a program in
the thermocycler that immediately cools the
tubes to 4 ºC.
• Random hexamers primered RT showed a
smaller yield than other options in controlled experiments. However, for silenced
gene expression analysis, oligo(dT)n
should only be used with intact RNA or
experiments that require examination of
splice variants. In that case, use 4–8 mL of
oligo(dT)n (complete with water until
8 mL).
Roche cat. Numbers: 05081955001/05091284001/05081
963001. http://www.roche-applied-science.com/proddata/
gpip/3_6_8_39_1_3.html.
5
Invitrogen cat. Numbers: 18080-093/18080-044/18080085. http://www.invitrogen.com/site/us/en/home/Productsand-Services/Applications/Nucleic-Acid-Amplification
-and-Expression-Profiling/Reverse-Transcription-andcDNA-Synthesis/RT___cDNA_Synthesis-Misc/
SuperScript.html.
4
284
J.J. de Vega-Bartol et al.
Table 22.1 Quantities per
tube for preparing a master
mix for cDNA synthesis.
Kit A: Roche Transcription
High Fidelity cDNA
Synthesis kit. Kit B:
Invitrogen Supercript III
Reverse Transcriptase
Buffer
DTT
dNTP mix
Reverse transcriptase
Inhibitor
Kit A
16
4
8
4.4
2 (Protector RNase Inhibitor)
34.4
Kit B
16
4
Previously added
4
4 (RNaseOUT)
28
Table 22.2 Conditions for cDNA synthesis. Kit A: Roche Transcription High Fidelity cDNA Synthesis kit. Kit B:
Invitrogen Supercript III Reverse Transcriptase
Preincubation
Elongation
Inactivation
Cooling
Random hexamers
Oligo(dT)
Kit A
Kit B
Kit A
29°C for 10 min
25°C for 5 min
48°C for 60 min
50–55°C for 60 min
45–55°C for 30 min
85°C for 5 min
70°C for 15 min
85°C for 5 min
4°C less than 2 h. Store it at −20°C until 6 months
• For kit B use the same quantities of primers
and also add 4 mL from the dNTP mix and
6.4 mL of water.
3. Prepare a master mix with the quantities per
tube that are shown in Table 22.1. Add 34.4 mL
(or 28 mmL) to each tube for a final volume of
50 mL.
4. Set up the conditions shown in Table 22.2 in
your thermocycler of choice according to the
kit used.
5. PCR amplification of synthesized cDNA and
quantification of the amplification products in
real time.
Quantitative Polymerase Chain Reaction
In this example, we expect the simultaneous
silencing of two highly similar transcription factors [26–28]. The wild-type strain and five transformed strains will be analyzed by using three
sets of primers for the 5¢, central and 3¢ regions of
each of the target genes, and 4 endogenous genes.
Also, controls for primer contamination and RNA
contamination will be included. Our qPCR equipment accepts 96rx plates, so three plates will be
needed. Also, a calibrator (a common sample) for
each gene must be included in all the plates to
allow later normalizing for interplate variation.
Place the tubes on ice during the whole process. Keep SYBR Green that contains reagents in
darkness.
Kit B
50–55°C for 60 min
70°C for 15 min
1. Mix 1,344 mL of 2X SYBR Green Buffer with
591.36 mL of water in a 2-mL tube and vortex.
Mix 5 mL of 2× Buffer with 3.2 mL of water
for each reaction and add a 5% more to the
final volume to prevent high volumes pipetting
deviation.
2. Place six 1.5 mL tubes on ice. Mark them
1 to 6. Transfer 648 mL (90rx) to the first tube
(1) and 216 mL (30rx) to the other (2 to 6). Set
two tubes for the negative controls and mark
them. Transfer 72 mL (10rx) and 43.2 mL (6rx)
to each of them.
3. Add 180 mL of wild-type cDNA (RT product)
to the first tube (1) and 60 mL of each of the
five transformants to the other tubes (2 to 6).
Add 20 ml and 4.8 mL of water to the respective negative controls. Always add cDNA
before primers even if the number of samples
is higher than the number of primers. Use 2 mL
of RT product per reaction.
4. Place 30 tubes (3 columns and 10 rows) in a
rack on ice and mark them 1 to 30. Add 27.6 mL
from tubes 1, 2 and 3 to each tube of the first,
second and third rack columns, respectively.
5. In ten new tubes mix 12 mL of forward and
reverse 10 mM primers of each set. Add 2.4 mL
of the primers mix to the three tubes of each
row. Three tubes in the first row from the first
primer tube, second row from the second
primer, etc.
22
Quantitative PCR Analysis of Double-Stranded RNA-Mediated Gene Silencing in Fungi
6. Centrifuge the tubes for 1 min at 12,000×g.
Place all the tubes on ice in darkness.
7. Mix by pipetting. Pipette 9 mL in three contiguous wells and discard the rest. If you followed the numerical order, the first 3 wells
will containing the tube 1 mix which corresponds with the first sample and first primer,
the next 3 wells will contain the tube 2 which
corresponds with the second sample and the
second primer, and so on, as indicated in the
previous schema. Pipetting 9 mL instead of
10 mL will prevent a smaller volume in the last
well by previous-steps pipetting errors.
8. In the last plate, pipette 9.2 mL from the first
negative control tube (10rx) in 10 contiguous
wells. Add 0.8 mL form the first primer mixture in the first well and so on. Pipette 8 mL
from the second negative control tube (6rx) in
6 contiguous wells. Add 2 mL of RNA extraction of the first sample (wild-type) in the first
well and so on.
9. Briefly centrifuge the qPCR plate and load it
in the thermocycler. Set up the appropriate
program.
Data Analysis
Determine PCR Efficiency from RAW Data
1. Export the fluorescence raw data (not the Ct
values) without baseline correction. Import
the data in an Excel workbook. In most cases6
raw data can be exported as a text or cvs file
that can be imported in Excel using the import
wizard. Set a sample per row and the
fluorescence values for each cycle in columns.
Keep the first row for the cycle numeration
R = (average Ct
wild - type , gene m
For Roche Lightcycler 480 there is an import tool available in http://www.hartfaalcentrum.nl/index.php?main=
files&sub=0
6
(1–40/45) and the first two columns for the
gene and sample information.
2. With the Excel program running, start the
LinRegPCR (http://www.hartfaalcentrum.nl/
index.php?main=files&sub=0) program [17].
Select the Excel workbook with the
fluorescence raw data and appropriate range
of rows and columns.
3. Determine the baselines. Select Amplicon
group for group-based window-of-linearity and
open the Amplicon Groups tab. Make a group
for each amplicon according to the information
in the first two columns. If you place the gene
information in the first column, select “base
groups on 1st part of the sample name from the
front” and press Group Samples and Set W-o-L
(Window of linearity) per group.
4. Save the results in the Excel workbook. You
should have an average PCR efficiency value
(column N) for each gene (group).
Data Normalization
1. Prepare a new Excel workbook with the average Ct and standard deviation for each combination of sample and gene. Values for each
three replicas should be homogeneous, otherwise discard the value that clearly deviates
from the other two. Place genes in the columns
and strains (samples) in the rows. In our example that would make for 6 rows and 10 columns (plus headers). Also, insert the previous
efficiency value of each gene into the new
Excel workbook.
2. Calculate the ratio (R) between two plates of
the average Cts for each gene in the calibrator
(wild-type sample). You can accept variations
lower than 3% (1.03 > Rn > 0.97).
)Reference plate ÷ (average C t
3. Calculate the ratio (R) between two plates of
the average Cts for each gene in the calibrator
(wild-type sample). You can accept variations
lower than 3% (1.03 > Rn > 0.97).
285
t - type , gene m
)Other plate
4. Calculate for each gene the difference (DCt)
between the average Ct of a transformant and
the wild-type strain, repeat it with the other
transformants. Calculate also the DCt of the
wild-type strain, which must be 1. For each
DCt also calculate the error (E) of the two
respective deviations.
286
∆C t
J.J. de Vega-Bartol et al.
sample n , gene m
= average C t
sample n , gene m
− average C t
wild - type , gene m
(
E sample n, gene m = (SDsample n, gene m )2 + (SD wild −type, gene m )2
0.5
)
Validate Reference Genes
1. Prepare a new Excel workbook with the DCt
for each combination of sample and (only)
endogenous gene. Place genes in the columns
and strains in the rows.
2. Open geNorm plugin (http://medgen.ugent.
be/~jvdesomp/genorm/) in a new Excel. Load
the previous table with DCt values. Press
Calculate. Check the result matrix and discard
RQ sample n, target gene i = Efficiency target gene i ( ∆C t
any endogenous gene with a stability value
(M) higher than 1.5.
sample n , target gene i
Quantify Relative Expression
Values
1. Check that your Excel workbook includes the
efficiency values of all the genes, the DCt and
prolongated error of all the gene and sample
combinations, and that unstable endogenous
genes have been discarded.
2. Calculate the relative expression of a target
gene with respect to the calculated mean of
the endogenous genes for each of the samples.
For each relative expression value, calculate
the expanded error (EE) of the DCt (target
gene and n endogenous genes).
) ÷ ∏ j (Efficiency endogenous gene j ( ∆Ctsample, endogenous gene j ))
(
EE sample n, target gene i = (E sample n, target gene i )2 + (E samplen, endogenous gene 1 )2 + ⋯ + (E sample n, endogenous gene j )2
0.5
)
3. Calculate the minimum and maximum relative expression considering the relative
expression error (EE).
RQ max
RQ max
sample n , target gene i
sample n , target gene i
(
(∆C
= Efficiency target gene i ∆Ct
= Efficiency target gene i
sample n , target gene i
t sample n , target gene i
)
(
+ EE sample n, target gene i ÷ ∏ j Efficiency endogenous gene j ( ∆Ct
)
(
− EE sample n, target gene i ÷ ∏ j Efficiency endogenous gene j ( ∆Ct
4. Express the result as a value and a range in the
form RQsample, target gene (RQ minsample, target gene RQ
minsample, target gene). Notice that though wild-type
relative expression is 1, its EE is not 0, so its
expression can also show a range of values.
5. dsRNA-mediated silenced transformants should
show a clear reduction of the relative expression of the analyzed gene with respect to the
wild-type strain. Analyze the results obtained
for each gene expression separately, as it is
likely that results vary depending on the primer
binding place in the target mRNA. If the results
obtained when using a certain set of primers
show a higher expression in the putative
silenced strains than in the wild-type strain discard them, as it can be the result of incomplete
degradation of the flanking mRNA regions.
sample , endogenous gene j
sample , endogenous gene j
)
)
)
)
References
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SE, Mello CC (1998) Potent and specific genetic
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73:962–970
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6. Nolan T, Hands RE, Bustin SA (2006) Quantification
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Semi-Nested PCR Approach
to Amplify Large 18S rRNA Gene
Fragments for PCR-DGGE Analysis
of Soil Fungal Communities
23
Miruna Oros-Sichler and Kornelia Smalla
Abstract
Denaturing gradient gel electrophoresis (DGGE) of 18S rRNA gene fragments PCR-amplified from total community DNA is a powerful tool for
the parallel comparative analysis of environmental fungal communities.
The 18S rRNA gene has the advantages of universality, high phylogenetic
information content, and a large number of sequences in the data banks.
The comparative analysis of soil fungal communities from large numbers
of samples by PCR-DGGE requires consistent amplification and separation efficiency, as achieved by the following semi-nested PCR-DGGE protocol based on two-step PCR of 1,650 bp rRNA gene fragments from bulk
soil DNA and their separation in DGGE.
Keywords
Denaturing gradient gel electrophoresis (DGGE) • PCR-DGGE • 18S rRNA
gene fragments • Soil fungal communities
Introduction
M. Oros-Sichler
Julius Kühn Institut, Institute for Epidemiology
and Pathogen Diagnostics, Zimmerstrasse 26,
Braunschweig, Lower Saxony 38106, Germany
K. Smalla (*)
Julius Kühn Institut, Federal Research Centre
for Cultivated Plants, Institute for Epidemiology
and Pathogen Diagnostics, Messeweg 11-12,
Braunschweig, Lower Saxony 38104, Germany
e-mail: kornelia.smalla@jki.bund.de
Along with bacteria, fungi are involved in soil
functionality, comprising physical, chemical, and
biological aspects [1]. As components of the
complex interactive soil food web, fungal assemblages in soil respond to changes of the different
soil trophic levels with community changes,
which, in turn, affect the soil properties. Therefore,
fungal community shifts may serve as indicators
for soil food web modifications and their analysis
is of crucial importance for the understanding of
soil ecosystems [2].
For a long time the soil fungal community
composition was studied only by methods based
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_23, © Springer Science+Business Media, LLC 2013
289
290
on isolation of fungi directly from environmental
samples plated onto nutrient media. However, the
isolation techniques are very fastidious and
confined by the boundaries set by unculturability
of many fungi. The analysis of the community
structure and dynamics of fungal communities
from soils achieved important advances in the
past two decades thanks to the molecular techniques [3]. Analysis methods based on PCR
amplification of marker gene fragments from
total DNA extracted from environmental samples
brought forth suitable approaches to analyze
comparatively high numbers of samples in a rapid
and efficient manner.
The genes of the ribosomal gene complex,
consisting of the small subunit (SSU) 18S rRNA
gene, the large subunit (LSU) 28S rRNA gene, the
internal transcribed spacer (ITS), and the intergenic spacer (IGS), are frequently used in fungal
community profiling [4]. These marker genes
comprise both highly conserved domains and
variable regions [5, 6], allowing the design of suitable primer systems and the high resolution analysis of fungal communities at taxonomical levels
ranging from phylum to strain [7, 8]. The fungal
sequence data bases are considerably informative,
especially for intensively studied taxonomic
groups (e.g., arbuscular mycorrhizal fungi) [9].
Several primer systems, group specific or fungal universal, were designed to amplify either
SSU, LSU rRNA gene fragments or the ITS/IGS
regions and used for the characterization of fungal diversity in soils [10]. Theoretical and practical evaluation of primers targeting the 18S rRNA
gene or the ITS region revealed that some of them
amplify also nonfungal sequences, or other primers exclude major fungal taxa. Four different
primer pairs were tested for their specificity
toward fungal rRNA genes and their suitability
for assessing the diversity of fungal communities
in grassland soils [11]. Based on cloning and
sequencing of amplicons obtained with each
primer system from soil DNA, the authors concluded that primer biases might be less significant
than previously expected.
Subsequent to the PCR amplification, the
amplicon pools can be identified taxonomically
M. Oros-Sichler and K. Smalla
by sequence analysis or separated by means of
molecular profiling methods that exploit the differences in their DNA sequence or conformation
and result in taxonomically anonymous
fingerprints that allow comparisons between different sample types.
Primer systems targeting fungal rRNA genes,
coupled with molecular fingerprinting techniques
such as denaturing gradient gel electrophoresis
(DGGE) to analyze PCR products obtained from
total community DNA, provide appropriate strategies for descriptive and comparative analysis of
soil fungal community structure [12]. Due to the
fact that the fungal rRNA genes are more conserved than bacterial 16S rRNA genes, the molecular fingerprints obtained for 18S rRNA gene
fragments are less complex and, thus, easier to
evaluate than the bacterial profiles. If different
taxa contribute to the same band in the DGGE
fingerprints [13, 14], the analysis resolution can
be improved by using taxon-specific primers (e.g.,
for Trichoderma community composition) [15].
The specificity of the primer system and
the phylogenetic information contained within
the amplified fragments are decisive factors for
the degree of resolution with which the community structure is revealed. The fungal universal
primer NS1 [7], combined with the fungusspecific reverse primer FR1 [8], amplifies about
1,650 bp of the 18S fungal rRNA gene and thus
allows the use of the most phylogenetic information contained by this gene. Initially designed
and used for the study of wood-inhabiting fungi
[8], this primer system was used later in only a
few studies—for example, to investigate the fungal communities associated with the bulk and
rhizosphere soil of maize from tropical climate
[14] and to compare the fungal community structure under different agricultural practices in soil
mini ecosystems [16] or in the endorhiza of different potato lines at two different sites [17].
These studies asserted the reproducibility of this
system and suitability even for a soil with high
contents of humics [16]. However, when analyzing different soils originating from 36 sites with
known properties, we encountered difficulties in
obtaining PCR amplicons from total community
23
Semi-Nested PCR Approach to Amplify Large 18S rRNA Gene Fragments…
DNA of a broad sample range using directly the
primer combination NS1-FR1-GC [18].
The literature mentioned several different factors affecting the efficiency of DNA amplification
from soil samples. For example, insufficient
amount or low priming availability of the template DNA is known as a limitation for successful
amplification [19, 20]. Also, co-extracted humic
substances were often reported to inhibit the yield
of amplicons from soil samples [19, 21]. Both the
proportion in which the fungal DNA contributes
to the total community DNA extracted, and the
amount of co-extracted contaminants (e.g., humic
acids), cannot be assessed by agarose gel electrophoresis and might reduce the amplification
efficiency [19]. In addition to that, it was already
speculated in the literature that the GC-clamp
necessary for the DGGE analysis [22] might bias
direct PCR amplification [23, 24].
Nested PCR approaches have the advantage of
enhanced sensitivity, allowing the detection of
problematic DNA (e.g., low target amount, high
contaminant amount) and the reduction of
nonspecific amplification [25]. Moreover, the
GC-clamp necessary for DGGE analysis can be
included without difficulties in the second PCR
step, after the specific templates reached a
sufficient amount in the first PCR step.
Therefore, we designed the semi-nested PCR
protocol presented in this chapter, which consists
of a first amplification with the novel primer
combination NS1-EF3, followed by a second
amplification with the primers NS-FR1-GC [8,
18]. EF3 was designed and used formerly in other
primer combinations [26–28]. The semi-nested
PCR protocol presented in this chapter was used
successfully for the comparative analysis of
soil fungal communities from 36 different sites
[18], as well as for the study on the impact of the
site, the sugar beet cultivar, the seasonal dynamics, and the rhizosphere effect on the fungal community structure at three different sites planted
with sugar beet [29].
However, the semi-nested procedure presented
in this chapter is far from free of limitations.
For example, inconsistent PCR amplification
was reported when using the protocol presented
in this chapter, resulting in a high variability of
291
the replicate DGGE patterns when investigating
fungi of the rhizosphere of strawberry and oil
seed rape at different sites [30]. The problem was
partially solved by these authors by switching to
a nested PCR version, replacing in the first PCR
step the primer NS1 with NS0, which is located
upstream from NS1.
Furthermore, two different studies reported on
the retrieval of nonfungal sequences when working with the PCR protocol presented in this chapter. Firstly, sequences of ubiquitous soil flagellates
were retrieved in the analysis of fungal communities from bulk soils of three different sites [14].
Secondly, it was impossible to compare fungal
communities from the gut of Diabrotica virgifera
feeding on maize roots in different soils with the
protocol in this chapter, as the DGGE patterns
generated were dominated by a band of D. virgifera [31].
Last but not least, because of the relatively
high conservation of the 18S rRNA gene within
the fungi, some of the fragments might not contain enough variation to allow DGGE separation
and thus migrate with similar electrophoretic
mobility, as previously observed [13, 14].
Therefore, the ITS should be mentioned here
as a valuable alternative marker for the analysis
of soil fungal communities. The use of ITS fragments, complementary or independently to the
analysis of 18S rRNA gene fragments, might
allow DGGE separation up to an intraspecific
level [32]. Thus, additional insights might be
gained for comparative studies (e.g., when analyzing potential effects of transgenic crops on the
microbial communities in comparison with nontransgenic lines) [33]. The number of ITS entries
in the GenBank (32,050) exceeded already by
June 2005 the number of submitted 18S rRNA
gene fragment sequences (30,651) [34] and
attained 50,956 fully identified and 27,364
insufficiently identified ITS sequences as of
February 2008 [35]. Several PCR-DGGE protocols based on ITS fragments are available for the
analysis of soil fungal communities, of which a
semi-nested protocol with the primers ITS1 and
ITS4 in the first PCR and the primers ITS1-GC
and ITS2 in the second PCR was described
recently in detail [36].
292
Equipment and Materials
Equipment
1. FastPrep® Instrument FP 120 for bead beating (BIO101, Carlsbad, California).
2. Microcentrifuge.
3. Vortex.
4. Magnetic stirrer.
5. Electrophoresis chamber with power supply
and accessories.
6. Thermocycler.
7. DGGE DCode™ Universal Mutation
Detection System (Biorad, München,
Germany) and accessories.
8. Gradient maker with peristaltic pump.
9. Gel documentation system with UV transilluminator and camera (e.g., UV System,
INTAS®, Mitsubushi Electric Corporation).
10. Fast DNA®Spin®Kit for Soil (BIO101,
Carlsbad, California).
11. GENECLEAN®SPIN®
Kit
(BIO101,
Carlsbad, California).
12. 1 kb molecular weight DNA marker (e.g.,
Invitrogen).
13. Ethidium bromide.
14. Agarose
15. AmpliTaq DNA Polymerase Stoffel Fragment
(Applied Biosystems, Foster City, California).
16. Deoxynucleotide Triphosphate Set (Roche
Diagnostics, Germany).
17. Primers (Table 23.1).
18. 2% dimethyl sulfoxide (DMSO).
19. 0.5 M EDTA pH 8.
20. Deionized formamide (Stir slowly for about
30 min 10 g Serdolit MB-1 and 1 L formamide. Filter through Whatman paper to
remove ionic exchange resin. Store at -20 ° C
as 50 mL Falcon tube aliquots).
21. 18% denaturant 7.5% acrylamide stock
solution (see Notes 1, 2).
22. 38% denaturant 7.5% acrylamide stock solution (see Notes 1, 3).
23. 10% Ammonium peroxodisulfate (APS)
(w/v) in MilliQ water (stored as aliquots at
-20 ° C).
24. Tetramethylethylendiamine (TEMED).
M. Oros-Sichler and K. Smalla
25. MilliQ (deionized) water.
26. 5× TBE Buffer (27.5 g boric acid, 54 g Tris
base, 20 mL 0.5 M EDTA pH 8.0 in 1 L distilled water).
27. 50× TAE Buffer: (242.2 g Tris base, 18.6 g
EDTA, 57.1 mL acetic acid in 1 L distilled
water; diluted 1:50 for DGGE run).
28. 6× DGGE loading buffer (25 mg bromophenol blue, 25 mg xylene cyanole, and 3 mL
glycerol in 10 mL distilled water, stored
at 4 ° C).
29. DGGE standard (see Note 4).
30. GelBond film (Lonza, Switzerland).
31. Reaction vials (1.5 and 2.0 mL).
32. Pipette tips (10, 20, 100, 200, 1,000 mL) and
capillary pipette tips.
33. Syringe needles.
34. 15 mL polypropylene Falcon tubes.
35. DGGE fixative solution (10 mL acetic acid
and 200 mL ethanol in 1,790 mL MilliQ
water).
36. DGGE staining solution (0.2 g silver nitrate
in 100 mL MilliQ water, made freshly for
each gel).
37. DGGE developing solution (400 mL 37%
formaldehyde in 100 mL 1.5% sodium
hydroxide, made freshly for each gel).
38. DGGE stopping solution (7.5 g of sodium
carbonate in 1 L MilliQ water).
39. DGGE conservation solution (250 mL ethanol and 100 mL glycerol in 650 mL MilliQ
water).
Methods
Initial Material
Total community DNA extracted directly from
replicate composite bulk soil samples taken from
random plots at different sites (see Notes 5–8).
Semi-Nested PCR
The First Amplification Step
The primer combination NS1-EF3 (see
Table 23.1) is used, amplifying almost the entire
23
Semi-Nested PCR Approach to Amplify Large 18S rRNA Gene Fragments…
293
Table 23.1 Primers used for the semi-nested PCR amplification of 18S rRNA gene fragments from bulk soil total
community DNA (GC-clamp underlined)
Primer
NS1
EF3
FR1-GC
Sequence
5¢-GTA GTC ATA TGC TTG TCT C-3¢
5¢-TCC TCT AAA TGA CCA AGTTTG-3¢
5¢-CCC CCG CCG CGC GCGGCG GGCGGG GCG GGG GCA CGG GCC
G-AIC CAT TCA ATC GGT AIT-3¢
References
White et al. [7]
Smit et al. [26]
Vainio and Hantula [8]
Fig. 23.1 Position of the annealing sites of the primers used in this chapter (NS1, FR1, EF3) on the 18S rRNA gene of
Saccharomyces cerevisiae and length of the amplicons generated with different primer combinations
18S rRNA gene (Fig. 23.1). Perform the reaction
with ca. 15–20 ng DNA extract in 25 ml volume
containing: Stoffel buffer (10 mM KCl, 10 mM
Tris–HCl pH 8.3), 0.2 mM deoxynucleoside
triphosphates, 3.75 mM MgCl2, 2% (w/v) dimethyl sulfoxide (see Note 9), 0.2 mM of each
primer, and 2 U/ml of Taq DNA polymerase
Stoffel fragment. PCR cycling program: 5 min
denaturation at 94 ° C, followed by 25 thermal
cycles of 30 s at 94 °C, 45 s at 47 °C, 3 min at
72 °C, and final extension at 72 °C for 10 min.
0.2 mM deoxynucleoside triphosphates, 3.75 mM
MgCl2, 2% (w/v) dimethyl sulfoxide, 0.2 mM of
each primer and 2 U/ml of Taq DNA polymerase
Stoffel fragment). PCR cycling program: 5 min
denaturation at 94 ° C, followed by 20 thermal
cycles of 30 s at 94 ° C, 45 s at 48 ° C, 3 min at 72 °
C, and final extension at 72 °C for 10 min.
The Second Amplification Step
The primer combination NS1-FR1-GC (see
Table 23.1) is used, amplifying 1,650 bp of the 18S
rRNA gene (see Fig. 23.1). Perform the reaction
with optimized dilutions of the amplicons from the
first PCR step in 25 mL volume containing Stoffel
buffer (10 mM KCl, 10 mM Tris–HCl pH 8.3),
See Figs. 23.2a,b and Note 10.
DGGE Fingerprinting of Soil Fungal
Communities
Gel Casting
Assembly of the gel sandwich:
1. Place the glass plates on a plane table.
Carefully clean the surface of the glass plates
with 97% ethanol.
294
Fig. 23.2 (a) DGGE fungal community fingerprints of
18S rRNA gene fragments amplified from bulk soil DNA
from three sites with different soil types: Lanes 1–4, replicates from Brackstedt (sandy soil); lanes 5–8, replicates
from Niedernjesa (alluvial silt); lanes 9–12, replicates
Rötzum (clay-rich blacksoil) loess loam; lane S, standard
mixture of PCR-amplified 18S rRNA gene fragments of
fungal isolates; lanes KB and KW, standard mixtures of
PCR-amplified 18S rRNA gene fragments cloned from
M. Oros-Sichler and K. Smalla
total soil DNA of similar soils. (b) Dendrogram based on
the Pearson correlation indices and UPGMA cluster analysis of the fungal community fingerprints of 18S rRNA
gene fragments amplified from bulk soil DNA from
Brackstedt, Niedernjesa, and Rötzum. The differences
between the profiles are indicated by percentage of similarity. Patterns of soil samples originating from different
site clusters separately
23
Semi-Nested PCR Approach to Amplify Large 18S rRNA Gene Fragments…
2. Spread a couple of tap-water drops on the
large glass plate.
3. Place the GelBond film with the hydrophobic
side in direct contact with the large glass plate,
ensuring the perfect alignment of the film to
the bottom of the glass.
4. Fix the film to the glass free of air bubbles
(e.g., with a Drigalski spatula). Remove excess
tap water.
5. Place two spacers at the outer sides of the
large glass plate and the small glass on the
top.
6. Insert the glass plate assembly within sandwich clamps ensuring that the bottoms correspond perfectly.
7. Place the sandwich assembly in a casting stand
with a rubber strip at the bottom to prevent
leakage. Ensure tight contact of the sandwich
assembly bottom with the rubber strip and a
stable position of the casting assembly. Close
both clamps with equal pressure using the
alignment card. Do not over-tighten clamps to
avoid usage of plates.
8. Insert the comb in the glass plate sandwich.
Casting of denaturing gradient gels and
polymerization:
1. Thaw and keep denaturing stock solutions on
ice.
2. Add 45 mL 10% APS and 26 ml TEMED to
each 15 mL of 18% respectively 38%
denaturing solution and mix by inverting the
vials. Work on ice to prevent premature
polymerization.
3. Place the gradient maker on a stir plate at
speed 300 rounds/min with a magnet stirrer in
the outlet port chamber.
4. Connect the gradient maker to the peristaltic
pump (the pump is off and the gradient maker
channel is closed). Provide the pump tube with
a syringe needle. Insert the needle centrally
between comb and small glass plate.
5. Pour the 38% denaturing solution into the outlet port chamber of the gradient maker. Briefly
open and close the valve to remove air between
chambers. Pour the 18% denaturing solution
in the remaining chamber.
6. Turn on the peristaltic pump and open simultaneously the valve between chambers.
295
An optimal flow of 5 mL/min is recommended.
Ensure the solutions flow without leaking
from the sandwich until air bubbles reach the
syringe needle.
7. After gel casting, remove the needle and
water-flush the gradient maker and tubing to
discard solution rests.
8. Let the gel polymerize unmoved for at least
1 h. Use the same day or keep at 4 ° C wrapped
in wet towels.
Pre-Run
1. Insert two gel sandwiches into the electrophoresis core. If only one gel is used, replace second gel with a glass plate sandwich without
spacers.
2. Place the core assembly into the buffer tank
filled with 1× TAE buffer. Renew 50% of the
buffer between runs. Check buffer level, set
up the temperature for 58 °C and start the
pump.
3. When the buffer reaches the run temperature,
turn off the system and remove the comb from
the gel.
Sample Loading and Electrophoresis
1. Adjust the volume of PCR products to load at
similar DNA concentrations. The different
concentrations of the samples can interfere
with software analysis of the gel.
2. Mix PCR products 1:1 with DGGE loading
buffer and load with microcapillary pipette
tips. Ensure that a maximum of 20 mL of sample is loaded to prevent well overflowing.
3. Load a DGGE standard in the outside lanes to
assess the gradient formation and the band
positions and to normalize the gel in further
software analysis.
4. Close the system; check buffer, temperature,
and pump; and start electrophoresis.
Gel Staining, Drying, and Scanning
1. Work on a switch rocker.
2. Transfer the gel in a recipient with 100 mL
fixative solution for 10 min. or unmoved
overnight.
3. Discard the fixative solution and add 100 mL
0.2% silver nitrate fresh solution for 15 min.
296
4. Discard the silver nitrate solution in a specific
waste. Wash the gel at least twice for 1 min
with MilliQ water.
5. Change the gel in a new recipient and add
100 mL fresh developing solution.
6. Discard the developing solution as pale bands
appear. Add 100 mL stopping solution for ca.
10 min.
7. Discard the stopping solution. Add 100 mL
conservation solution for at least 7 min.
8. Transfer the gel on a rigid frame. Cover the
gel without air bubbles with a cellophane
film soaked with conservation solution. Fix
with clamps. Air-dry at room temperature for
2 days.
9. Transform the gel image in a digital picture
using any transparency scanning system
available.
10. Analyze the digitalized gel by means of software—e.g., GelCompar 4.0 (Applied Maths,
Ghent, Belgium)—with unweighted pair
group method using arithmetic averages
(UPGMA) cluster analysis.
11. Apply any statistical method available to
ensure statistical significance of results (see
reference [37]).
Notes
1. Consider that 100% denaturant solution contains 40% deionized formamide and 7 M
urea.
2. 18% denaturant 7.5% acrylamide stock solution: Dissolve 18.93 g urea in 100 mL MilliQ
water. Add 5 mL 50× TAE, 18 mL deionized
formamide, and 62.5 mL acrylamide
Rotiphorese Gel 30 (37.5:1) (Roth, Germany).
Adjust the volume to 250 mL in a volumetric
flask and filter. Aliquot 15 mL solution in
Falcon tubes. Store at -20 °C.
3. 38% denaturant 7.5% acrylamide stock solution: Dissolve 39.94 g urea in 100 mL MilliQ
water. Add 5 mL of 50× TAE, 38 mL deionized formamide and 62.5 mL acrylamide
Rotiphorese Gel 30 (37.5:1) (Roth, Germany).
Adjust the volume to 250 mL in a volumetric
flask and filter. Aliquot 15 mL solution in
Falcon tubes. Store at -20 ° C.
M. Oros-Sichler and K. Smalla
4. The DGGE standard is an artificial mixture
of 18S rRNA gene fragments PCR-amplified
from single fungal isolates or clones known
to migrate with different electrophoretic
mobilities in DGGE protocol used.
5. Ensure at least four replicate samples per site
for representative statistical results.
6. Dig a number of bulk soil cores per plot representative for the plot dimensions; e.g.,
8–10 cores (15–20 cm of top soil) per 10 m2.
Avoid root material if bulk soil analysis is
intended. Mix well by sieving.
7. Ensure an amount of 0.3–1 g dry weight of
soil per replicate for DNA extraction to minimize eventual heterogeneous distribution of
fungal DNA targets and to ensuring representative results.
8. For total DNA extraction, use one of the
commercial kits for soil, preferably BIO101
Fast DNA®Spin®Kit for Soil, combined with
a harsh cell lysis to break fungal cell walls,
e.g., with the FastPrep® Instrument for 1 min
at 4,000 rpm. Purify the crude DNA from
eventual humic contaminants, e.g., with the
GENECLEAN®SPIN® Kit. Store DNA
extracts at -20 °C until further procedures.
9. Dimethyl sulfoxide in the reaction mixture is
known to enhance PCR by eliminating
nonspecific amplification and to improve the
primer annealing efficiency by destabilizing
secondary structures within the template.
10. All DGGE materials, gel casting procedures,
and running conditions presented in this
chapter are strictly referred to the DGGE
DCode™ Universal Mutation Detection
System (Biorad, München, Germany). Use
an 18–38% denaturing gradient.
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CRC Press, Boca Raton, FL, pp 355–386
Proteomic Protocols for the Study
of Filamentous Fungi
24
Raquel González Fernández
and Jesús V. Jorrín Novo
Abstract
In the last few years, proteomics has experienced rapid improvement in
technologies and applications. Gel-based strategies have become the
method of choice for both identification and quantification of proteins in
most studies. The workflow of a standard gel-based proteomic experiment
includes experimental design, sampling, protein extraction, protein separation, mass spectrometry analysis, protein identification, data statistical
analysis, validation of the identification, quantification, and data analysis.
The appropriate protocol to be used depends on and must be optimized for
the biological system (i.e., fungal species, plant species, organ, tissue,
cells). Preliminary steps are relevant. The choice of a good extraction protocol in a proteomic experiment is crucial because only if you can extract
and solubilize a protein you have a chance of detecting and identifying it.
This is more important in the case of filamentous fungi, which, owing to
their particular cellular characteristics, can be considered recalcitrant biological material, making it difficult to obtain quality protein samples to
proteomic analysis.
Fungi have an exceptionally robust cell wall, consisting largely of chitin, which makes up the majority of the cell mass. Because of its rigidity,
cell lysis is an important element in fungal proteomics. For protein extraction, various buffer- and precipitation-based protocols are available. In
most of these protocols, trichloroacetic acid (TCA) and/or acetone are
used for protein precipitation, or a phenol extraction is made, where proteins are solubilized in the phenolic phase and then are precipitated with
methanol and ammonium sulfate. These methods also eliminate some
R.G. Fernández (*) • J.V.J. Novo
Department of Biochemistry and Molecular Biology,
University of Córdoba, Agro-forestry and Plant
Biochemistry and Proteomics Research Group,
Ed. Severo Ochoa, planta baja. Campus de Rabanales,
Córdoba 14071, Spain
e-mail: q42gofer@uco.es
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_24, © Springer Science+Business Media, LLC 2013
299
300
R.G. Fernández and J.V.J. Novo
contaminants abundant in fungal material (such as polysaccharides, lipids,
nucleic acids, or phenolic compounds) that affect the protein isoelectrofocusing and electrophoresis processes.
Key words
fungal proteomics • fungal secretome • cell lysis • protein precipitation
• protein isoelectrofocusing
Introduction
In a post-genomic era, proteomic technologies
have become a powerful tool to study the proteome and to examine alterations in protein
profiles [1]. Similar to genomics and transcriptomics, proteomics has evolved to incorporate
high-throughput processes, which allow faster
analysis of a larger number of proteins [2, 3].
Proteomics involves the combined applications
of advanced separation gel-based, namely monoand two-dimensional electrophoresis (1-DE and
2-DE), and gel-free, such as liquid chromatography (LC) techniques, identification techniques
such as mass spectrometry (MS) analysis and
bioinformatics tools to characterize the proteins
in complex biological mixtures [4].
Plant pathogenic fungi cause significant yield
losses in crops. Molecular studies of the fungal
biological cycle and their interaction with their
hosts are necessary, in order to develop efficient
and environment-friendly crop protection strategies [5, 6]. Proteomics, in combination with other
techniques, constitutes a successful tool for providing important information about pathogenicity and virulence factors. Moreover, proteomics
also allows location-specific analysis (i.e., subproteomes at the level of organelles, cell membranes, cell wall, secretory proteins, etc.), the
study of post-translational modifications [7] and
interactions of host-pathogen, as well as hostpathogen-biocontrol agents [8, 9]. As a consequence, proteomics is opening up new possibilities
for crop disease diagnosis and crop protection.
Several areas can be defined in proteomics,
including descriptive and differential expression
proteomics. In the case of fungi, a new area can
also be defined as secretomics (the secretome is
defined as being the combination of native proteins and cell machinery involved in their secretion), since many fungi secrete an arsenal of
proteins to accommodate their saprotrophic lifestyle, namely proteins implicated in the adhesion
to the plant surface, host-tissue penetration, and
invasion effectors, together with other virulence
factors [10]. Fungal proteomics research has
experienced great advances over the last years,
because of the availability of powerful proteomics technologies and the increasing number of
fungal genome sequencing projects. Currently,
more than 50 pathogenic fungal genomes have
been sequenced.1 Excellent reviews on fungal
proteomic methodologies have been recently
published [4, 11]. The workflow of a fungal gelbased proteomics experiment includes, among
others, the following steps: experimental design,
fungal growth, sampling, sample preparation,
protein extraction, separation, MS analysis, protein identification, statistical analysis of data,
quantification, and data analysis, management,
and storage.
Most of plant pathogenic fungi are filamentous
fungi. This type of fungi can be considered, similarly to plants, recalcitrant biological material, so
the preparation of protein samples is a critical
step. Cell breakdown and protein extraction are
difficult because of the presence of a cell wall
that makes up the majority of the cell mass [12].
To overcome this challenge, early studies were
performed using mechanical lysis via glass beads
[13–15], a cell mill [16], or sonication [17–19],
because these methods are more efficient than
Broad Institute Database, http://broadinstitute.org/
science/project/fungal-genomeinitiative
1
24
Proteomic Protocols for the Study of Filamentous Fungi
those based on chemical or enzyme extraction
[20]. Shimizu and Wariishi [21] employed an
alternative approach to avoid the difficulty of
lysing the fungal cell wall by generating protoplasts of Tyromyces palustris. A better 2-DE pattern was obtained from protoplast than from
intact cells. The most widely used method for
cell disruption consists of pulverizing the mycelium in liquid nitrogen using a mortar and pestle
[17, 22–33]. The production of high-quality protein samples is also a crucial step for proteomic
analysis. The protocol most widely employed for
fungal proteins uses protein precipitation media
containing organic solvents, such as trichloroacetic acid (TCA), followed by solubilization of
the precipitate in an appropriate buffer. This
method
minimizes
protein
degradation/
modification. Furthermore, it removes interfering
compounds such as polysaccharides, polyphenols, pigment and lipids, which may be a problem during IEF [34], and prevents protease
activities [35]. TCA-treatment complicates subsequent protein solubilization for IEF, especially
with hydrophobic proteins. These problems have
been partly overcome by the use of chaotropes
(urea and thiourea) [36], new zwitterionic detergents [37–41], and by a brief treatment with
sodium hydroxide [35], which led to an increase
in resolution and capacity of 2-DE gels. Other
protein extraction methods have reported an
improvement when using acidic extraction solution to reduce streaking of fungal samples caused
by their cell wall [42], as well as using a phosphate buffer solubilization before the precipitation [23, 24]. Finally, the combined use of TCA
precipitation and phenol extraction provides a
better spot definition, due to the fact that it reduces
streaking and leads to a higher number of detected
spots [22, 43]. Alternative protocols for protein
extraction from spores of Aspergillus ssp. have
been optimized, since they use acidic conditions,
step organic gradient, and variable sonication
treatment (ultrasonic homogenizer and sonic
water bath) [19].
Special protocols are required for secreted proteins, due to the fact that there may be problems
such as a very low protein concentration, sometimes below the detection limit of colorimetric
301
methods (Bradford, Lowry, or BCA), or the
presence of polysaccharides, mucilaginous
material, salts, and secreted metabolites (lowmolecular organic acids, fatty acids, phenols, quinones, and other aromatic compounds). The
presence of these extracellular compounds may
impair standard methods for protein quantification
and may result in a strong overestimation of total
protein number [44]. This determination can also
be affected by the high concentration of reagents
from the solubilization buffer (such as urea,
thiourea, or DTT) that may interfere in the
spectrophotometric measurement, producing an
overestimation of the total amount of protein in
which, depending on the method, the differences
varied in the order of two magnitudes [45].
Comparison of different standard methods for
protein precipitation has demonstrated their limited applicability to analyzing the whole fungal
secretome [45–54].
Electrophoresis is almost the only protein separation technique employed in fungal research.
Despite its simplicity, 1-DE remains as quite a
valid technique providing relevant information,
especially in the case of comparative proteomics
with large numbers of samples to analyze. Thus,
it is possible by using this technique to distinguish between phenotypes of different wild-type
strains of Botrytis cinerea and to identify proteins
involved in the pathogenicity mechanisms
(Gonzalez-Fernandez et al., personal communication). With appropriate software, 1-DE is a
simple, reliable technique for finger-printing
crude extracts, and it is especially useful in the
case of hydrophobic and low-molecular-weight
proteins [53]. Therefore, the 1-DE is a good
approach to obtain preliminary results before carrying on 2-DE analysis [54–58].
Two-DE is the dominant platform in fungal
proteomics. Briefly, the 2-DE consists of a tandem pair of electrophoretic separations: in the
first dimension, proteins are resolved according
to their isoelectric points (pIs), normally using
IEF; while in the second dimension, proteins are
separated according to their approximate molecular weight using SDS-PAGE. Excellent reviews
describing and discussing the features and protocols of electrophoretic separations in proteomics
302
strategies have been published [34, 59]. The main
advantages of 2-DE are its high protein separation capacity and the possibility of making largescale protein-profiling experiments. Nevertheless,
reproducibility and resolution of this technique
are still remaining challenges. This method was
reported to under-represent proteins with extreme
physicochemical properties (size, isoelectric
point, transmembrane domains), as well as those
with a low abundance [60].
After separating proteins, they can then be
detected by a variety of staining techniques [34,
59] namely: (1) organic dyes, such as colloidal
Coomassie blue staining; (2) zinc–imidazole
staining; (3) silver staining; and (4) fluorescencebased detection, such as Sypro Ruby. The criteria to choose the staining method are the level of
sensibility and its compatibility with MS. Gels
are digitalized, and bands or spots are studied by
specific software of image analysis (i.e.,
Quantity-One, PD-Quest, BioRad). Bands or
spots are excised from gels and prepared for MS
analysis.
The limitations of gel-based analysis have led
to the more recent development of techniques
based on LC separation of proteins or peptides,
including two-dimensional liquid-phase chromatography 2-D LC-MS/MS (based on a high performance chromatofocusing in the first dimension
followed by high-resolution reversed-phase chromatography in the second) [61], and one-dimensional electrophoresis(1-DE)-nanoscale capillary
LC-MS/MS, namely GeLC-MS/MS (this technique combines a size-based protein separation
with an in-gel digestion of the resulting fractions)
[62]. This GeLC-MS/MS strategy paves the way
toward the analysis on a large-scale fungal
response environmental cues on the basis of
quantitative shotgun protein- profiling experiments. The case of Multidimensional Protein
Identification Technology (MudPIT), which
allows the identification of a much larger number
of proteins compared to gel-based methods, is
drawback being the lack of quantitative data [2,
63]. MudPIT was used to analyze the mechanisms of germling growth in Uromyces appendiculatus by comparing germinating asexual
uredospores with inactive spores [64].
R.G. Fernández and J.V.J. Novo
MS is the basic technique for global proteomic
analysis due to its accuracy, resolution, and sensitivity (in the femtomole to attomole concentration range), and due to the fact that is has the
capacity for a high throughput. Not only does it
allow one to profile a proteome, but more importantly, it allows one to identify the protein species
and
characterize
post-translational
modifications and interactions. Proteins are
identified from mass spectra of intact proteins
(top-down proteomics), or peptide fragments
obtained after enzymatic (mostly digested with
trypsin) or chemical treatment (bottom-up proteomics). Protein species are identified by comparison of the experimental spectra, while the
theoretical ones were obtained in silico from protein, genomic, ESTs sequence, or MS spectra
databases. For that purpose, different instrumentation, algorithms, databases, and repositories
are available [65, 66].
Although methods for proteomic analysis of
limited fungal species have been published [4,
11, 67–69], procedures for protein extraction and
2-DE gel analysis conditions are progressively
evolving according to individual characteristics
of fungal species.
Materials
(See Note 1)
1. Distilled water.
2. Liquid nitrogen.
3. Freeze-dryer.
4. Mortar and pestle.
5. Cell strainer, 100 mm Nylon (BD Falcon).
6. Vortexer.
7. Micropestles.
8. Ultrasonic homogenizer.
9. Microcentrifuge and centrifuge.
10. Disposable microcentrifuge tubes: 1.5 mL
and 2 mL.
11. Centrifuge tubes: 50 mL.
12. Trichloroacetic acid (TCA) (10% w/v)/acetone (80% v/v) solution.
13. 0.1 M Ammonium acetate/methanol (100%
and 80% v/v) solution.
14. Acetone (80% v/v) solution.
24
Proteomic Protocols for the Study of Filamentous Fungi
15. Phenol solution equilibrated with 10 mM
Tris–HCl pH 8 (Sigma-Aldrich).
16. SDS buffer: 30% (w/v) sucrose, 2% (w/v)
SDS, 5% (v/v) b-mercaptoethanol, 0.1 M
Tris–HCl pH 8.
17. Solubilization solution: 9 M urea, 2 M thiourea, 4% (w/v) CHAPS, 0.5% (v/v)
Tritón-X100, 20 mM DTT.
18. Microtube mixer.
19. Bradford solution (Sigma-Aldrich).
20. Extraction buffer: 8 M urea, 1% (w/v) SDS,
1 mM EDTA, 100 mM DTT, 50 mM Tris–HCl
pH 8.
21. TE buffer for secreted proteins: 50 mM EDTA,
2% (v/v) b-mercaptoethanol, 1 mM PMSF,
10 mL/mL buffer of protease inhibitor cocktail
for fungi (Sigma-Aldrich), Tris–HCl pH 8.
22. Running buffer: 192 mM Glycine, 1% (w/v)
SDS, 50 mM Tris–HCl pH 8.
23. Vertical electrophoresis equipment; for
example, Criterion System (BioRad).
24. Precast free stain gels (Criterion System,
BioRad): 4–20% Tris–HCl multi-wells for
1-DE and 8–16% Tris–HCl IPG + 1 for 2-DE.
25. IPG strips, 11 cm, pH 5–8 (BioRad).
26. IPG strips rehydration solution: 7 M urea,
2 M thiourea, 4% (v/v) CHAPS, 2% (v/v)
ampholytes (BioRad), 20 mM DTT.
27. Equilibration buffer: 6 M urea, 20% (v/v)
glycerol, 2% (w/v) SDS, 1.5 M Tris–HCl pH
8.8.
28. Shaker.
29. Densitometer; for example, GS-800
(BioRad).
Methods
The methods below have been optimized to
mycelium, secreted proteins in liquid media, and
conidia from B. cinerea, although these procedures can be applied in proteomic analysis of
filamentous fungi in general.
Sample Collection
For in vitro cultures, conidia are produced using
rich-media plates at 22 °C under constant black
303
light (UV) for 3–4 weeks. Mycelium and secreted
proteins can be obtained from liquid cultures
inoculated with conidia or nonsporulating mycelia (see Note 2). Mycelia and media can be separated by centrifugation and filtration, frozen in
liquid nitrogen, and lyophilized.
Protein Extraction by TCA/
Acetone-Phenol/Methanol Method
Protein extraction is carried on by using the TCA/
acetone–phenol/methanol method [70, 71] with
some modifications [4] and adapted to started
material (conidia, mycelium or secreted
proteins).
Mycelium
The lyophilized mycelium is ground to a fine
powder in liquid nitrogen using a cooled mortar
and pestle, and stored at −80 °C for later analysis
(see Note 3). For protein extraction, the following protocol is applied:
1. Transfer 50–100 mg of mycelial powder into
a 2-mL tube.
2. Add 1 mL of 10% (w/v) TCA/acetone and
mix well using a micropestle and then by
vortexing.
3. Sonicate 3 × 10 s (50 W, amplitude 60) at
4 °C, breaking on ice at 1 min.
4. Fill the tube with 10% (w/v) TCA/acetone.
Mix well by vortexing.
5. Centrifuge at 16,000 × g for 5 min (4 °C) and
remove the supernatant by decanting (see
Note 4).
6. Fill the tube with 0.1 M ammonium acetate
in 80% (v/v) methanol. Mix well by
vortexing.
7. Centrifuge at 16,000 × g for 5 min (4 °C) and
discard the supernatant.
8. Fill the tube with 80% (v/v) acetone. Mix
well by vortexing.
9. Centrifuge at 16,000 × g for 5 min (4 °C) and
discard the supernatant.
10. Air-dry at room temperature to remove residual acetone.
11. Add 1.2 mL of 1:1 phenol (pH 8, SIGMA)/
SDS buffer. Mix well using a pipette and by
vortexing. Incubate for 5 min in ice.
304
12. Centrifuge at 16,000 × g for 5 min. Transfer
the upper phenol fase into a new 1.5-mL tube
(see Note 5).
13. Fill the tube with 0.1 M ammonium acetate
in 100% (v/v) methanol, mix well, and allow
the precipitation overnight at −20 °C.
14. Centrifuge at 16,000 × g for 5 min (4 °C) and
discard the supernatant (a white pellet should
be visible).
15. Wash the pellet with 100% methanol and
mix by vortexing.
16. Centrifuge at 16,000 × g for 5 min (4 °C) and
discard the supernatant.
17. Wash the pellet with 80% (v/v) acetone and
mix by vortexing.
18. Centrifuge at 16,000 × g for 5 min (4 °C) and
discard the supernatant.
19. Dry the pellet at room temperature.
20. Dissolve the proteins in solubilization solution for 2 h, shaking in a microtube mixer at
4 °C (see Note 6).
21. Quantify proteins using Bradford method
[72].
22. Store the protein extracts at −20 °C for
further analysis.
Secreted Proteins
Lyophilized media are re-suspended in 5 mL of
TE buffer and proteins are precipitated according
to the following protocol:
1. Transfer the medium resolubilized into a
50-mL tube and add 2/1 (v/v) (10 mL) of
20% (w/v) TCA/acetone. Mix well by vortexing and allow protein precipitation overnight at 4 °C.
2. Centrifuge at 16,000 × g for 10 min (4 °C)
and remove the supernatant by decanting
(see Note 7).
3. Add a volume 4/1 (v/v) (20 mL) of 0.1 M
ammonium acetate in 80% (v/v) methanol.
Mix well by vortexing.
4. Centrifuge at 16,000 × g for 10 min (4 °C)
and discard the supernatant.
5. Add a volume 4/1 (v/v) (20 mL) of 80% (v/v)
acetone. Mix well by vortexing.
6. Centrifuge at 16,000 × g for 10 min (4 °C)
and discard the supernatant.
R.G. Fernández and J.V.J. Novo
7. Air-dry at room temperature to remove
residual acetone.
8. Add 4 mL of 1/1 (v/v) phenol (pH 8, SIGMA)/
SDS buffer. Mix well by vortexing and transfer the 4-mL to two 2-mL eppendorf. Incubate
for 5 min in ice.
9. Centrifuge at 16,000 × g for 10 min. Transfer
the upper phenol phase into a new 2-mL tube
(1 mL per 2-mL tube).
10. Fill the tube with 0.1 M ammonium acetate
in 100% (v/v) methanol, mix well, and allow
to precipitate overnight al −20 °C.
11. Centrifuge one 2-mL tube at 16,000 × g
for 5 min (4 °C) and discard the supernatant
(a slight pellet should be visible). Fill the
same 2-mL tube with the other eppendorf
(mix well before changing). Centrifuge at
16,000 × g for 5 min (4 °C) and discard the
supernatant.
12. Follow the steps in Mycelium section, starting with step 15.
Conidia
Conidia can be harvested with H2Od with 0,01%
Tween-80 scraping on the surface of agar plate.
The conidia suspension is filtered through a cell
strainer, concentred in 1.5-mL tubes, centrifuged at 16,000 × g for 5 min (4 °C), lyophilized
and stored at −80 °C for further analysis. For
protein extraction, the TCA/acetone-phenol/
methanol [70, 71] method was used, with some
modifications [4, 19].
1. Add 300 mL of extraction buffer to conidia. Mix
well using a micropestle and by vortexing.
2. Sonicate 3 × 10 s (50 W, amplitude 60), breaking on ice at 1 min. Mix well using a micropestle and by vortexing.
3. Centrifuge at 16,000 × g for 5 min (4 °C).
4. Fill the tube with 10% (w/v) TCA/acetone.
Mix well using by vortexing.
5. Centrifuge at 16,000 × g for 5 min (4 °C) and
discard the supernatant.
6. Fill the tube with 0.1 M ammonium acetate in
80% (v/v) methanol. Mix well using a
micropestle and by vortexing.
7. Follow the steps in Mycelium, starting with
step 5.
24
Proteomic Protocols for the Study of Filamentous Fungi
Protein Separation
One-Dimensional Electrophoresis
Proteins can be separated by SDS-PAGE according to Laemmly electrophoresis system, [73] for
example, using Criterion System (BioRad) with
precast Criterion Stain Free Gels, Tris–HCl,
4–20% linear gradient (BioRad). The 1-DE is
visualized using the Image Lab System (BioRad),
and stained by CBB (Coomassie Blue Brilliant)
method [74] (see Note 8). After the staining of
proteins, bands can be analyzed using the
Quantity-One software (BioRad).
Two-Dimensional Electrophoresis
Isoelectrofocusing
Focusing conditions will vary with sample composition, sample complexity, and strip pH range. In
our conditions, the 11 cm IPG strips, pH 5–8
(BioRad), are rehydrated with 50 mg of protein
extract in 185 mL rehydration solution applying
50 V for 16 h (active rehydratation) at 20 °C. Before
the focusing a wet wick is inserted under each end
of the strip (catode). The conditions for IEF have
been adapted to our system from reference [45]:
150 V for 1 h, 1 h at 200 V, 1 h at 500 V, 1,000 V·h
at 1,000 V, followed by 2.5 h gradient from 1,000 to
8,000 V, and finally focused for 30,000 V·h at
8,000 V, with a cell temperature of 20 °C (see
Note 9). After IEF, IPG strips are stored at −20 °C.
Before the second dimension, IPG strips are
equilibrated in two steps. Firstly, it is carried on
with 2% (w/v) DTT in equilibration buffer for
10 min in agitation at room temperature; secondly, it is done with 2.5% (w/v) iodoacetamide
in equilibration buffer for 10 min in agitation at
room temperature.
The second dimension is performed in the
same way as SDS-PAGE, but using precast
Criterion Stain Free Gels, Tris–HCl, 8–16% linear gradient for IPG strips (BioRad). After the
staining of proteins, spots can be analyzed using
the PD-Quest software (BioRad).
Protein Identification
The bands or spots are cut out and digested with
trypsin. Tryptic peptides are analyzed in a mass
305
spectrometer; for example, a 4,800 Proteomics
Analyzer
MALDI–TOF/TOF
(Applied
Biosystems). In this case, the most abundant peptide ions are subjected to MS/MS analysis.
A PMF search and a combined search (+MS/MS)
are performed in nrNCBI database of proteins
using the MASCOT algorithm (see Note 10).
Notes
1. Gloves and lab coat should be used in these
procedures, and particular care should be taken
when handling TCA and phenol (consult safety
data sheets) because they are corrosive products. Steps involving phenol and b-mercaptoethanol should be performed in a fume hood.
2. Examples of rich-media are PDAB (potato,
dextrose, agar + bean leaves), solid synthetic
complete medium (CM) [75], or solid malt
extract medium (1.5% w/v).
3. Be careful to work with liquid nitrogen
because its cool temperature (−195.8 °C)
could cause severe frostbite. The nitrogen
evaporated reduces the concentration of oxygen in the air and can act as an asphyxiant,
especially in confined spaces, so it may be
dangerous because nitrogen is odorless, colorless, and tasteless, and could cause suffocation without any sensation or warning.
4. Be careful to not throw out the pellet.
5. Three phases appear, namely: the upper
phase (which is the phenolic phase where are
the proteins), a white interphase, and a lower
aqueous phase. Try to not to get part of the
white interphase.
6. The volume of solubilization solution added
will depend on quantity of precipitated proteins. It is advisable that samples are well
concentrated.
7. In this case, maybe that the precipitated pellet is faint because the proteins secreted to
medium are at very low concentration.
8. More details about 1-DE and 2-DE separation methods are described in two excellent
reviews [34, 59].
9. The condition of protein focusing must be
optimized for each system of study. In our
case, we use the PROTEAN IEF cell by
306
BioRad. The conditionating phase involves
the application of previous steps at low voltage that allow to remove ions and other contaminants containing the sample, and that
interfere on protein focusing. The current
should not exceed 50 mA per strips. For more
information see the 2-D Electrophoresis for
Proteomics Manual by BioRad.
10. More details about MS analysis are described
in references [65, 68, 76].
Acknowledgments This work was supported by the
Spanish Ministry of Science and Innovation (BotBank
Project, EUI2008-03686), the Regional Government of
Andalusia (Junta de Andalucía) and the University of
Córdoba (AGR-0164: Agricultural and Plant Biochemistry
and Proteomics Research Group).
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Detection and Quantification
of Endoprotease Activity Using
a Coomassie Dye-Binding Assay
25
Anthony J. O’Donoghue and Cathal S. Mahon
Abstract
Traditional methods for detecting proteases in fungi require the separation
of product from substrate. These methods are time-consuming, laborious,
and not amenable to high-throughput analysis. A simple alternative method
is described here that utilizes Coomassie dye reagent to follow the timedependent proteolytic loss of a macromolecular protein substrate.
Keywords
Protease assay • Peptidase • Fungi • Colorimetric • Coomassie dye
Introduction
Fungi possess multiple proteases with highly
diverse functions that are central to their physiology, metabolism, and development. These functions range from simple digestion of protein for
food to complex regulation of signal transduction
pathways. There are six classes of protease found
in fungi that are defined based on the nature of
the functional group in the active site and
are therefore termed aspartic, glutamic, serine,
threonine, cysteine, and metallo proteases. Some
enzymes in these families exhibit exopeptidase
A.J. O’Donoghue (*) • C.S. Mahon
Department of Pharmaceutical Chemistry,
University of California—San Francisco,
600 16th Street, San Francisco, CA 94158, USA
e-mail: aodonoghue@picasso.ucsf.edu
activity, catalyzing the release of mono, di-, or
tripeptides from the amino or carboxy termini of
a polypeptide. However, most of the proteolytic
enzymes isolated and characterized from fungi
possess endoprotease activity and cleave polypeptides distal from the termini [1].
There are several methods to detect and quantify endoproteolytic activity in fungus. One technique utilizes synthetic peptide substrates with
colorimetric or fluorescent labels that produce a
signal upon protease cleavage. While these substrates are highly specific and sensitive to proteolysis, knowledge of the protease specificity is a
prerequisite in choosing a substrate. Where substrate specificity is unknown, there are several
general endoprotease assays for detection of proteolytic activity in fungal extracts and spent
media. Traditional techniques employ common
macromolecular substrates such as casein, albumin, and hemoglobin. Proteolytic cleavage can
be detected by SDS PAGE followed by Coomassie
staining or trichloroacetic acid precipitation
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_25, © Springer Science+Business Media, LLC 2013
309
310
followed by spectrophotometry [2]. These techniques are time-consuming, laborious, and not
compatible with microplate assay formats.
Fluorescently labeled proteins such as casein,
elastin, and gelatin are useful for high-throughput
protease assays [3]. These macromolecular substrates are heavily conjugated with dye such that
fluorescence is efficiently quenched. Proteolytic
release of dye-labeled peptides results in an
increase in fluorescence that correlates with
enzyme activity.
A simple, rapid, and quantitative colorimetric
assay is described here that is a modification of
assays developed by Saleemuddin and co-workers
[4] and Buroker-Kilgore and Wang [5]. The protocol utilizes Coomassie Brilliant Blue G-250 dye to
follow the decrease of protein substrate following
endoprotease digestion and functions on the basis
that the dye does not bind to peptides less than
~3,000 Da [6]. This assay can be used to investigate endoprotease activity using any number of
proteins as substrate and digestion produces a
quantitative and time-dependent loss of substrate.
Variations of this technique have been used successfully to detect and characterize endoproteases
from bacteria [7], protozoa [8], and multiple fungi
[9–12]. We used the dye-binding endoprotease
assay described here to detect and subsequently
purify two acid-acting proteases from the fungus
Talaromyces emersonii [13]. In our experimental
strategy we utilized bovine serum albumin (BSA)
as the protein substrate but other groups have successfully used casein [11], gelatin [7], IgG, hemoglobin, and mucin [8]. Furthermore, proteolytic
activity in the presence of inhibitors can be readily
performed [13]. This assay is pH- and temperature-independent, amenable to microplate assay
format and requires only dye, a source of protein
substrate and a spectrophotometer.
A.J. O’Donoghue and C.S. Mahon
5. Phosphate buffered saline (PBS).
6. Microcentrifuge with 1.5-mL microcentrifuge tubes.
7. Coomassie Brilliant Blue G-250 in phosphoric acid and methanol. Available from
Thermo Scientific (Coomassie Plus Bradford
Assay Reagent), Sigma-Aldrich (Coomassie
Protein Assay Reagent) or BioRad (Quick
Start Bradford 1X Dye Reagent).
8. Protease sample; cell-free culture supernatant, or cell lysate diluted to <0.1 mg/mL in
assay buffer.
9. Protease assay buffer; choice of buffer will
depend on pH preference of protease.
Suggested buffers include 25 mM citrate–
phosphate buffer pH 2.5–6.0, 25 mM
Phosphate buffer pH 6.0–8.0, 25 mM Tris–
HCl buffer pH 7.5–9.0 or 20 mM Glycine–
NaOH pH 9.0–10.5. All buffers can be
supplemented with salts, reducing reagents,
and detergents at concentrations compatible
with Coomassie dye-binding reagent (see
supplier’s compatibility chart).
10. 2 mg/mL of BSA or any other protein that
produces a linear response curve to Coomassie
dye binding.
11. Microplate spectrophotometer with 580- to
610-nm filter (595 nm is optimal).
12. 96-well microtiter assay plates.
13. PCR tubes.
14. Thermal cycler.
15. Protease inhibitors (1,10-Phenanthroline,
Pepstatin A, PMSF, and E-64; all available
from Sigma-Aldrich).
16. Aspergillopepsin
(Sigma-Aldrich,
Catalogue# P2143).
17. Trypsin (Sigma-Aldrich, Catalogue #
T8003).
Method
Materials
1. 50 mL of sterile culture media in 250-mL
shake flask.
2. Cheese cloth.
3. Liquid nitrogen.
4. Mortar and pestle.
Fungal Culture and Enzyme Sample
Preparation
1. Inoculate a flask of sterile liquid media with
fungal cells and grow for 2–5 days under optimal conditions.
25
Detection and Quantification of Endoprotease Activity…
2. Separate cells from media using several layers
of cheese cloth (or centrifugation).
3. Transfer 1.5 mL of filtered media to a microcentrifuge tube and spin at 12,000 × g for
20 min at 4 °C. Remove supernatant and store
at 4 °C. This is the cell-free media sample.
4. “Squeeze dry” filtered cells and freeze with
liquid nitrogen. Grind to a fine powder using
mortar and pestle.
5. Transfer powder to a microcentrifuge tube and
dissolve in PBS. Spin at 12,000 × g for 20 min
at 4 °C.
6. Remove supernatant and determine protein
concentration. Dilute to <0.1 mg/mL in PBS.
3.
4.
5.
6.
Generation of Standard Curve
for Protein Substrate
1. Remove Coomassie dye reagent from refrigerator and equilibrate to room temperature.
2. Make a 2-mg/mL stock solution of BSA (or
other protein substrate) in assay buffer.
3. Make a dilution series of substrate in assay
buffer starting at 1.5 mg/mL.
4. Pipette 180 mL of each dilution in triplicate
into wells of a microplate, known as the protease assay (PA) plate.
5. Add 20 mL of control protease sample (fresh
media or PBS) to each well, mix by pipetting,
and remove 20 mL into wells of a second plate,
known as the dye-binding (DB) plate.
6. Add 180 mL of Coomassie dye reagent, shake
for 10 s, and incubate at room temperature for
5 min.
7. Measure the absorbance between 580 and
610 nm (595 nm is optimal) and generate a
standard curve. (see Note 1)
Endoprotease Assay
1. Choose a substrate concentration that produces the highest absorbance on the linear
portion of the standard curve (Fig. 25.1a).
Make a 20-mL stock in assay buffer.
2. Add 180 mL of this protein stock into a
PA well or PCR tube. A PCR tube and a
7.
311
thermocycler are important for assays
performed at elevated temperature (>30 °C).
Combine 20 mL of cell-free media or cell
lysate with substrate. Mix by pipetting and
immediately remove 20 mL into triplicate
wells on the DB plate.
Add 180 mL of Coomassie dye reagent and
measure absorbance as outlined previously.
This sample is the zero minute time point (T0).
(see Note 2)
Remove 20 mL in triplicate from the reaction
after 5, 10, 15, and 20 mins and repeat treatment with Coomassie dye reagent in DB
plate.
Use the standard curve to calculate the concentration of substrate remaining at each time
point and plot versus time (see example in
Fig. 25.1b). Note: If no significant loss of substrate is observed after 20 min, the assay can
be run for several hours to days. Use a PCR
tube to minimize evaporation.
Optional: Set up a control reaction with either
0.2 mg of Aspergillopepsin I at pH 1.5–4.5 or
Trypsin at pH 7–9 to monitor time-dependent
cleavage of protease substrate.
Inhibition Assay
1. Combine an equal volume of fungal sample
containing protease with 20× Protease
Inhibitor. Mix briefly and incubate for 15 min
at room temperature. Set up a control reaction
containing no inhibitor.
2. Pipette 20 mL of this reaction into a well in the
PA plate or PCR tube containing 180 mL of
substrate, incubate, and repeat dye-binding
assay as outlined above. (see Note 3).
Calculation of Protease
Cleavage
Loss of substrate at each time point can be calculated using the standard curve and the following
formula:
∆ (delta )S = [S0 ]− [SX ]
312
A.J. O’Donoghue and C.S. Mahon
Fig. 25.1 (a) Standard curve of BSA absorbance at 595 nm
following Coomassie dye binding. The highest substrate
concentration in the linear portion of the graph was chosen
as the substrate concentration for the protease assay. (b)
Time-dependent cleavage of BSA measured by Coomassie
dye-binding assay. Spent media from shake-flask cultures
of Talaromyces emersonii were assayed for proteolytic
activity as described in Methods section. Assays were carried out in citrate-phosphate pH 3.3, ammonium acetate
pH 5.0, and sodium phosphate pH 6.75 and 7.5.
where S0 and SX are substrate concentrations at T0
and TX minutes, respectively. It is important to
always take a T0 sample because the addition of
enzyme to substrate may result in an overall
increase in absorbance, particularly when using a
cell lysate as the source of protease.
Notes
1. To increase sensitivity of the assay, a higher
ratio of protein substrate to Coomassie dye
reagent may be used. Some protocols use an
equal volume of protein sample to dye.
25
Detection and Quantification of Endoprotease Activity…
2. For most assays, addition of Coomassie dye
will instantly quench reaction due to the low
pH of the reagent. However, for acid-acting
enzymes quenching can be performed by heat
denaturation in a thermocycler or by the addition of inhibitor to dye reagent.
3. The total enzyme concentration is 50 % relative to the previous assay so extended incubation may be required to observe cleavage.
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Protocol of a LightCycler™
PCR Assay for Detection
and Quantification of Aspergillus
fumigatus DNA in Clinical Samples
of Neutropenic Patients
26
Birgit Spiess and Dieter Buchheidt
Abstract
The increasing incidence of life-threatening systemic fungal infections
emphasizes the need to improve molecular diagnostic tools. Polymerase
chain reaction method (PCR) was used to establish sensitive and rapid
molecular detection assays of pathogens not detectable in cultures. Using
the LightCycler™ technology that combines amplification of DNA with
an immediate fluorescence detection of the amplicon, a real-time PCR
assay was established to achieve an improved, specific, sensitive, and rapid
method for quantification of the Aspergillus fumigatus fungal load in clinical samples of hematological patients in order to improve antifungal treatment monitoring.
Keywords
Aspergillus fumigatus • Polymerase chain reaction (PCR) • Clinical
samples • Fungal load • Neutropenic patients
Introduction
The increasing incidence of life-threatening
systemic fungal infections, especially invasive
aspergillosis (IA), correlates with an increased
number of immunocompromised patients [1].
Patients at highest risk are those who received
B. Spiess (*) • D. Buchheidt
3rd Department of Internal Medicine,
Hematology and Oncology, Scientific Laboratory,
Mannheim University Hospital, Pettenkoferstr. 22,
Mannheim 68169, Germany
e-mail: birgit.spiess@medma.uni-heidelberg.de
intensive cytotoxic chemotherapy for acute leukemia or bone marrow or allogeneic hematopoietic stem cell transplantation, which can lead to
prolonged periods of neutropenia [2–7].
Due to the limited prognosis of patients with
invasive Aspergillus infections, all diagnostic
approaches primarily aim at an early confirmation of an infection, to optimize antifungal
treatment [8].
It remains difficult to diagnose IA at all, since
the current diagnostic tools either lack specificity
or sensitivity or both, at worst. At present, only
positive results from conventional cultures or histological examination provide the definitive proof
of invasive aspergillosis. However, establishing
cultures from blood, bronchoalveolar lavage
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_26, © Springer Science+Business Media, LLC 2013
315
316
(BAL), or other clinical samples is often unsuccessful because of the low yields of colony-forming units of the pathogen [9].
Polymerase chain reaction method (PCR) was
used to establish sensitive and rapid molecular
detection assays of pathogens not detectable in
cultures.
Against this background we established and
evaluated a highly sensitive and Aspergillus
specific two-step PCR assay for the analysis of
clinical samples (blood, BAL, CSF, tissue samples) [3–7,10,11].
In order to estimate the fungal burden and to
monitor and evaluate the response to antifungal
drugs, the quantification of the fungal burden is
of great clinical relevance.
Using the LightCycler™ technology that
combines amplification of DNA with an immediate fluorescence detection of the amplicon, we
established an assay for clinical samples and
achieved an improved, specific, sensitive, and rapid
method for quantification of the A. fumigatus fungal
load [12].
Materials
1. Sterile distilled water.
2. Puffer 10× RCLB: 1.55 M NH4Cl, 0.1 M
NH4HCO3, 1 mM EDTA pH 7.4.
3. Puffer 1× PBS.
4. Lyticase.
5. Proteinase K.
6. 10% SDS.
7. 2× APEX: 400 mM Tris/Cl, 20 mM EDTA,
2% SDS.
8. Phenol:chloroform (1:1).
9. 70% isopropanol.
10. 70% ethanol.
11. LightCycler Fast Start DNA Master
Hybridization Probes kit (Roche Applied
Science, Mannheim, Germany).
12. Oligonucleotide primers and labeled probes
(custom-made TIB MOLBIOL, Berlin,
Germany).
13. Microcentrifuge
(Heraeus,
Biofuge,
Frankfurt, Germany).
14. LightCycler (Roche Applied Science,
Mannheim, Germany).
B. Spiess and D. Buchheidt
15. LightCycler glass capillaries.
16. Agarose, molecular biology grade (BioRad,
Munich, Germany).
17. Puffer 10× TBE.
18. Ethidium bromide.
19. Horizontal electrophoresis equipment (e.g.,
BioRad, Munich, Germany).
20. U.V. transilluminator and camera suitable for
imaging agarose gels (e.g., ChemiDoc XRS+,
BioRad, Munich, Germany).
Methods
Clinical Samples from
Immunocompromised Patients
Blood samples were obtained under sterile conditions by venipuncture, in a sterile vessel containing potassium EDTA to a final concentration of
1.6 mg EDTA per milliliter of blood. The sample
volume was 1–5 mL.
Bronchoalveolar lavage samples: bronchoscopy was performed by experienced physicians
according to guidelines [9] and BAL samples were
obtained in a sterile vessel without conservation
media. The mean sample volume was 8–10 mL.
CSF and tissue samples were obtained in a
sterile vessel without conservation media by
puncture under sterile conditions, according to
specific clinical guidelines.
All clinical samples were drawn after informed
consent of the individual patient.
DNA Preparation
1. 3–5 mL peripheral blood were mixed with 5
volumes of RCLB (10× RCLB; red cell lysis
buffer: 1.55 M NH4Cl, 0.1 M NH4HCO3,
1 mM EDTA pH 7.4) and incubated on ice for
10 min for lysis of the erythrocytes.
2. After incubation, the sample was centrifuged
for 10 min at 300×g. Supernatant was discarded, the leukocytes were washed once
with 1× PBS (10× PBS; phosphate buffered
saline: 1.4 M NaCl, 50 mM KCl, 90 mM
Na2PO4⋅2H2O, 20 mM KH2PO4, pH 7.4) and
recentrifuged.
26
Protocol of a LightCycler™ PCR Assay for Detection and Quantification…
3. BAL samples were transferred into 1.5 mL
tubes and centrifuged for 5 min at 300×g. The
sedimented leukocyte pellet was resuspended
in 300 ml 1× PBS and incubated with 10–125 U
of lyticase (50,000 U, Sigma-Aldrich,
Deisenhofen, Germany) for 30 min at 37 °C to
achieve degradation of the fungal cells.
4. 500–1000 mg proteinase K (Roche Molecular
Biochemicals, Mannheim, Germany) and
0.5% SDS (Sigma-Aldrich) were added
and the suspension was incubated at 55 °C
for 1 h.
5. By treatment with additional 100 ml 2× APEX
(2× APEX; Aspergillus extraction buffer:
400 mM Tris/Cl, 1 M NaCl, 20 mM EDTA,
2% SDS) for 30 min at 65 °C, residual cell
material was lysed. DNA isolation was performed under a laminar flow.
6. The purification of the fungal and human DNA
mixture was performed by conventional phenol–chloroform extraction [13].
7. The DNA was precipated with 70% (v/v) of
isopropanol and the DNA pellet was washed
once with 70% ethanol and air dried.
8. DNA concentration of human DNA was measured by spectrophotometry at 260 nm/
280 nm.
317
Alignment of the DNA sequences was performed using the program Geneworks
(Intelligenetics, Inc.) with standard algorithms.
After testing of 16 theoretically wise combinations of primers and hybridization probes, the
optimum pairs were chosen for all subsequent
PCR assays.
Out of the mitochondrial Cytochrome B gene
sequence of Aspergillus fumigatus (GenBank
accession no. AB025434) the sequence for the
forward primer was 5¢-AATGCACGATACTGTA
GGATCTG-3¢ (AfLC2s), and for the reverse
primer 5¢-TGCATTGGATTAGCCATAACA-3¢
(AfLC2as). The length of the amplified fragment
was 194 bp. Hybridization probes were selected
from the region between forward and reverse
primers of the primer pair.
For labeling of one probe at the 5¢ end, the
LightCycler Red 640 fluorophore was used:
5¢-TAATCTATCATAATTACCAGAAATACCT
AAAGGA-3¢ (Cyt3A). The other probe was
labeled at the 3¢ end with fluorescein:
5 ¢ -AATCTTTAAATACAAAGTAAGGAGCG
AAAG-3¢ (Cyt3B) [12]. Primers and hybridization probes were obtained from TIB MOLBIOL,
Berlin, Germany.
Quantification of A. fumigatus DNA
Primers and Hybridization Probes
for the LightCycler™-based PCR Assay
PCR primers and probes were derived from fungal mitochondrial Cytochrome B genes.
Several regions of the mitochondrial Cytochrome B gene of minor homology between four
Aspergillus, four Candida species, and human
DNA were the presupposition for the design of
the primers and hybridization probes (A. fumigatus (GenBank accession no. AB025434), A. flavus
(GenBank accession no. AB000596), A. terreus
(GenBank accession no. AB000603) and A. niger
(GenBank accession no. AB000597), C. albicans
(GenBank accession no. AB0044919), C. parapsilosis (GenBank accession no. AB044929),
C. glabrata (GenBank accession no. AB044922),
C. tropicalis (GenBank accession no. AB044930),
and human (GenBank accession no. M28016)
(Fig. 26.1).
Amplification and quantification of A. fumigatus
DNA was performed using the LightCycler™
PCR and detection system (Roche Applied
Science, Mannheim, Germany). The Hot Start
PCR reaction was performed in glass capillaries
using the LightCycler Fast Start DNA Master
Hybridization Probes kit (Roche Applied Science)
as described by the manufacturer. PCR mixture
contained 1× Fast Start reaction mix including
the Fast Start Taq DNA polymerase, reaction buffer, dNTPs and 10 mM MgCl2, all together
3.5 mM MgCl2, 20 pmol of each primer, and
60 nmol of hybridization probes.
In a volume of 20 ml PCR was performed under
following conditions: initial denaturing for 8 min
at 95 °C, 45 cycles with 4 s at 95 °C, annealing for
8 s at 58 °C, and enzymatic chain extension for
20 s at 72 °C. Each analysis done by PCR included
a H2O negative control without any template
318
B. Spiess and D. Buchheidt
Fig. 26.1 Multiple nucleotide sequence alignment of
mitochondrial Cytochrome B genes of A. fumigatus
(GenBank accession no. AB025434), A. flavus (GenBank
accession no. AB000596), A. terreus (GenBank accession
no. AB000603), C. albicans (GenBank accession no.
AB0044919), C. glabrata (GenBank accession no.
AB044922), C. tropicalis (GenBank accession no.
AB044930), and human (GenBank accession no.
M28016). Locations of primers and hybridization probes
are underlined. Homologous regions are in boxes
DNA to monitor for possible contamination.
Aliquots of DNA from healthy control persons
were prepared in parallel to clinical sample DNA
specimens and analyzed as negative controls.
A serially diluted standard of genomic
A. fumigatus DNA was used. 1 × 106 copies of the
mitochondrial Cytochrome B gene were corresponding to 1.32 ng and 1 × 10 copies to 13.2 fg
of genomic A. fumigatus DNA or 1–5 CFU per
mL blood.
The logarithmic linear phase was distinguished
from the background by online monitoring
26
Protocol of a LightCycler™ PCR Assay for Detection and Quantification…
319
Fig. 26.2 Determination and quantification of serially
diluted standard of Aspergillus fumigatus DNA (a) using
the LightCycler PCR technique. (b) LightCycler standard
curve report for serially diluted Aspergillus fumigatus
DNA. A representative evaluation is shown
(Figs. 26.2a, b). The amounts of Aspergillus DNA
in unknown samples were calculated by comparing the Cytochrome B gene copy numbers of the
logarithmic linear phase of the sample with the
copy numbers of the standards (Fig. 26.3) [12].
10 healthy control persons in the LightCycler
PCR assay. Only DNA from A. fumigatus (DSM
819 CS) was detectable in the LightCycler PCR
assay. All PCR assays with other fungal and bacterial strains were negative.
PCR has been shown to be a highly sensitive
and specific diagnostic tool for the detection of
Aspergillus species in clinical samples. We aimed
to extend the diagnostic value of our nested PCR
assay [10] to a quantification of the pathogen
load in order to improve the antifungal treatment
monitoring.
For clinical evaluation we investigated clinical samples from neutropenic patients suffering
Specificity of the LightCycler PCR Assay
DNA from several fungal and bacterial strains
was subjected to the LightCycler PCR to determine the specificity of the assay. Cross reactivity
of the primers and hybridization probes with
human DNA was excluded by testing of DNA of
320
Fig. 26.3 Correlation between LightCycler quantification
of defined amounts of Aspergillus fumigatus DNA and
sample amounts (1–5 ng) in vitro. Data represent
means ± SD from five separate experiments
from malignant hematological diseases. BAL and
blood samples gave positive results in the
LightCycler PCR assay and were also tested with
our previously described [10] nested PCR assay.
The PCR-mediated quantification of the fungal
burden showed 15–269,018 CFU per mL of BAL
and 298–104,114 CFU per mL of blood sample.
BAL and blood samples from subjects without
evidence for invasive pulmonary aspergillosis
were PCR-negative [12].
In studies, up to now, we screen clinical samples of high-risk patients first with our nested
PCR assay providing highest sensitivity and general specificity for Aspergillus species. Samples
tested positive in the nested PCR assay are subsequently quantified with the LightCycler PCR
assay [4–6].
Notes
In summary, our highly specific and sensitive
LightCycler-based real-time PCR assay is applicable for the rapid and early detection of Aspergillus
species and the quantification of the fungal load
from clinical samples of high-risk patients.
References
1. Buchheidt D (2008) Molecular diagnosis of invasive
aspergillosis in patients with hematologic malignancies—new answers to a diagnostic challenge? Expert
Opin Med Diagn 2:753–761
B. Spiess and D. Buchheidt
2. Lehrnbecher T, Frank C, Engels K, Kriener S, Groll
AH, Schwabe D (2010) Trends in the postmortem epidemiology of invasive fungal infections at a university hospital. J Infect 61:259–265
3. Buchheidt D, Baust C, Skladny H, Baldus M,
Brauninger S, Hehlmann R (2002) Clinical evaluation
of a polymerase chain reaction assay to detect
Aspergillus species in bronchoalveolar lavage samples
of neutropenic patients. Br J Haematol 116:803–811
4. Buchheidt D, Hummel M, Schleiermacher D, Spiess B,
Schwerdtfeger R, Cornely OA et al (2004) Prospective
clinical evaluation of a LightCycler-mediated polymerase chain reaction assay, a nested-PCR assay and a
galactomannan enzyme-linked immunosorbent assay
for detection of invasive aspergillosis in neutropenic
cancer patients and haematological stem cell transplant
recipients. Br J Haematol 125:196–202
5. Buchheidt D, Baust C, Skladny H, Ritter J, Suedhoff
T, Baldus M et al (2001) Detection of Aspergillus species in blood and bronchoalveolar lavage samples
from immunocompromised patients by means of
2-step polymerase chain reaction: clinical results.
Clin Infect Dis 33:428–435
6. Hummel M, Spiess B, Kentouche K, Niggemann S,
Bohm C, Reuter S et al (2006) Detection of Aspergillus
DNA in cerebrospinal fluid from patients with cerebral aspergillosis by a nested PCR assay. J Clin
Microbiol 44:3989–3993
7. Hummel M, Spiess B, Roder J, von Komorowski G,
Durken M, Kentouche K et al (2009) Detection of
Aspergillus DNA by a nested PCR assay is able to
improve the diagnosis of invasive aspergillosis in paediatric patients. J Med Microbiol 58:1291–1297
8. Verweij PE, Maertens J (2009) The changing face
of febrile neutropenia-from monotherapy to moulds
to mucositis. Moulds: diagnosis and treatment.
J Antimicrob Chemother 63:i31–i35
9. Maschmeyer G, Beinert T, Buchheidt D, Cornely OA,
Einsele H, Heinz W et al (2009) Diagnosis and antimicrobial therapy of lung infiltrates in febrile neutropenic patients: guidelines of the infectious diseases
working party of the German Society of Haematology
and Oncology. Eur J Cancer 45:2462–2472
10. Skladny H, Buchheidt D, Baust C, Krieg-Schneider F,
Seifarth W, Leib-Mösch C et al (1999) Specific detection of Aspergillus species in blood and bronchoalveolar lavage samples of immunocompromised patients
by two-step PCR. J Clin Microbiol 37:3865–3871
11. Hummel M, Spiess B, Cornely OA, Dittmer M,
Mörz H, Buchheidt D (2010) Aspergillus PCR testing:
results from a prospective PCR study within the
AmBiLoad trial. Eur J Haematol 85:164–169
12. Spiess B, Buchheidt D, Baust C, Skladny H, Seifarth
W, Zeilfelder U et al (2003) Development of a
LightCycler PCR assay for detection and quantification
of Aspergillus fumigatus DNA in clinical samples
from neutropenic patients. J Clin Microbiol 41:
1811–1818
13. Sambrook J, Fritsch E, Maniatis T (1989) Molecular
cloning: a laboratory manual. Cold Spring Harbor
Laboratory Press, Cold Spring Harbor, NY
Application of Polymerase Chain
Reaction and PCR-Based Methods
Targeting Internal Transcribed
Spacer Region for Detection and
Species-Level Identification of Fungi
27
K. Lily Therese, R. Bagyalakshmi, and H.N. Madhavan
Abstract
This chapter focuses on the application of molecular technique based on
polymerase chain reaction (PCR) targeting internal transcribed spacer
(ITS) region for detection and identification of fungi from normally sterile
body fluids and tissue biopsies; detection of nucleotide polymorphisms in
Aspergillus flavus; PCR-based DNA sequencing for identification of nonsporulating molds (NSM); and detection and identification of dermatophytes from dermatological specimens by PCR-based restriction fragment
length polymorphism (PCR-RFLP).
Keywords
ITS region • PCR • PCR-based DNA sequencing • Nonsporulating molds
(NSM) • PCR-based RFLP • Dermatophytes
Introduction
Conventional methods for the detection of fungal
infections are less sensitive because the microbial
threshold is low and the techniques are laborious
and time-consuming. Rapid diagnosis by molecular methods aid in the institution of specific anti-
K.L. Therese (*) • R. Bagyalakshmi
H.N. Madhavan
Larsen and Toubro Microbiology Research Centre,
Kamal Nayan Bajaj Research Centre,
Vision Research Foundation,
41 (Old No. 18) College Road, Chennai,
Tamil Nadu 600006, India
e-mail: drklt@snmail.org
fungal drug and management [1]. Molecular
methods are rapid, extremely sensitive, and
specific. Fungi have a ribosomal DNA (rDNA)
complex that includes a sequence coding for the
18S rDNA gene, an internal transcribed spacer
region 1(ITS1), the 5.8S rDNA gene coding
region, another ITS region called ITS2, and the
sequence coding for 28S rDNA gene [1, 2].
Polymerase chain reaction (PCR) assays have
been developed and applied targeting the 28S
rDNA [3, 4] and 18S rDNA [5]. The ITS region
is a multicopy gene and consists of considerable
variation to differentiate the fungal species.
Several studies that target the ITS region to
detect and identify fungal genome using PCR
and PCR-based methods from clinical specimens
have been carried out. Target genes used in
molecular diagnosis of fungal infections include
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_27, © Springer Science+Business Media, LLC 2013
321
322
single and multicopy nuclear and mitochondrial
genes. Multicopy genes provide a better detection threshold than single-copy genes and are
employed in different molecular assays [6–16].
Molecular diagnostic assays involve the use of
PCR followed by restriction fragment length
polymorphisms (RFLP) [17, 18] to identify the
species and to analyze the strain variations and
DNA sequencing for identification, detection of
nucleotide polymorphisms, and mutations existing in the fungal genome [19–21].
Application of a Semi-Nested
Polymerase Chain Reaction Targeting
the Internal Transcribed Spacer
Region
Protocol for Detection of Panfungal
Genome by Application of snPCR
Targeting ITS Region
Materials
Standard Strains of Fungi
1. C. albicans ATCC 24433
2. C. albicans ATCC 90028
3. C. parapsilosis ATCC 90018
4. C. parapsilosis ATCC 22019
5. C. krusei ATCC 6258
6. C. tropicalis ATCC 750
7. A. fumigatus ATCC 10894
Reagents (Commercially Available QIAGEN
Kit, Germany)
1. Proteinase K
2. Lysis buffer
3. Absolute ethanol
4. Washing buffer-1 and 2
5. Elution buffer
6. Microfuge (Eppendorf, Germany)
7. New sterile disposable polypropylene
microfuge tubes, 1.5 mL
8. Waterbath
Methods
DNA extraction is carried out using QIAamp
DNA extraction kit (Qiagen, Germany) according to the manufacturer’s instruction.
K.L. Therese et al.
1. Pipette 20 ml Qiagen Proteinase K into the
bottom of a 1.5-mL microfuge tube.
2. Add 200 ml of the sample to the microfuge
tube. Use up to 200 ml whole blood, plasma,
serum, buffy coat or body fluids or up to 5 × 106
lymphocytes in 200 ml PBS.
3. Add 200 ml of lysis buffer (AL buffer) to the
sample. Mix by pulse vortexing for 15 s.
4. Incubate at 56 °C in a water bath for 10 min.
(see Note 1)
5. Add 200 ml of ethanol (96–100%) to the sample, mix by gentle pipetting for 15 s.
6. Carefully apply the mixture from step 5 to the
QIAamp mini-spin column (in a 2-mL collecting tube) without wetting the rim. Close the
cap, and centrifuge at 8,000 rpm for 1 min.
Place the QIAamp mini-spin column in a clean
2-mL collecting tube and discard the tube containing the filtrate.
7. Carefully open the QIAamp mini-spin column
and add 500 ml buffer AW1 without wetting
the rim. Close the cap and centrifuge at
8,000 rpm for 1 min. Place the QIAamp minispin column in a clean 2-mL collecting tube
and discard the tube containing filtrate.
8. Carefully open the QIAamp mini-spin column
and add 500 ml buffer AW2 without wetting
the rim. Close the cap and centrifuge at
14,000 rpm for 3 min, followed by an empty
spin at 14,000 rpm for 1 min.
9. Place the QIAamp mini-spin column in a new
1.5-mL microfuge tube and discard the tube
containing filtrate. Carefully open the QIAamp
mini-spin column and add 200 ml buffer AE.
Incubate at room temperature for 1 min, and
then centrifuge at 8,000 rpm for 1 min. Discard
the column and store the DNA at −20 ° C.
Semi-Nested PCR Targeting
the Internal Transcribed Spacer
Region
The snPCR targeting ITS region involves the use
of primers designed by Ferrer et al. [22].
Materials
1. Deoxyribonucleotide triphosphates (dNTPs)—
dATP, dGTP, dCTP, dTTP (10 mM each,
Bangalore Genei, India) stored at −20 °C.
27
Application of Polymerase Chain Reaction and PCR-Based Methods…
2. Thermostable Taq DNA polymerase 3 units/ml
(Bangalore Genei, India) supplied with a10×
PCR buffer (Tris–Cl buffer pH 8.8) which
contains 500 mM KCl, 15 mM Magnesium
chloride.
3. 25 mM Magnesium chloride (Bangalore
Genei, India).
4. Oligonucleotide primers (custom made,
Bangalore Genei, India).
5. PCR tubes thin walled (0.5 mL, 0.2 mL,
Axygen, USA).
6. Thermal
cycler
(Eppendorf,
Germany
PerkinElmer 2700, USA).
Procedure
1. A 50-ml reaction, consists of 8 ml of dNTPs
(200 mmol), 5 ml of 1× PCR buffer (1.5 mM
MgCl2 ,50 mM KCl, 10 mM Tris–Cl,0.001%
gelatin), 6 ml of 25 mM MgCl2 (1 in 10 diluted
to get a final concentration of 3 mM), 1
microlitre of forward primer (10 picomoles)
of, ITS1—5¢-TCC GTA GGT GAA CCT GCG
G-3¢and 1 microlitre of reverse primer ITS4—
5¢-TCC TCC GCT TAT TAT GC-3¢ targeting
ITS region, 1 unit of Taq polymerase, 18.7
microlitre of milli-Q water and 10 ml of template DNA [22].
2. Amplification is allowed to occur in a PCR
machine (Perkin Elmer Model 2700). The first
round of amplification yields 520–611 bp
product depending on the fungal species present in the clinical specimen.
3. The thermal profile includes initial denaturation
at 95 °C for 5 min followed by 35 cycles of
denaturation at 95 °C for 30 s, annealing at
55 °C for 60 s and extension at 72 °C for 60 s
followed by final extension at 72 °C for 6 min.
4. This is then subjected to semi-nested
amplification using same PCR conditions as
that of the first round with forward primer
ITS86—5¢-GTG AAT CAT CGA ATC TTT
GAA C-3¢ and reverse primer ITS 4 as indicated above.
5. 5 ml of amplified product is transferred from
the first round to the second round and subjected to amplification.
6. The thermal profile consists of initial denaturation at 95 °C for 5 min followed by 35 cycles
323
of denaturation at 95 °C for 30 s, annealing at
55 °C for 30 s, and extension at 72 °C for 30 s
followed by final extension at 72 °C for 5 min.
Detection of Amplified Products
1. Agarose molecular grade (SRL, India).
2. Tris–borate EDTA buffer (TBE buffer) consisting of 50 mM Tris, 50 mM boric acid, and
1 mM EDTA. A 1-in-10 dilution is made and
used to dissolve agarose.
3. Ethidium bromide −0.5 mg/mL stock.
4. Tracking dye: Bromophenol blue (Hi Media,
India) consisting of 0.1% Bromophenol blue,
40% (w/v) sucrose, 0.1 M EDTA. 1-in-10
dilution is made with electrophoresis buffer
and used.
5. Parafilm for aliquoting the amplified
products.
6. Horizontal electrophoresis equipment with
power pack (Sri Balaji Scientific Supplies,
India) along with gel-casting accessories.
7. Molecular weight marker Hinf I digest of
fX174 Bacteriophage DNA.
8. Gel documentation system (Vilber Lourmat,
France).
Preparation of 2% Agarose Gel
1. Clean the gel trough with ethanol and seal
the ends with cellophane tape with the combs
placed within groove to form wells.
2. Prepare 2% agarose gel by dissolving 1 g of
agarose in 50 mL of 1× TBE buffer. The agarose is dissolved by boiling it in a microwave
oven and add 8 ml of ethidium bromide.
(Final concentration in the gel 0.5 mg/mL).
3. Mix thoroughly and pour onto the trough.
4. The gel is allowed to solidify for 30 min.
5. After the gel gets solidified, the combs are
carefully removed.
6. Place the gel in the submarine electrophoresis tank and remove the combs.
7. Mix 10 microlitre of PCR amplified product
using 2 microlitre of Bromophenol blue in a
parafilm strip.
8. Load the negative control, samples, positive
control, and then the molecular weight marker.
9. Carry out the electrophoresis at 100 V for
half an hour.
324
K.L. Therese et al.
Table 27.1 Amplicon size in base pairs of fungi
Fungi
Candida albicans
Candida parapsilosis
Candida glabrata
Candida krusei
Aspergillus flavus
Aspergillus fumigatus
Aspergillus niger
Aspergillus terreus
Fusarium solani
Fusarium oxysporum
Cryptococcus neoformans
Scedosporium apiospermum
Alternaria alternata
I Round
586
524
820
510
595
596
599
608
569
544
556
611
570
II Round
282
270
360
294
300
299
300
308
286
283
320
329
292
10. Carry out the gel documentation using the
gel documentation system Vilber Lourmat,
France and analyze using BioID software.
11. The amplified product size varies according
to the species of fungi and the respective
amplicon size of representative fungi is given
in Table 27.1.
Sensitivity of snPCR Targeting ITS
Region for Detection of Fungal Genome
1. 1 ml of the extracted DNA is dissolved in
999 ml of water and quantified spectrophotometrically at 260 and 280 nm.
2. The reading at 260 nm gives the nucleic acid
concentration. The ratio of readings at OD
260/280 nm gives the purity of the nucleic
acid.
3. The sensitivity of snPCR is determined using
serial tenfold dilutions of standard strain of
C. albicans ATCC 24433.
4. 10 mg of C. albicans and A. fumigatus are used
as the template DNA.
Specificity of snPCR
The specificity of snPCR is determined using
DNA extracts of microorganisms; C. albicans
(ATCC 90028), C. tropicalis (ATCC 750),
C. parapsilosis (ATCC 90018), C. krusei (ATCC
6258), A. flavus (ATCC 204304), A. fumigatus
(ATCC 10894), A. niger (ATCC16404), F. solani
(ATCC 36031), laboratory isolates of A. terreus,
Curvularia and Alternaria species, S. aureus
(ATCC 12228), Pseudomonas aeruginosa (ATCC
27853), M. tuberculosis (H37Rv), M. fortuitum
(ATCC 1529), M. chelonae (ATCC 1524) and
laboratory isolates of Nocardia asteroides,
Actinomyces species, Herpes Simplex virus
(ATCC 733 VR), Acanthamoeba polyphaga
(ATCC 30461), and human leukocyte DNA.
Standardization of PCR: Sensitivity
of snPCR Targeting ITS Region
1. The expected sensitivity of ITS primers is
1–10 fg of C. albicans DNA (single cell of
C. albicans) and 10 fg of A. fumigatus and
F. lichenicola DNA.
2. The ITS PCR is specific amplifying only the
fungal DNA. The application of semi-nested
PCR targeting ITS region on clinical specimens is shown in Fig. 27.1.
PCR targeting ITS region of fungi is a rapid
and sensitive diagnostic test used for specieslevel identification of fungi from clinical
specimens [23].
Application of Semi-Nested PCR
Targeting ITS Region to Determine
the Nucleotide Polymorphisms
Associated with Ocular Isolates
of Aspergillus flavus
Strain typing of medically important fungi (i.e.,
the ability to identify them to the species level
and to discriminate among individuals within
species) has been galvanized by new methods of
tapping the tremendous variation found in fungal
DNA. There are several methods like multilocus
enzyme electrophoresis, electrophoretic karyotype analysis, RFLP, randomly amplified polymorphic DNA, sequence confirmed amplified
region analysis, DNA fingerprinting, PCR-based
DNA sequencing with repetitive sequences
27
Application of Polymerase Chain Reaction and PCR-Based Methods…
325
Fig. 27.1 Agarose gel electrophoretogram showing application of ITS PCR on ocular specimens. Lane 1: Negative control II round; Lane 2: Negative control I round; Lane 3:
AH-positive (Aspergillus flavus) (300 bp); Lane 4:
VF-positive Fusarium solani (286 bp); Lane 5: VF-negative;
Lane 6: AH-negative; Lane 7: AH-positive Aspergillus terreus (308 bp); Lane 8: VF positive Candida tropicalis
(293 bp); Lane 9: VF positive (Aspergillus flavus) (300 bp);
Lane 10: AH-positive Candida albicans (282 bp); Lane 11:
AH-positive A. niger 300 bp; Lane 12: Positive control :
Candida albicans ATCC 24433 (282 bp); Lane MW:
Molecular weight marker Hinf I digest of Phi X174 bacteriophage DNA. Note: The species identity detected by ITS PCR
was further confirmed by PCR-based DNA sequencing
available for studying the strain variation existing among the species of fungi rRNA genes. The
use of DNA sequence diversity in the ribosomal
regions as an aid to species identification has
been exploited using PCR amplification of targets followed by either fragment length analysis
[19], DNA probe hybridization [20], or DNA
sequence analysis.
the method provided in Semi-Nested PCR
(snPCR) Targeting the ITS Region.
Study on Strain Variations
of Aspergillus flavus
In a study conducted in the authors’ center, seven
ocular isolates of A. flavus from aqueous humor
(AH, 2), vitreous fluid (VF, 2), corneal scraping
(1), eviscerated material (1), corneal button (1)
were isolated from four patients clinically suspected to have fungal endophthalmitis.
Molecular Microbiological Investigations:
DNA Extraction from Isolates
DNA extraction from clinical specimens and fungal isolates is done using QIAamp kit as described
in “Methods”.
PCR Targeting ITS Region
PCR targeting ITS region is carried out on culture isolates of Aspergillus flavus according to
PCR-Based Restriction Fragment
Length Polymorphism
1. PCR-RFLP is carried out on ITS amplicons
(first round products) using restriction enzyme
Hae - III.
2. For a 25 ml reaction, 2.5 ml of Buffer C, 0.5 ml
of Hae – III enzyme and 10 ml of amplified
product are added and incubated at 37 ° C for
3 h.
3. The digested products are loaded on 4% agarose gel incorporated with ethidium bromide
and the electrophoresis is carried out at 100 V
for 45 min.
4. The digested pattern (385,188,82 bp) is visualized and documented using gel documentation system (Vilber Lourmat, France).
DNA Sequencing of ITS Amplicons
DNA sequencing of ITS amplicons is carried out
after purification of amplified products.
In our study, all A. flavus isolates showed similar
pattern of digestion with Hae-III. In an earlier
study Henry et al. [24]. have reported a variation
of 1% in epidemiological analysis of A. flavus
strains. Alignment of contiguous fungal sequences
demonstrated that both single nucleotide differences and short lengths of sequence diversity due
to insertion or deletion existed in the ITS regions
326
among the pathogenic A. flavus strains. A. flavus
isolates in the study had a BLAST score of 97.7%
identity with the standard strain of A. flavus
(ATCC 16883)—GenBank accession no.
AB008415. The inspection of BLAST alignments generated with A. flavus ITS1 and ITS 2
data from GenBank revealed that many A. flavus
sequences in the data base had truncated ends
and/or heterogeneities at positions found to be
conserved at the subgeneric level among reference sequence of type and authenticated culture
collection strains. The study by Bagyalakshmi
et al. [25] on A. flavus isolates revealed a variation of 2.3% as compared with the standard strain
of A. flavus. The sequences have been deposited
in GenBank and the accession nos. DQ683118,
DQ683119, DQ683120, DQ683121, DQ683122,
DQ683123, DQ683124 have been assigned for
the isolates. The nucleotide polymorphisms existing among A. flavus strain in the study was novel
and the first to be reported in literature [25].
Application of PCR-Based DNA
Sequencing Targeting ITS Region
for Genus- and/or Species-Level
Identification of Nonsporulating
Molds
In the clinical laboratory, isolates that cannot be
identified by reproductive structures are described
as Mycelia sterilia or nonsporulating molds
(NSM), a name indicating a filamentous fungus
that displays no distinguishing phenotypes recognized by routine clinical laboratory analysis.
Such fungi can be identified using PCR-based
DNA sequencing.
Identification of NSM by PCR-based
DNA Sequencing
1. DNA extraction from fungal isolates of NSM
should be done using Qiagen kit according to
the manufacturer’s instructions (see Methods).
2. PCR is carried out as per the procedure given
in “Semi-Nested PCR Targeting the ITS
Region”.
K.L. Therese et al.
Table 27.2 Details of DNA sequencing technique (the
four nucleotide bases with the respective acceptor dyes
and color emission)
Terminator
A
C
G
T
Acceptor dye
dR6G
dROX
dR110
dTAMRA
Color of raw data
on ABI PRISM310
electrophoretogram
Green
Red
Blue
Black
3. The amplified products are subjected to cycle
sequencing, purified, and then subjected to
DNA sequencing.
Sequencing in ABI Prism 310/3100
AVANT Genetic Analyzer
1. The sequence of the PCR amplified DNA is
deduced with the help of the ABI Prism.
2. 310/3100 AVANT genetic analyzer that works
based on the principle of Sanger dideoxy
sequencing.
3. The fluroscent-based detection by automated
sequencer adopts the Sangers method and
incorporates the fluorescent dyes into DNA
extension products using 5¢-dye labeled primers or 3¢-dye labeled ddNTPs (dye terminators
called commercially as RR MIX).
4. Each dye emits light at a different wavelength
when excited by an argon ion laser (Table 27.2).
All four colors and, therefore, all four bases
can be detected and distinguished in a single
gel lane or capillary injection.
5. The protocol and thermal profile for cycle
sequencing are provided in Tables 27.3 and
27.4, respectively
6. The amplified products with the dye at the terminated 3’end are subjected to capillary electrophoresis by an automated sample injection.
7. The emitted flurorescence from the dye labels
on crossing the laser area is collected in the
rate of one per second by cooled, chargecoupled device camera at particular wavelength bands (virtual filters) and stored as
digital signals on the computer for processing
that are analyzed by software called as the
27
Application of Polymerase Chain Reaction and PCR-Based Methods…
Table 27.3 Protocol for cycle sequencing
Components
Amplfied products
Sequence buffer
Primer (2 pmol/ml)
RRMIX
Water
Volume (ml)
28S rRNA
amplicons
1.0
3.0
2.0
1.0
3.0
Volume (ml)
ITS
amplicons
1.0
2.5
2.0
1.5
3.0
Table 27.4 PCR conditions for cycle sequencing
PCR step
Initial denaturation
Denaturation
Annealing
Extension
Temperature (°C)
96
96
50
60
Time
1 min
10 s
5s
4 min
The reaction to be carried out for 25 cycles.
sequence analysis softwares (Sequence
Navigator in ABI 310 and seqscape manager
in ABI 31000 AVANT machine).
8. The ABI AVANT genetic analyzer can be
upgraded from 4 capillary to 16 capillary so
that facilitates the electrophoresis of 16 samples at a given time.
Purification of Extension Products
The products are purified to remove the unincorporated dye terminators before sujecting the samples to capillary electrophoresis.
Procedure
1. 2 ml of 125 mM EDTA and 2 ml of 3 M sodium
acetate (pH 4.8) 10 microlitre of Milli Q water
is mixed with 10 microlitre of cycle sequenced
prodcuts followed by the addition of 50 ml of
absolute ethanol and incubated at room temperture for 15 min follwed by centrifugation at
12,000 rpm for 20 min to precipitate the
amplified product and remove the unutilized
ddNTPs, primer (short-length molecules),
etc.
2. The pellet is washed twice with 70% ethanol
at 12,000 rpm for 10 minutes followed by air
drying. The purified samples is suspended in
formamide and subjected for capillary electro-
327
phoresis in ABI PRISM 310/3100 genetic
analyzer.
3. The sequence is then analyzed in Sequence
Navigator software (version 1.0.1; ABI Prism
310) or seqscape manager (version 2.1; ABI
Prism 3100 AVANT).
The use of PCR-based DNA sequencing has several advantages over the conventional methods in
terms of rapidity, accuracy, and definite
identification. Out of the 50 NSM fungal isolates, 27 were found to be emerging pathogens
involving 7 genera (Botryosphaeria species,
Lasiodiplodia species, Thielavia tortuosa,
Glomerulla singulata, Macrophomina phaseolina, Rhizoctonia bataticola, Podospora species)
and 23 as established pathogens involving 8 genera (Aspergillus, Fusarium, Bipolaris, Pythium,
Cochliobolus, Exserohilum, Pseudallescheria
and Scedosporium species) (Table 27.5) [26].
Application of Polymerase Chain
Reaction-Based Restriction Fragment
Length Polymorphism for SpeciesLevel Identification of Dermatophytic
Fungi
Molecular Microbiological
Investigations
DNA Extraction from Isolates
DNA extraction from clinical specimens and fungal isolates is done using QIAamp kit as described
in “Methods”.
PCR Targeting ITS Region
PCR targeting ITS region is carried out on dermatological specimens according to the method
provided in “Semi-Nested PCR Targeting the ITS
Region”.
PCR-Based Restriction Fragment Length
Polymorphism
1. PCR-RFLP is carried out on ITS amplicons
(first round products) using restriction enzyme
Hae-III.
328
K.L. Therese et al.
Table 27.5 Nonsporulating molds identified by PCR-based DNA sequencing targeting ITS region
Newer emerging pathogens identified
by PCR-based DNA sequencing
targeting ITS region
Emerging pathogens 27
Botryosphaeria species 10
Botryosphaeria rhodina 8
Botryosphaeria dothidea 1
Botryosphaeria species 1
Lasiodiplodia theobromae 2
Rhizoctonia bataticola 5
Glomerulla singulata 3
Cochliobolus species 3
Cochliobolus species 2
Cochliobolus heterophrynus 1
Macrophomina phaseolina 2
Podospora species 1
Thielavia tortuosa 1
Established pathogens: 27
Pythium insidiosum 9
Fusarium species 7
Fusarium solani 2
F. solani 1, F. falciforme 2b 3
Fusarium proliferatum 2
Exserohilum species 1
Aspergillus terreus 1
Aspergillus fumigatus 1
Scedosporium species 1
Pseudallescheria species 1
Bipolaris species 2
Ocular specimens
GenBank accession number
Corneal scraping 4a
Conjunctival scraping 1a
Corneal necrotic tissue 1a
Corneal button 2a
Corneal button 1
Corneal scraping 1
Corneal scraping 1
Eviscerated material 1
Corneal scraping 4
Eviscerated material 1
Corneal scraping 1
Corneal button 1
Eviscerated material 1
Corneal scraping 2
Corneal button 1
Corneal scraping 2
Corneal scraping 1
Corneal button 1
EF446281
EF446291
EF446282
EF446289
EF446288
EF446287
Corneal scraping 6, Corneal
button 2, Vitreous aspirate 1
Corneal scraping 2
Corneal scraping 3
Corneal scraping 1, Corneal button 1
Corneal scraping 1
Corneal scraping 1
Corneal button 1
Infected suture 1
Corneal button 1
Corneal scraping 1
Donor corneal rim 1
EF446283
EF446284
a
The five isolates of Botryosphaeria rhodina were obtained from the same patient (corneal scraping 2, conjunctival
scraping, corneal button, necrotic tissue)
b
The three Fusarium species were identified by species specific PCR as F. solani (1) and F. falciforme (2) by PCR-based
DNA sequencing on 28S rRNA gene
2. For a 25 ml reaction, 2.5 ml of buffer C, 0.5 ml
of Hae-III enzyme (Bangalore Genei, India),
12 microlitre of milli-Q water and 10 ml of
amplified product are added and incubated at
37 °C for 3 h.
3. The digested products are loaded on 4% agarose gel incorporated with ethidium bromide
and the electrophoresis is carried out at 100 V
for 45 min.
27
Application of Polymerase Chain Reaction and PCR-Based Methods…
329
4. The digested pattern (Microsporum gypseum
420, 95 bp, Trichophyton rubrum (300, 200,
95 bp), Epidermophyton floccosum (350,
95 bp) [27] is visualized and documented
using gel documentation system (Vilber
Lourmat, France).
PCR-based RFLP is a rapid method for detection
and identification of dermatophytes (24 h) as
compared to conventional culture (which may
require up to 21 days for growth) and always may
not result in successful isolation.
10. Gel incorporated with ethidium bromide and
pipette tips should be wrapped in a foil, discarded in a labeled container and disposed
off by incineration.
11. While transferring the first round products to
the second round, always the negative control should be transferred first followed by
the specimens and finally the positive control
to avoid amplicon carryover.
12. The place of transfer of negative controls and
positive controls should be physically
separated.
Notes
Acknowledgments The financial support in the form of
a Research Grant by Department of Science and
Technology, (DST) Government of India, and the infrastructure facility provided by Vision Research Foundation,
Chennai are gratefully acknowledged.
1. Depending on the purulence of body fluids
and thickness of tissue biopsies incubation
can be extended up to 1 h at 56 °C. Before
proceeding with the next step, it is absolutely
essential that complete digestion of the specimen is attained. When complete digestion
takes place, the specimen becomes clear.
While extracting DNA from tissue specimens, tissue lysis buffer provided in the kit
should be used.
2. Wear laboratory coat, gloves, safety goggles
while handling clinical specimens.
3. Extraction of DNA and preparation of PCR
premix should be carried out in a Laminar
flow hood.
4. While extracting DNA from standard strains
it is strictly recommended to handle the fungal strains in a level III biosafety cabinet.
5. For all molecular mycological procedures,
autoclaved deionized water should be used
or molecular grade water available commercially can be used.
6. New pre-sterilized vials to be used for PCR.
7. The pre-amplification, amplification, and
post-amplification areas need to be physically
separated and dedicated pipettes should be
used for DNA extraction, preparation of PCR
cocktail, and post-amplification analysis.
8. Filter-guarded tips should be used to avoid
cross contamination.
9. Wear face mask, cap, and gloves while handling ethidium bromide since it is
carcinogenic.
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Real-Time PCR Assay in Fungi
28
Naomichi Yamamoto
Abstract
Traditional growth-based methods for characterizing environmental fungi
are biased by selection of culture media and incapable of detecting noncultivable fungi, which still retain allergenicity and/or pathogenicity.
Meanwhile, real-time quantitative polymerase chain reaction (qPCR)-based
methods have been recently developed and used to characterize fungal
concentrations in environmental samples such as air and house dust. As
qPCR-based methods are independent of fungal cultivability or viability,
they are expected to be toxicologically more relevant than conventional
growth-based methods for assessing health effects caused by environmental fungi. This chapter presents protocols for the collection of environmental fungal samples and subsequent qPCR analysis.
Keywords
Fungi • qPCR • Allergens • Spores • DNA • 18S rRNA
Introduction
Growth-based methods have been traditionally
and widely used to detect environmental fungi [1,
2]. However, the results of these methods are
often biased by selection of culture media [1].
Furthermore, identification of fungal species by
growth-based methods is often based on observation of their microscopic and macroscopic
N. Yamamoto (*)
Department of Environmental Health, Graduate School
of Public Health, Seoul National University,
1 Gwanak-ro, Gwanak-gu, Seoul 151-742, Korea
e-mail: naomichi.yamamoto@yale.edu
morphologies, and accuracy of the identification
relies on operator’s experience and level of expertise. Most importantly, growth-based methods
are incapable of detecting non-cultivable microorganisms [3, 4], which are still potential allergens and pathogens [5]. For instance, research
has indicated that growth-based methods underestimated concentrations of fungi in dust by two
to three orders of magnitude compared to realtime quantitative polymerase chain reaction
(qPCR) assays [3].
In recent years, several studies have utilized
qPCR-based methods to characterize fungal concentrations in environmental samples such as air
[6–9] and house dust [3, 4, 10–12]. qPCR-based
methods are expected to be toxicologically more
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_28, © Springer Science+Business Media, LLC 2013
331
332
relevant than conventional growth-based methods
because they are not affected by cultivability or
viability of the fungi. Although qPCR-based
methods have become common in recent years,
care must be taken in their implementation as the
inadequate reporting of experimental detail and
the use of flawed protocols lead to the technically
inappropriate results [13]. Here I introduce a protocol for measurements of fungi in air by qPCR
[9, 14]. Sections include creation of a DNA standard, Andersen impactor environmental sampling, personal sampling, DNA extraction, qPCR
protocol, avoiding qPCR inhibition, and precision, accuracy, and method detection limit (MDL)
of qPCR.
Materials
1. Sterile distilled water.
2. Pipettes: 0.1–10 mL, 10–100 mL,
100–1,000 mL.
3. Disposable filter pipette tips: 0.1–10 mL,
10–100 mL, 100–1,000 mL.
4. Petri dishes: 90 mm diameter.
5. Disposable polypropylene tubes: 15-mL
conical.
6. Disposable polypropylene microcentrifuge
tubes: 1.5-mL conical.
7. Sterile nylon-flocked swabs.
8. Ethanol (70%).
9. Vortexer.
10. Tween 20 solution (0.1% in deionized
water).
11. Crystal violet.
12. Microscopic counting chamber (depth
20 mm).
13. Optical microscope (×400).
14. Tally counter.
15. Microcentrifuge.
16. DNA extraction kit; e.g., PowerSoil DNA
Isolation Kit (Mobio Laboratories, Carlsbad,
CA, USA).
17. Disposable polypropylene screw-capped
microcentrifuge tubes: 2.0-mL conical.
18. Sterile glass beads: 0.1- and 0.5-mm
diameters.
19. Bead beater; e.g., Model 607 (Bio Spec
Products Inc., Bartlesville, OK, USA).
N. Yamamoto
20. TE buffer (10 mM Tris–HCl, 1 mM EDTA,
pH 8.0).
21. Sterile tweezers.
22. For airborne fungal sampling: eight-stage
nonviable Andersen samplers or the Personal
Environmental Monitor Model 200 (PEM)
(New Star Environmental, Inc., Roswell,
GA, USA). The Andersen sampler uses
quartz glass fiber filters (81-mm diameter;
New Star Environmental Inc., Roswell, GA,
USA) while the PEM uses polycarbonate
track etch (PCTE) membrane filters (37-mm
diameter, 0.8-mm pore size; SKC Inc., Eighty
Four, PA, USA).
23. Sterile scissors.
24. Molecular biology grade water.
25. TaqMan Universal PCR Master Mix; e.g.,
Applied Biosystems, Carlsbad, CA, USA.
26. Primers.
27. TaqMan probes; e.g., The BHQplus™ probe
with FAM dye (Biosearch Technologies Inc.,
Novato, CA, USA).
28. Real-time PCR well reaction plate; e.g.,
Applied Biosystems MicroAmp® Optical
96-Well Reaction Plate.
29. Real-time PCR clear adhesive seal.
30. Centrifuge; e.g., 5804R (Eppendorf,
Hamburg, Germany).
31. Real-time PCR system; e.g., Applied
Biosystems ABI 7500.
Methods
Preparation of Standard Fungal DNA
Samples
This method has been used to prepare standard
fungal DNA samples containing known numbers
of fungal spores for use in qPCR. Fungal species
producing unicellular amerospores such as
Aspergillus fumigatus, Cladosporium cladosporioides, and Penicillium chrysogenum are considered here.
1. Swab and re-suspend the spores in 10 mL of
ethanol (70%) in a 15-mL tube. Harvest only
the spores of the entire surface of fungal colonies grown on a 90-mm petri dish. Multiple
dishes may be used if needed.
28
Real-Time PCR Assay in Fungi
2. Aliquot the suspension into 1.5-mL microcentrifuge tubes to make several subsamples.
Transfer 1-mL suspension into each microcentrifuge tube. Vortex the 15-mL conical tube
each time before transfer to prevent gravitational settling of the spores in the suspension.
3. Centrifuge the microcentrifuge tubes at
10,000×g for 3 min to pellet the spores.
4. Remove ethanol supernatant.
5. Select 5 subsamples for cell enumeration by
microscopy. The remaining subsamples are
used for DNA extraction. The samples used
for the DNA extraction are added with the
Mobio power lysis solution (750 mL), and the
suspensions are transferred to 2-mL screwcapped microcentrifuge tubes. For DNA
extraction, follow the steps in “Extraction of
Fungal DNA”, starting from step 2.
6. For cell enumeration, add 1-mL Tween 20
solution (0.1%) and 10 mL crystal violet to
each subsample tube.
7. Pipette 10 mL of the suspension into the microscopic counting chamber.
8. Enumerate the spores on an entire area of the
counting chamber (=1 mm2) using an optical
microscope (×400) and a tally counter.
Calculate the total number of fungal spores in
each subsample. 108–109 spores per sample
are desirable. Dilution of the suspension may
be necessary. The optimum counts per
1/25 mm2 are in the 5–15 cell range.
333
4. Cut each filter to one-eighth piece and shred
them into smaller pieces using sterile scissors
and tweezers.
5. Place all the shredded pieces of the oneeighth filter to 2-mL screw-capped microcentrifuge tube and follow the steps for DNA
extraction in “Extraction of Fungal DNA”,
starting from step 1.
Collection of Airborne Spores by
Personal Environmental Monitor
The PEM is used to collect airborne fungi in the
PM2.5 or PM10 size ranges on a polycarbonate
filter. This sampler is suitable to assess personal
exposures to airborne fungi in the PM2.5 or
PM10 fractions, which are generally respirable or
inhalable fractions, respectively.
1. Load a 37-mm PCTE filter on the PEM.
2. Operate the PEM with a specified air flow
rate. The sampling time will vary depending on the expected fungal concentrations
in the air.
3. Cut the PCTE filter to one-fourth piece using
sterile scissors.
4. Place the cut filter to 2-ml screw-capped
microcentrifuge tube, and follow the steps for
DNA extraction in “Extraction of Fungal
DNA”, starting from step 1.
Extraction of Fungal DNA
Collection of Airborne Spores by
8-Stage Nonviable Andersen Sampler
The 8-stage nonviable Andersen sampler allows
for collection of size-fractionated airborne fungal
cells on glass fiber filters [15]. This sampler is
suitable to characterize particle size distributions
of airborne fungi.
1. Load glass fiber filters on each stage of the
Andersen sampler using sterile tweezers.
2. Operate the sampler with an air flow rate of
28.3 L min-1. The sampling time will vary
depending on the expected fungal concentrations in the air.
3. After sampling, recover the filters and place
them in 90-mm Petri dishes.
This method has been used to extract DNA from
fungal cells in environmental samples such as
those collected on air filters from the above or
other methods. A protocol by the PowerSoil DNA
Isolation Kit is modified to use for the isolation
of fungal DNA from air samples. This kit is suitable to remove PCR inhibitors from environmental samples.
1. Add 750 mL of Mobio lysis solution.
2. Add the Mobio power beads (1.0 g) supplemented with 0.1-mm diameter glass beads
(300 mg) and 0.5-mm diameter glass beads
(100 mg).
3. Disrupt fungal cells for 5 min at 3,450 rpm by
a bead beater.
334
4. Follow the Mobio’s protocol (see Note 1) and
elute DNA into 50 mL of TE buffer.
5. For the standard DNA extracted from the samples containing known numbers of fungal
spores prepared according to the steps in
Preparation of Standard Fungal DNA Samples,
make tenfold dilution series from 1/10 to
1/107. Calculate the numbers of fungal spores
in 1 mL of each diluted standard (see Note 2).
N. Yamamoto
3.
4.
5.
6.
Quantitation of Fungal Spores
Using Real-Time qPCR
7.
The ribosomal DNA (rDNA) is the most commonly targeted region for fungal PCR [16].
Sensitive fungal detection is possible by targeting rDNA since there are large numbers of gene
copies present in each fungal genome. For
instance, A. fumigatus has an average of 54 copies (range from 38 to 91 depending on the strain)
of rDNA per genome [17] (see Note 3). Selecting
species-specific rDNA regions for PCR primer
sites allows for species-specific fungal detection.
For instance, the U.S. Environmental Protection
Agency (EPA) has provided the DNA sequences
that can be used for qPCR primers and probes to
characterize more than 130 of the major indoor
air fungal species.1 [18] The following qPCR
protocol can be used with primers and probes
reported by the U.S. EPA for species-specific
fungal quantification. This method uses TaqMan
chemistry. Here the ABI 7500 is used as a realtime PCR system, but other instruments with
equivalent functions may also be used.
1. Prepare a sufficient amount of TaqMan reaction mixture for the number of samples to be
analyzed. Each 24 mL of the master mix contains 1× TaqMan Universal PCR Master Mix,
1 mM of each primer, and 0.08 mM of the
probe. Vortex and centrifuge the mixture,
and distribute 24 mL aliquots into each well
of the reaction plate.
2. Add 1 mL of the DNA samples and standard
DNA to the appropriate well of the reaction plate. The samples are in triplicate. The
tenfold dilution series (from 1/10 to 1/107) of
8.
1
http://www.epa.gov/nerlcwww/moldtech.htm.
9.
10.
11.
12.
the standard DNA prepared according to the
steps in Extraction of Fungal DNA and the
no-template control; i.e., molecular biology
grade water, are used for qPCR calibration.
Each standard is in duplicate.
Use a clear adhesive seal to seal the PCR
well reaction plate.
Centrifuge the plate briefly.
Turn on the real-time PCR machine.
Load the reaction plate in the real-time PCR
machine.
Follow the procedure specified by the manufacturer of the real-time PCR system (e.g.,
ABI 7500 system).
Enter the information of each sample and
standard including concentrations of each
standard and the names of each sample using
the ABI 7500 software. For the standard,
enter the number of fungal spores in 1 mL
calculated in “Extraction of Fungal DNA”,
starting from step 5.
Run the appropriate thermal condition: 50 °C
for 2 min, 95 °C for 15 min of initial denaturation and 45 cycles of 95 °C for 15 s of dissociation, and 60 °C for 1 min of annealing
and extension. For the ABI 7500 system, use
the standard mode. The fast mode is not
recommended.
When cycling is complete, create a calibration plot and calculate the concentrations of
each sample. For the ABI 7500 system, this
can be done by selecting “Auto Ct” as an
analysis setting and clicking the “Analyze”
button on the sheet of “Amplification Plot.”
Use the auto function to calculate cycle
threshold (Ct) values.
Export the file containing the concentration
results (e.g., csv format).
Process the data using spreadsheet software
(e.g., MS-Excel) (see Note 4).
PCR Inhibition Assay
PCR can be inhibited by a wide range of materials including those originating from environmental compounds (e.g., phenolic compounds,
humic acids, and heavy metals), constituents of
microbial cells, non-target DNA, etc. [19]. PCR
28
Real-Time PCR Assay in Fungi
inhibitors affect amplification efficiencies and
lead to inaccurate qPCR measurements [20]. To
test for PCR inhibition, sample extracts are spiked
with subsets of the standard fungal DNA, and
standard curves are produced according to the
steps described in PCR Inhibition Assay. The
standard DNA used to spike the samples should
have a significantly higher concentration than
any fungal DNA present in the sample itself. The
amplification efficiencies with and without spiking the sample extracts are compared to assess
the presence of PCR inhibitors. PCR inhibition
should be mitigated if a significant reduction in
the amplification efficiencies is observed.
Dilution of the sample extracts is a simple method
that can increase amplification efficiencies as it
dilutes inhibiting materials, although it also
dilutes target DNA [6]. Alternatively, the samples
may be re-analyzed using the remaining pieces of
air filters with special care taken during the washing step of the DNA extraction (see Note 1).
Accuracy, Precision, and Method
Detection Limits of Fungal
Measurement by qPCR
Accuracy of fungal measurement by qPCR is
limited by the efficiency of DNA extraction from
cells and whole-cell recovery from sampling air
filters while precision is separated into the total
precision (reproducibility) and precision associated with the analytical instrument (instrument
repeatability). For the MDL, a probability distribution is used to estimate the qPCR MDL as PCR
detection is based on a logarithmic signal
amplification and is binary, i.e., either positive or
negative. Further details about methods to determine the accuracy, precision, and MDLs of fungal measurement by qPCR were reported in our
previous work [12, 14].
335
ethanol-based washing buffer, is critical to
eliminate all traces of residual ethanol.
2. For example, if there are 108 spores in the
standard sample prepared according to the
steps in Preparation of Standard Fungal DNA
Samples, you can assume 2 × 106 spores in
1 mL of the extract as a final elution volume of
the extract is 50 mL. The tenfold dilution series
(from 1/10 to 1/107) of this extract will contain
2 × 105 to 2 × 10-1 spores in 1 mL of each dilute,
respectively.
3. Variations in the numbers of rDNA gene copies as well as those in the numbers of nuclei in
a fungal cell [21, 22] can lead to the biases in
the qPCR measurements. Regardless of these
uncertainties, we use a cell count as a basis of
qPCR calibration since it allows for more
intuitive data presentation. Alternatively, you
may calibrate qPCR based on the number of
gene copies. In this case, synthesized DNA
oligomers with known sequences containing
the region to be amplified, including the sites
for forward and reverse primers, can be used
as standard DNA. The number of gene copies
in the synthesized DNA oligomers can be estimated by the following [23]: gene copy number = DNA content (pg) × 0.978 × 109 (bp/pg)/
genome size (bp). The mass of synthesized
DNA oligomers can be obtained by spectrophotometer or fluorescent methods.
4. The DNA quantities, as a basis of the number
of fungal cells, obtained here are those in 1 mL
of the extract. As fungal DNA is eluted in
50 mL of TE buffer in the extraction process,
these values must be multiplied by a factor of
50 to obtain fungal quantities on the analyzed
filters (i.e., one-eighth of glass fiber filter for
the Andersen sampler and one-fourth of polycarbonate filter for the PEM).
Acknowledgment I thank Karen Dannemiller at Yale
University for invaluable comments on the manuscript.
Notes
1. Care must be taken in the washing step of the
DNA extraction as residual ethanol in the
washing buffer can inhibit PCR. The second
spin of the empty columns, after removing the
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units in outdoor and indoor environments in
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(2002) Profiles of airborne fungi in buildings and outdoor environments in the United States. Appl Environ
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3. Meklin T, Haugland RA, Reponen T, Varma M,
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6. Yamamoto N, Kimura M, Matsuki H, Yanagisawa Y
(2010) Optimization of a real-time PCR assay to
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fungi measurements by active and passive sampling
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Auvinen P, Nevalainen A et al (2008) Analysis of fungal flora in indoor dust by ribosomal DNA sequence
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Quantitative Sampling Methods for
the Analysis of Fungi: Air Sampling
29
Mary C. O’Loughlin, Katherine D. Turner,
and Kevin M. Turner
Abstract
Quantitative sampling of fungi can be carried out in a variety of ways for
a large array of purposes. These include approaches such as tape sampling,
settled dust sampling, bulk material sampling, and air sampling followed
by macroscopic analysis, polymerase chain reaction (PCR), or immunochemical methods for quantitation. Air sampling is widely used in a variety of industries and settings as a means of isolating material for the
identification and potential enumeration of fungal strains thus we have
chosen to discuss this approach to sampling in detail in this chapter. PCR
overcomes the limitations of traditional culturing and macroscopic methods as it is not dependant on culturability or viability of the microorganisms. This chapter discusses a general approach to carrying out such air
sampling and PCR-based quantitation.
Keywords
Quantitative sampling • Air sampling • Polymerase chain reaction
Introduction
M.C. O’Loughlin (*)
Department of Life Sciences, University of Limerick,
Castletroy, Limerick, Ireland
e-mail: maryc.oloughlin@gmail.com
K.D. Turner
Centre for Chromosome Biology, School of Natural
Sciences, National University of Ireland Galway,
Galway, Ireland
K.M. Turner
Manufacturing Sciences and Technology, Pfizer Ireland
Pharmaceuticals, The Pfizer Biotech Campus at Grange
Castle, Clondalkin, Dublin 22, Ireland
In the nineteenth century Louis Pasteur disposed
of the supposition of spontaneous generation by
showing that airborne microscopic organisms
account for biologic growth on previously sterile
media. It has been estimated in recent times that up
to 40% of homes in Northern Europe and Canada
have mould contamination [1]. Various health
effects, such as respiratory symptoms, allergic
rhinitis, asthma, and hypersensitivity pneumonitis,
are associated with mould exposure. Traditional
methods for the isolation and identification of fungal spores can be time-consuming and laborious.
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_29, © Springer Science+Business Media, LLC 2013
337
338
Sampling should be performed by using validated
methods and must be planned so that the smallest amount of sampling and interpretation is
done to meet the information requirements. Air
sampling is widely used in the detection of such
moulds for eventual identification and if possible
enumeration [2–5].
These air techniques are generally categorized as both passive (gravitational) and active
(volumetric). Traditionally, passive air sampling (Settle plate) has been used and is still
commonly in use to determine the types of
microorganisms by exposing a Petri dish containing nutrient rich agar medium to the air
[6, 7]. The method is somewhat criticized for
being considered semiquantitative with a
potential for bias towards microorganisms of
larger spore size. This is primarily due to its
reliance on gravity. However, it does have the
potential to mimic the natural deposition of
airborne spores on the surface of food products
and some regard it as a dependable method to
assess airborne microbial food contamination.
The technique is straightforward to perform
and does not involve additional investment on
specialized equipment.
Active air sampling uses devices that draw a
predetermined volume of air at a particular
speed over a definite period of time for the
assessment of viable airborne microorganisms.
Though both are widely used, there has been
criticism that the methodologies for sampling
and analysis are neither consistent nor definitive
[8]. Quantitation of sampled specimens is by
and large carried out by direct microscopy, culture, or biochemical analysis but new DNAbased methods for fungal detection can now be
used to enumerate the spores of fungi. Airborne
spores can be collected and identified by PCR
allowing identification of the species [9, 10].
However, the sample volume, short collection
period, and artificial air disturbance produced
by the devices may in fact affect the types and
quantities of fungi captured [6]. This chapter
outlines protocols for both passive and active air
sampling in the context of quantitation of fungal
strains from indoor areas.
M.C. O’Loughlin et al.
Materials
All chemicals sourced from Sigma-Aldrich, St.
Louis, MO, USA, unless specified.
1. SAS-super-180 sampler (PBI International,
Milan, Italy).
2. Reuter Centrifugal Air Sampler (Biotest,
Frankfurt, Germany) with high volume pump
(Gast Inc., Benton Harbor, MI, USA).
3. Rotameter (Zefon International, Ocala, FL,
USA).
4. DG18 media composed of glucose, 10 g/L;
peptone, 5 g/L; NaH2PO4, 1 g/L; Mg2SO4,
0.5 g/L; dichloran, 0.002 g/L; agar,15 g/L;
glycerol, 220 g/L; and chloramphenicol,
0.5 g/L.
5. Rose Bengal agar medium, MEA (Malt
extract agar), CYA (Czapaek yeast extract
agar), YES (Yeast extract sucrose agar),
CREA (Creatine sucrose agar), and NO2
(Nitrite sucrose agar) (Difco; Becton
Dickinson, Sparks, MD, USA).
6. 9 cm Petri dish (Fisher Scientific Company,
Pittsburgh, PA).
7. Parafilm-M™ (Fisher Scientific Company,
Pittsburgh, PA).
8. 70% Ethanol.
9. 0.05% Tween 80.
10. 13-mm mixed cellulose ester filter 0.8 mm
pore size (Fisher Scientific Company,
Pittsburgh, PA).
11. Acetone-vaporizing
unit
(Quickfix,
Environmental Monitoring Systems, Cha
rleston, SC).
12. Glycerin jelly (20 g gelatin, 2.4 g phenol
crystals, 60 mL glycerol, and 70 mL water).
13. 0.1% IGEPAL CA-630®.
14. Acid-washed Ballotini beads (8.5 grade,
400–455 mm in diameter) and ball mill (Glen
Creston, Stanmore, UK).
15. 2× Lee and Taylor lysis buffer (100 mM
Tris–HCl pH 7.4, 100 mM EDTA, 6% SDS,
2% b-mercaptoethanol).
16. Phenol:chloroform (1:1).
17. 20 mg/mL Glycogen (Roche diagnostics Ltd.,
Lewes, UK).
29
Quantitative Sampling Methods for the Analysis of Fungi: Air Sampling
18. 6 M ammonium acetate.
19. Isopropanol.
20. TE buffer: 10 mM Tris–HCl pH 7.5 (25°C),
0.1 mM EDTA.
21. RNaseA: 10 mg/mL in TE.
22. Phenol.
23. ABI 7000 Fast Real-time PCR System
(Applied Biosystems, CA, USA).
24. Universal fungal primers NS5 (5¢-AA
C T TA A A G G A AT T G A C G G A A G - 3 ¢ )
and NS6 (5¢-GCATCACAGACCTGTTATTG
CCTC-3¢).
25. SYBR® Premix Ex Taq™ II (×2) and ROX
Reference Dye (×50) (Takara Bio., Shiga,
Japan).
Methods
Active Air Sampling Using a Portable
SAS-Super-180 Sampler
1. Operate the SAS-super-180 sampler at a sampling rate of 180 L air/min.
2. Follow the instruction manual provided by the
producer.
3. Charge the portable battery fully prior to use.
4. Sample 500 L of air from the center of the
room at a height of 1 m above the floor.
5. Use a 9-cm Petri dish containing Dichloran
18% glycerol agar medium (DG 18) [11] to
trap viable fungal particles.
6. Disinfect the sampler with 70% ethanol before
each use.
7. Cover the Petri dish with the lid immediately following sampling and seal with
Parafilm-M™.
8. Incubate the Petri dishes containing DG18 at
25°C for 1 week in the dark and examine every
24 h.
Active Air Sampling Using a Reuter
Centrifugal Air Sampler
1. Operate the Reuter centrifugal air sampler
(RCS) at a flow rate of 40 L air/min [8, 12–14].
339
2. Follow the instruction manual provided by the
producer.
3. Calibrate the high volume pump to 28.3 L/min
using a rotameter.
4. Collect 15 samples of 1–15 min duration in
random order at a height of 1 m above the
floor.
5. Use a 9-cm Petri dish containing Rose Bengal
agar medium supplemented with 100 mg/L
chloramphenicol to trap viable fungal particles.
6. Disinfect the sampler with 70% ethanol before
each use.
7. Cover the Petri dish with the lid immediately
following
sampling
and
seal
with
Parafilm-M™.
8. Incubate the Petri dishes containing Rose
Bengal agar at 25°C for 1 week and examine
every 24 h.
Isolation and Enumeration
of Mycological Samples
1. Count the numbers of fungal colonies on each
Petri dish and then subculture on Petri dishes
with suitable agar media for species
identification.
2. Prepare all media as per manufacturers’
instructions.
3. Plate moulds belonging to Penicillium on the
following media; MEA, CYA, YES, CREA,
and NO2.
4. Plate other moulds and yeasts on MEA and
PDA (Potato dextrose agar) [11].
5. Incubate MEA, CYA, YES, and PDA in the
dark at 25°C and CREA and NO2 at 20°C for
7 days [15].
6. Convert the number of CFU per plate to the
number of CFU/L of air and analyze data
using an appropriate statistical package.
Identification of Mycological Samples
1. Suspend individual colonies in 20 mL 0.05%
Tween® 80 prepared with sterile deionized
water in a test tube [5].
340
2. Vortex for 10 s at 20,000×g.
3. Filter each sample through a 0.8 mm mixed
cellulose ester filter and then place on a glass
slide.
4. Allow slides to dry overnight.
5. Clear the slides using a modified instant acetone-vaporizing unit
6. Mount a 25 × 25-mm cover glass on the slide
using glycerin jelly.
7. Observe the slides and identify the collected
fungal spores at ×400 magnification using a
light microscope.
Extraction of DNA from Mycological
Samples
1. Suspend individual colonies in 0.5 mL 0.1%
IGEPAL CA-630® prepared with sterile
deionized water in a tube.
2. Adjust the spore suspensions were adjusted
to 2–3 × 104 spores/mL.
3. Transfer 0.4 mL of the spore suspension to a
fresh tube and add 0.4 g of acid-washed
Ballotini beads (8.5 grade, 400–455 mm in
diameter).
4. Shake the mixture for 8 min in a ball mill.
5. Add 0.4 mL 2× Lee and Taylor lysis
buffer [16].
6. Vortex the sample and incubate at 65°C for 1 h.
7. Add 0.8 mL phenol:chloroform (1:1), vortex
briefly, and centrifuge at 20,000×g for 15 min
8. Transfer the top aqueous layer to a clean
tube.
9. Add 1 mL glycogen (20 mg/mL), 40 mL 6 M
ammonium acetate, and 600 mL isopropanol
10. Invert tube gently to mix and incubate at
−20°C for 10 min.
11. Centrifuge at 20,000×g for 2 min and remove
the supernatant.
12. Resuspend the pellet in 50 mL TE buffer containing RNase A (10 mg/mL) to the pellet
and incubate at 37°C for 15 min.
13. Add 150 mm TE and then add 200 mL phenol
14. Vortex briefly and centrifuge at 20,000×g for
6 min.
M.C. O’Loughlin et al.
15. Transfer the top aqueous layer to a clean
tube.
16. Add 10 mL 6 M ammonium acetate and
600 mL isopropanol.
17. Invert tube gently to mix and incubate at
−20°C for 10 min.
18. Centrifuge at 20,000×g for 10 min and
remove the supernatant.
19. Add 800 mL ice cold 70% ethanol, centrifuge at 20,000×g for 2 min, and remove the
supernatant.
20. Centrifuge at 20,000×g for 10 s and remove
the remaining liquid.
21. Dry pellet for 20 min in a fume hood and
resuspend in 50 mL TE buffer.
Quantitation of Mycological Samples
Using RT-PCR
1. Use the universal fungal primer pair NS5
(5¢-AACTTAAAGGAATTGACGGAAG-3¢)
and NS6 (5¢-GCATCACAGACCTGTTATT
GCCTC-3¢) to amplify the 310 bp of 18S
rDNA region [17].
2. Thaw the reagents and DNA preparations and
keep on ice until required.
3. Dilute the extracted fungal DNA in sterile
PCR-grade water to several ratios; i.e., 0.5/20,
1/20, 2/20, 4/20, and 8/20.
4. Prepare a 50-mL reaction mixture consisting
of 25 mL of SYBR® Premix Ex Taq™ II (×2),
1 mL of ROX Reference Dye (×50), 2 mL of
each primer (10 mM), and 20 mL of the diluted
DNA extracts.
5. Program the ABI 7000 fast real-time PCR
System, or equivalent system, with the following cycling conditions 95°C for 10 s, 4
cycles of 95°C for 30 s, 60°C for 31 s, and
then 40 cycles of 95°C for 4 s and 60°C for
31 s.
6. Set a threshold level of 0.2 and use the
auto-baseline function on the ABI 7000
software.
7. Carry out quantitative analysis of the sample
using the ABI 7000 software.
29
Quantitative Sampling Methods for the Analysis of Fungi: Air Sampling
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Indoor fungal composition is geographically patterned
and more diverse in temperate zones than in the tropics. Proc Natl Acad Sci U S A 107:13748–13753
Lee SB, Taylor JW (1990) Isolation of DNA from fungal mycelia and single spores. In: Innis MA, Gelfand
DH, Sninsky JJ, White TJ (eds) PCR protocols: a
guide to methods and applications. Academic, San
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Wu Z, Wang X-R, Blomquist G (2002) Evaluation of
PCR primers and PCR conditions for specific detection
of common airborne fungi. J Environ Monit 4:377–382
Transformation of Filamentous
Fungi in Microtiter Plate
30
Bianca Gielesen and Marco van den Berg
Abstract
The polyethylene glycol (PEG) protoplast method for transformation of
filamentous fungi is developed for use in microtiter plates, enabling one to
perform a multitude of transformations in parallel.
Keywords
Polyethylene glycol • PEG protoplast • Genetic transformation
• Filamentous fungi • Microtiter plates
Introduction
Genetic transformation is one of the most commonly used tools in molecular biology. For
filamentous fungi, various methods have been
described, including Agrobacterium-mediated
transformation [1], particle bombardment [2],
and electroporation [3]. Still, the method mostly
used is the classical polyethylene glycol (PEG)mediated protoplast transformation [4]. Although
all methods do work, yielding from a few dozen
to well over 1,000 transformants per microgram
DNA, none of them has been described so far as
a high-throughput method. With the availability
B. Gielesen • M. van den Berg (*)
DSM Biotechnology Center, Alexander Fleminglaan 1,
Delft, Zuid, Holland 2613 AX, The Netherlands
e-mail: marco.berg-van-den@dsm.com
of many fungal genome sequences, such a method
is needed to embark on genome-wide functional
studies.
In this chapter we describe a protocol for protoplast transformation and subsequent regeneration of filamentous fungi in microtiter plate
(MTP). The examples given are obtained with the
ß-lactam antibiotic producing Penicillium chrysogenum, but similar results were obtained for
Aspergillus niger; therefore, the protocol can be
used as a basis for other fungal species as well.
Materials
1. Penicillium YGG medium: per liter: 8.0 g
KC, 16.0 g glucose, 6.7 g Difco yeast nitrogen base, 1.5 g citric acid, 6.0 g K2HPO4,
2.0 g yeast extract, pH 6.2, supplemented
with penicillin and streptomycin (Gibco).
2. KC buffer: per liter: 60 g KCl, 2 g citric acid,
pH 6.2.
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_30, © Springer Science+Business Media, LLC 2013
343
344
3. Glucanex (GLX) stock solution. Dissolve 5 g
GLX in 50 mL KC, cool overnight at 4 °C,
filter sterilize, and store in small portions at
−20 °C.
4. STC buffer: per liter: 145.6 g sorbitol, 2.8 g
CaCl2, 3.0 g Tris, pH 8.0; sterilize in
autoclave.
5. Aurintricarboxylic acid (ATA) stock solution. Dissolve 53 mg ATA in 50 mL and store
in small portions at −20 °C.
6. 20 % PEG solution. Dissolve 200 g PEG4000
in 1 L water; sterilize in autoclave.
7. 30 % PEG solution. Dissolve 300 g PEG4000
in 1 L water; sterilize in autoclave.
8. 10 % Acetamide stock solution in water;
filter sterilize.
9. 1.5 M CsCl stock solution in water; filter
sterilize.
10. Trace element solution: per liter (in g): citric
acid (150), FeSO4·7H2O (15), MgSO4·7H2O
(150), H3BO3·(0.0075), CuSO4·5H2O (0.24),
CoSO4·7H2O (0.375), ZnSO4·7H2O (5),
MnSO4·H2O (2.28), CaCl2·2H2O (0.99); filter
sterilize.
11. Acetamide regeneration medium with
sucrose: (1) 15.0 g agar, 342.0 g sucrose,
5.0 g glucose, 35.0 g lactose, 2.9 g Na2SO4,
5.2 g KH2PO4, 4.8 g K2HPO4 in 900 mL; (2)
set pH 6.5 at before sterilization; (3) after
sterilization in autoclave and cooling to
~50 °C add 10 mL acetamide stock, 10 mL
CsCl stock and 10 mL of a trace element
solution, and fill up to 1 L with sterilized
water.
12. Acetamide regeneration medium with KCl:
(1) 15.0 g agar, 22.4 g KCl, 5.0 g glucose,
35.0 g lactose, 2.9 g Na2SO4, 5.2 g KH2PO4,
4.8 g K2HPO4 in 900 mL; (2) set pH 6.5 at
before sterilization; (3) after sterilization in
autoclave and cooling to ~50 °C add 10 mL
acetamide stock, 10 mL CsCl stock and
10 mL of a trace element solution, and fill up
to 1 L with sterilized water.
13. 96-well MTP, standard 200 mL shallowwell, 200 mL wide-well, and 1 mL deepwell.
14. 24-well deep-well MTP.
15. Filters 14 × 14 mm; sterilize in autoclave.
B. Gielesen and M. van den Berg
Methods
Protoplastation
1. Inoculate 25 mL of YGG with 104–105
spores/mL of P. chrysogenum.
2. Incubate o/n at 25° C and 250 rpm.
3. Dilute the overnight culture 1:30 into 100 mL
fresh YGG and incubate o/n at 25 °C and
250 rpm.
4. Spin down the cells for 5 min at 5,000 rpm.
5. Wash the cell pellet thoroughly with 10 volumes of KC.
6. Resuspend the cells in 15 mL of KC and add
4 mL of GLX.
7. Incubate at 25 °C and 100 rpm.
8. Monitor the protoplast formation under a
light microscope every 30 min and cool on
ice when >90 % mycelium is converted into
protoplasts.
9. Add cold KC up to 50 mL and spin down
protoplasts for 5 min at 1500 rpm.
10. Discard the supernatant and resuspend the
protoplasts in cold KC up to half the volume
of the tube and add another half of cold STC.
Spin down for 5 min at 1,500 rpm.
11. Discard the supernatant. Wash the protoplasts in cold STC. Spin down for 5 min at
1,500 rpm.
12. Discard the supernatant and resuspend the
protoplasts in cold STC.
13. Count the protoplasts in the supernatant and
adjust towards 108 protoplasts/mL. The protoplasts are now ready for direct use or can
be frozen at −80 °C for later use.
Transformation
In order to study the robustness towards varying
types of MTP, transformations were performed in
standard (i.e., shallow-well) 96-well MTP, widewell 96-well MTP, and deep-well 96-well MTP.
Transformants were obtained with all MTP types.
For our experiments, we typically use either the
standard (i.e., shallow-well) 96-well MTP or
wide-well 96-well MTP.
30
Transformation of Filamentous Fungi in Microtiter Plate
345
Fig. 30.1 Comparison of manual versus automated
handling of Penicillium chrysogenum MTP transformation. P. chrysogenum strain DS17690 was transformed
with plasmid pHELY-A1 according to the classical manual method [6] and the MTP method as described in this
chapter. The transformation mixtures were plated out on
acetamide-selective agar in petri dishes. y-axis, number of
transformants per 50 mL of protoplasts (or 2 mg of DNA)
as obtained on standard agar plates; x-axis, different
handling protocols. C control, i.e., the classical glass
tube protocol with manual pipetting and swirling; A1
MTP method with automated pipetting and swirling;
A2 MTP method with automated pipetting and mixing;
M1 MTP method with manual pipetting and swirling; M2
MTP method with manual pipetting and mixing
1. Pipette in each well of a standard 96-well MTP
4 mL of DNA (typically, 0.5–2 mg) or H2O as
control. In our experiments we used as DNA
0.25 mg/mL of the plasmid pHELY-A1 harboring an acetamidase expression cassette [5].
Upon uptake of the DNA, this expression cassette enables transformants to grow on media
with acetamide as the sole nitrogen source.
2. Mix together the protoplasts, the nuclease
inhibitor (ATA) and 20 % PEG in the ratio of
10:1:5.
3. Add 40 mL of the mixture from step 2 to the
DNA-containing wells. This can be done
either manually (pipetting using a repetitive
or a multichannel pipette) or automated (e.g.,
using a Multidrop from Thermo Electron
Corporation). Large differences between the
number of transformants obtained with either
the manual or the automated method
(Fig. 30.1) were not detected.
4. If step 3 is done manually, the mixture is
carefully mixed, either via swirling, or via
pipetting up-and-down.
5. Subsequently, the MTP is incubated on ice
for 30 min.
6. Add to each well 190 mL 30 % PEG. As in step
4, this can be done manually or automated. No
additional mixing is required at this step.
7. Incubate for 15 min at 25 °C.
8. Add to each well 700 mL STC. No additional
mixing is required at this step.
9. Spin down the protoplasts via centrifugation
for 5 min at 1,500 rpm. Discard the supernatant and try not to disturb the pellet.
10. Resuspend the pellet by adding 100 mL STC
per well. The mixtures are now ready for
plating out on selective media and regeneration of the protoplasts.
Regeneration
In order to study the robustness towards varying
conditions for regeneration, we looked at four
parameters: (1) the type of osmotic stabilizer, i.e.,
sucrose or KCl; (2) the concentration of KCl as
osmotic stabilizer; (3) the MTP type, i.e., 24- and
96-well (both shallow- and deep-well); (4) control of water surface tension/humidity by plating
on a filter or not. As illustrated by Figs. 30.2 and
30.3, all variations do lead to transformants, with
slight differences in the numbers obtained.
Typically 24-well deep-well plates, filled with
3 mL of medium (acetamide selective medium
with 0.3 M of KCl) and sterile filters placed on
top were used.
346
B. Gielesen and M. van den Berg
Fig. 30.2 Comparison of osmostabilizers during MTP
transformation of Penicillium chrysogenum. P. chrysogenum strain DS17690 was transformed with plasmid
pHELY-A1 according to MTP transformation method as
described in this chapter. (a) Osmostabilizing either by
0.6 M KCl or 1 M sucrose does not lead to a drastic difference, either with (gray bars) or without (white bars)
filters. (b) Increasing the concentration of KCl does have
a clear effect on the number of transformants, whereas
plating on filters (gray bars) leads to, on average, 50 %
more transformants as compared to plating without filters
(white bars). y-axis, average number of transformants per
well, with standard deviation indicated (n = 3-5); x-axis,
osmostabilizer concentration
1. The protoplast suspensions are transferred at
different dilutions to selective acetamide MTP,
either manually (using a multichannel pipette)
or automated (any pipetting station with sterility control will do)
2. Incubate the MTP for 5–7 days at 25 °C. For
improved humidity control one might leave the
lids off the MTP in a flow cabinet for 1 h before
closing and transfer to incubator. Or, alternatively, after the first day in the incubator, the
plates are put in a box inside the incubator.
3. Transfer obtained transformants to a second
selective plate for purification.
4. If needed, apply further characterization techniques to selected transformants in order to
verify the altered genotype and the subsequent
altered phenotype. Typically this method
results in several dozens of transformants per
mg DNA.
Notes
1. Instead of the YGG medium, the standard
yeast medium (YEPD, per liter: yeast extract
10 g, pepton 10 g, glucose 20 g) can be used,
or any other medium most suitable to the fungus of choice.
2. It is advisable to determine the best effective
MTP format, regenerative medium, and osmostabilizer (concentration) for each specific
filamentous fungal species.
3. When selection markers other than acetamidase are used (e.g., phleomycin, nourseotricin,
or hygromycin resistance), the regeneration
medium needs to be adapted accordingly.
4. The MTP transformation method can be very
well combined with the so-called NHEJ
mutants of filamentous fungi, which have a
significant improved frequency of homologous
30
Transformation of Filamentous Fungi in Microtiter Plate
347
Fig. 30.3 Penicillium chrysogenum transformants regenerated in
MTP. P. chrysogenum strain
DS17690 was transformed with
plasmid pHELY-A1 according to
the MTP transformation method
as described in this chapter.
(a) Regeneration in standard
shallow-well 96-well MTP on
acetamide medium with 1 M
sucrose. (b) Regeneration in
deep-well 24-well MTP on
acetamide medium with 0.3 M
KCl on a filter
recombination (example as reported in reference [6]).
5. The transfer of transformants to the second
selective plates preferably is done manually
with a toothpick and re-streaking to single
spore-derived colonies.
References
1. de Groot MJ, Bundock P, Hooykaas PJ, Beijersbergen AG
(1998) Agrobacterium tumefaciens-mediated transformation of filamentous fungi. Nat Biotechnol 16:839–842
2. Aboul-Soud MA, Yun BW, Harrier LA, Loake GJ
(2004) Transformation of Fusarium oxysporum by particle bombardment and characterisation of the resulting
3.
4.
5.
6.
transformants expressing a GFP transgene.
Mycopathologia 158:475–482
Chakraborty BN, Patterson NA, Kapoor M (1991) An
electroporation-based system for high-efficiency transformation of germinated conidia of filamentous fungi.
Can J Microbiol 37:858–863
Tilburn J, Scazzocchio C, Taylor GG, ZabickyZissman JH, Lockington RA, Davies RW (1983)
Transformation by integration in Aspergillus nidulans.
Gene 26:205–221
van den Berg MA, Bovenberg RAL, Raamsdonk LML,
Sutherland JD, de Vroom E, Vollinga RCR (2007)
Cephem compound. PCT/NL2004/000367
Snoek IS, van der Krogt ZA, Touw H, Kerkman R,
Pronk JT, Bovenberg RA et al (2009) Construction of a
hdfA Penicillium chrysogenum strain impaired in nonhomologous end-joining and analysis of its potential
for functional analysis studies. Fungal Genet Biol
46:418–426
Molecular Fingerprinting of Fungal
Communities in Soil
31
Roberto A. Geremia and Lucie Zinger
Abstract
Study of fungal spatio-temporal dynamics requires analyzing a high
number of samples with robust methods in a reasonable time. Fingerprinting
DNA-based methods are better adapted than classical cultural methods for
such purposes. Although molecular fingerprinting does not allow
identification of phylotypes, they provide a snapshot of fungal communities, allowing comparison of a large number of samples.
Keywords
Fungal communities • Phylogenetic structure • Molecular fingerprinting
• Molecular signature • Spatio-temporal dynamics • Soil
Introduction
The fungal biomass represents a large amount of
soil biota, and sometimes it is the major biotic
community of soil [1]. Soil fungal communities
hold a large panel of functional properties that
place them as the first step of the soil food web.
Indeed, the fungal extracellular enzymes are
involved in degradation of vegetal organic
matter [2]. They also influence plant development
and productivity by mutualistic nutrient relocation
through mycorrhiza [2–5]. However, little is
known about the factors that influence the dynamics and functional implications of fungal commu-
R.A. Geremia (*) • L. Zinger
Laboratoire d’Ecologie Alpine, CNRS/UJF,
Université Joseph Fourier, BP 53 Bat D Biologie,
Grenoble 38041, France
e-mail: roberto.geremia@ujf-grenoble.fr
nities as well as the systematic study of different
habitats. Indeed, the identification of situations
that lead to changes in the genetic structure of fungal communities would allow better understanding
of the ecological role of fungi in different habitats.
For instance, studies in alpine tundra habitats show
the prominent role of snow as well as plant cover
in the distribution and dynamics of fungal communities [6–8]. Molecular tools for characterization of fungal communities play a prominent role
in this kind of study.
The phylogenetic structure of fungal communities is most commonly achieved using molecular methods. Actually, cultural methods are
time-consuming and only allow one to cultivate
less than 20% of the fungal strains [9]. The simplest molecular methods to assess the phylogenetic structure of fungal communities are the
so-called molecular fingerprinting or molecular
signature. Although these methods do not provide
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_31, © Springer Science+Business Media, LLC 2013
349
350
taxonomic identification, they display a
snapshot of fungal diversity. The emergence or
loss of fungal phylotypes would be detected as
changes in the electrophoretic pattern of the
community. Total soil DNA is used as a template for the PCR-amplification of a marker
gene, the most common marker being the internal transcribed spacer 1 (ITS1) [10]. The PCR
amplicons, actually a mixture of different fungal phylotypes, are separated through three
kinds of methods relying on electrophoresis.
First, the marker gene is separated according
its length (size polymorphism); the most common are fragment length analysis (FLA) and
automated ribosomal intergenic spacer analysis (ARISA). Second, the marker gene is separated according to its conformation (sequence
polymorphism) either by denaturant (temperature gradient gel electrophoresis, TGGE; denaturing gradient gel electrophoresis, DGGE), or
native electrophoresis (single-stranded conformation polymorphism, SSCP). Finally, the
third technique relies on both previous methods: PCR amplicons are digested according to
their sequence with restriction enzyme, and
then separated by their length (restriction fragment length polymorphism (RFLP) and terminal-RFLP (T-RFLP)).
The robustness of final results requires a high
sampling density and PCR replicates, which
increases the number of samples to be processed.
To cope with such constraints, a high-throughput
and robust strategy was set up in our laboratory.
We first adapted a protocol of the commercial
Power-Soil kit for extraction. Second, to reduce
the experimental steps, we chose a straightforward method like SSCP. Finally, we used capillary electrophoresis (CE), which provides the
possibility of analyzing a large number of samples in a few hours. Although CE has already
been coupled to T-RFLP for soil fungal community studies [11, 12], the use of CE-SSCP remains
limited. Additionally, and owing to different
fluorophores, CE allows the analysis of at least
three different primer pairs. We have used
CE-SSCP to study fungal, bacterial, and
chrenoarcheal communities. Here, we present a
protocol allowing the study of fungal and bacterial communities.
R.A. Geremia and L. Zinger
Materials
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
25.
26.
27.
28.
29.
Manual drill.
Plastic gloves.
Sieves <5 and <2 mm.
Disposable polypropylene tubes: 50 mL
conical.
Ultra-high-quality (UHQ) water.
PowerSoil® DNA Isolation Kit catalog no.
12888-100 (Mobio Lab. Inc, Carlsbad, CA).
FastPrep®-24 Instrument (MP Biomedicals,
Inc., Illkirch, France).
Vortexer.
Centrifuge.
PCR hood.
Disposable polypropylene microcentrifuge
tubes: 1.5 mL conical.
Spectrophotomer UV.
PCR machine.
Automatic micropipettes (i.e., Pipetman).
Filter tips, 0.01–1 mL.
96-well PCR Plate, rigid semi-skirted, ABI.
Fungal primers (12): ITS5 (5¢-GGAAGTAAAAGTCGTAACAACG-3¢), ITS2-FAM
(5¢-fluorescein phosphoramidite GCTGCGTTCTTCATCGATGC-3¢).
AmpliTaqGold
(Applied
Biosystems,
Stockholm, Sweden).
Agarose.
Ethidium bromide: 0.5 mg/mL stock.
Gel loading mixture: 40% (w/v) sucrose,
0.1 M EDTA, 0.15 mg/mL bromophenol blue.
Horizontal electrophoresis equipment.
UV transilluminator and camera suitable for
photographing agarose gels, e.g., Syngene
Gene Genius Bioimaging system.
TBE buffer.
Mark XIV gene ladder.
Hi-Di™ Formamide (Applied Biosystems).
NaOH 0.3 M.
HD-Rox-400 markers (Applied Biosystems).
Multi-well plate for sequencer.
Methods
The methods described as follows were designed
to obtain a fungal molecular profile specific to a
31
Molecular Fingerprinting of Fungal Communities in Soil
given habitat. It allows one to withdraw a soil
sample, extract its DNA, amplify the ITS1 from
this DNA, obtain the SCCP profile, and generate
distance trees. Depending on the content and
quality of soil organic matter, relative amount of
fungal DNA, some modifications should be introduced at the PCR level. Our suggestions are indicated in the Notes.
351
agarose gel; DNA is considered suitable when
only a high molecular weight (MW) with no or
low smear. DNA concentration and purity are
estimated by absorbance at 260/280 nm. DNA
could be stored a few days at 4°C, but for longer
time it should be stored at −20°C.
Amplification of ITS1 Fragments by PCR
Soil Sampling
The DNA extraction requires 0.25 g of soil.
However, in order to minimize the bias due to
spatial heterogeneity in fungal distribution, larger
samples from different points of the sampling site
should be obtained. As previously suggested [12],
at least five samples per habitat replicate should
be obtained. Usually, we obtain 50 g of soil per
sampling point.
1. Select the points to be sampled based on the
principles of the experiment.
2. Drill the first 15 cm of the soil; wearing gloves
place them in a plastic bag or directly sieve
them (see Note 1).
3. To perform the sieving wear gloves and keep
the sample out of the refridgerator for as little
time as possible.
4. Manually desegregate the soil.
5. Sieve the soil first at 5 mm.
6. Recover the material passing through the sieve.
7. Sieve the soil at 2 mm.
8. Recover the material passing through the
sieve; transfer to a polypropylene 50-mL tube.
9. Store the tube at −20°C.
DNA Extraction
This step is performed in a dedicated laboratory,
to prevent contamination with DNA of other
organisms or PCR products. The protocol used is
the one provided by the manufacturer for individual tubes (available at www.mobio.com/images/
custom/file/protocol/12888.pdf) (see Note 2),
with only one modification in step 5: instead of the
flat vortex, we used the Fastprep 24 instrument at
maximum power two times 20 s. DNA integrity
is estimated by electrophoresis in a 1.6% W/V
The protocol below uses the primers ITS5 and
ITS2, which work nicely in our laboratory [13–
15]. This protocol was also used to obtain amplicons for pyrosequencing, which allowed to test
their specificity; only 1.3% of the obtained
sequences were assigned to Virideplantae, 0.01%
to Metazoa, and 0.02% to Alveolata, Rhizaria,
and Rhodophyta [16]. Fungal ITS1 was amplified
with the primers ITS5 and ITS2-FAM labeled
[17]; however, new primers have recently been
proposed that are even more specific [18], but we
do not have experience on SSCP.
1. The setup of the PCR reaction is performed in
a dedicated hood, located in the DNA extraction laboratory. Wear gloves through all the
preparation.
2. For each sample, make three independent
PCR reactions and prepare at least three negative controls.
3. The PCR reactions (25 mL) consisted of
2.5 mM of MgCl2, 1 U of AmpliTaq GoldTM
buffer, 20 g/L of bovine serum albumin
(BSA), 0.1 mM of each dNTP, 0.26 mM
of each primer, 0.2 U of AmpliTaqGold
DNA polymerase (Applied Biosystems), and
around 10 ng of DNA template in 1 mL.
4. Thaw the reagents and DNA preparations.
Reactions are carried out in 25 mL. Prepare a
reaction mixture sufficient for the number of
samples to be tested.
5. To prepare the reaction mixture, use a set of
dedicated micropipettes.
6. Calculate the amount of the reaction mixture
to be prepared using 24 mL/sample. Each
96 mL (for 4 samples) of the master mix, contains 62.4 mL of UHQ water, 10 mL of 25 mM
MgCl2, 4 mL of a mixture containing 2.5 mM
of each dNTP, 4 mL of each 5 mM ITS5 and
5 mM ITS2-FAM, 0.8 mL of L 20 mg/mL
352
7.
8.
9.
10.
11.
12.
R.A. Geremia and L. Zinger
BSA, and 0.8 mL of AmpliTaq Gold polymerase (1 U/mL) (see Note 2.) Gently vortex,
centrifuge, and add 24 mL aliquots into the
wells of a suitable multi-well PCR plate.
With a micropipette equipped with filter tips,
add 1 mL of soil DNA (10–20 mg/mL) to the
PCR reaction well; mix briefly with the pipet
tip. To the negative controls add 1 mL of
UHQ water. Cover the plate with a suitable
adhesive film and centrifuge. Carry out the
PCR reaction as follows: an initial step at
95°C (10 min), followed by 30 cycles at
95°C (30 s), 56°C (15 s), and 72°C (15 s),
and final step at 72°C (7 min).
The PCR products were checked on a 1.6%
agarose gel. A positive reaction shows one or
several bands between 150 and 500 bp (see
Note 3).
The samples where amplification was successful are further submitted to CE-SSCP.
Calculate the amount of denaturing mixture considering 10 mL of Hi-Di™
Formamide (Applied Biosystems) (see
Note 4), 0.5 mL of 0.3 M NaOH, and 0.2 mL
of internal DNA molecular weight (MW)
standard Genescan-400HD ROX (Applied
Biosystem) per sample. Aliquot 10 mL of
the mixture in an ABI sample plate, add
1 mL of the sample, and place the plate
cover (see Note 5).
Denature for 3 min at 95°C in a PCR machine.
Cool on ice to prevent re-annealing of double
strands. Keep at 4°C until running.
Perform the SSCP run on an ABI Prism 3130xl
Genetic Analyzer™ (Applied Biosystem)
using a capillary 36 cm in length. The nondenaturing polymer consists of 5% Genscan
polymer, 10% glycerol, and 16Tris–borate–
EDTA (TBE) (Euromedex, France). Running
buffer consists of 10% glycerol and 1× TBE
Injection time and voltage should be set to
22 s and 6 kV. Electrophoreses are performed
at 32°C; data were collected for 25 min.
Data Analysis
In order to analyze the data by statistical
methods, the electrophoregram should be recov-
ered as a table. We have developed software to
produce this kind of table, standardizing with
the molecular weight markers (Supplementary
Material 1, see Note 6). The fluorescence units
might vary a lot according to PCR or electrophoresis’ capillaries. To make these profiles
suitable to use for abundance studies, it is necessary to normalize peak height. A Scilab script
is available under request (see Note 6) that also
selects the size of fragments (150–400 bp) to
include in the final analysis. The resulting tables
are then suitable to construct distance-based
trees or multivariate analysis. It worth mentioning that peaks often shows “shoulder” precluding the use of SSCP data for study of
alpha-diversity. On the other hand, these data
are suitable for beta-diversity analysis. We currently use the R packages ade4 or vegan for
these purposes.
Notes
1. It is highly recommended to sieve the soil at
the sampling site. However, it is possible that
field conditions do not allow one to do so, in
which case the soils should be stored at 4°C
and sieved as soon as possible. Sieving 96
samples by three persons can be achieved in 1
day of work.
2. A kit based on multi-well plates is also available at MoBio. However, the possibility of
cross contamination between wells makes the
use of individual tubes preferable.
3. In certain cases, no PCR products are found.
This can be due to the presence of inhibitors
or a tiny amount of fungal DNA. In both
cases the best solution to get amplicons is to
increase the number of cycles to 32–35. In
the second case, up to 50 mg of DNA can be
added.
4. Work under a chemical hood.
5. It is possible that the fluorescence in the region
of interest is saturated, in which case a 1/10 or
1/50 dilution should be done.
6. The software “Lucieanalyzer” to read the *.fsa
or *.ab1 files from the ABI Prism Genetic
Analyzer™ is available under request to
roberto.geremia@ujf-grenoble.fr.
31
Molecular Fingerprinting of Fungal Communities in Soil
SSCP Data Analysis
353
354
R.A. Geremia and L. Zinger
31
Molecular Fingerprinting of Fungal Communities in Soil
355
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Development of Microsatellite
Markers from Fungal DNA Based
on Shotgun Pyrosequencing
32
Shaobin Zhong
Abstract
Traditional methods for the isolation and identification of fungal microsatellite
markers mostly rely on construction of microsatellite-enriched DNA libraries and Sanger sequencing of clones from these libraries. These methods are
time-consuming, labor-intensive, and relatively expensive. In this chapter, a
quick and cost-effective approach is described for the discovery of microsatellites in fungi based on direct shotgun pyrosequencing. With this approach,
high molecular weight DNA is extracted from the fungus of interest and
subjected to 454 genome sequencing. The sequence reads are assembled
into contigs or unique sequences in a fasta format. Then, free softwares
such as MSATCOMMANDER and QDD are used for microsatellite search
and primer design. The designed primer pairs are tested for PCR
amplification and polymorphism using DNA samples from a diverse
collection of fungal isolates.
Keywords
Microsatellite • Single sequence repeat • Fungi • DNA • Shotgun pyrosequencing • 454 Genome Sequencer FLX
Introduction
Microsatellites or single sequence repeats (SSRs)
are short, tandemly repeated motifs of 1–6 bases,
which are found in genomes of all eukaryotes,
including plants, animals, and fungi [1, 2]. Due to
their high level of polymorphism, co-dominance,
S. Zhong (*)
Department of Plant Pathology, North Dakota State
University, Walster Hall 306, Fargo, ND 58102, USA
e-mail: shaobin.zhong@ndsu.edu
ease to score, and reproducibility, SSR markers
have been widely used in many fields of biology,
such as genome mapping and population genetics
[3, 4]. However, development of SSR markers is
still a big challenge, especially for the non-model
organisms, which have no or limited genomic
sequences available and/or relatively low frequency of microsatellites in their genomes [5, 6].
Traditional methods for the development of SSR
markers mostly rely on construction of SSRenriched libraries and then sequencing of the
clones from these libraries by the Sanger sequencing technology [2]. Although this approach has
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_32, © Springer Science+Business Media, LLC 2013
357
358
been used for developing SSR markers in a large
number of organisms, the labor-intensive and
time-consuming procedure as well as the relatively high Sanger sequencing costs prevent it
from becoming a high-throughput method for
large-scale SSR discovery. With the development
of next-generation sequencing (NGS) technology
[7], rapid and cost-effective discovery of microsatellite markers in species without prior genome
sequence information has become feasible [6, 8–
10]. In general, two approaches have been used
for microsatellite discovery based on NGS. One
approach is to generate random genomic DNA
sequences from the organism of interest by direct
shotgun pyrosequencing and then search for those
sequences or contigs containing microsatellites
[8–12]. The other approach uses the shotgun
pyrosequencing technology to sequence microsatellites-enriched DNA libraries [13–15]. For
those organisms with low microsatellite abundance or with large genome size, microsatellite
enrichment increases the amount of target microsatellites in the sequences generated [13–15], but
this can lead to systematic biases in the type of
microsatellite detected [6]. Due to the decreasing
costs of shotgun pyrosequencing and the relatively small sizes of fungal genomes, the development of SSR markers based on the direct
shotgun pyrosequencing approach has become
one of the most favorable choices. In this chapter,
I will describe procedures for the development
and characterization of SSR markers from
sequence reads generated by 454 Genome
Sequencer FLX.
Materials
Equipment and Consumables
1. Computers (see Note 1).
2. LI-COR 4300 DNA Analyzer (Li-Cor Inc.,
Lincoln, NE, USA).
3. Bench-top microcentrifuge, e.g., Eppendorf
5415D.
4. Refrigerated centrifuge, e.g., Sorvall RC-5
Refrigerated Centrifuge.
S. Zhong
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
Vortexer.
Lyophilizer.
Mortars and pestles.
Rubber policeman.
Flasks (1,000 mL).
Beakers (250 mL).
Miracloth (CalBiochem, EMD Chemicals,
Inc., San Diego, CA, USA).
Cheesecloth.
Environmental incubator shaker.
pH meter.
Horizontal electrophoresis equipment.
UV transilluminator and camera suitable for
photographing agarose gels.
Water bath.
Petri plates.
Microcentrifuge tubes.
Centrifuge tubes (50 mL).
PCR tubes.
Thermal cycler, e.g., Master 100.
Compound microscope.
Hemocytometer.
Media and Reagents
1. Sterile distilled water.
2. V8-PDA (Add 150 mL V8 juice, 10 g PDA,
3 g CaCO3, 10 g Agar, and 850 mL H2O to
make a total volume of 1,000 mL).
3. Potato dextrose broth (PDB).
4. DNA extraction buffer (50 mM Tris–HCl pH
8.0, 150 mM EDTA, 1% Sarkosyl [n-lauroyl
sarcosine], and 300 mg/mL proteinase K [add
fresh]).
5. Electrophoresis-grade agarose.
6. TE (10 mM Tris–HCl pH 8.0, 1 mM
EDTA).
7. Proteinase K.
8. Isopropanol.
9. Ethanol (70%).
10. Phenol solution equilibrated with 10 mM
Tris–HCl pH 8, 1 mM EDTA (EMD
Chemicals, Inc., San Diego, CA, USA).
11. Phenol:chloroform:isoamyl alcohol (25:24:1)
(USB Corporation, Cleveland, OH, USA).
12. Chloroform:isoamyl alcohol (24:1).
32
Development of Microsatellite Markers from Fungal DNA…
13. RNaseA (10 mg/mL in TE, heat treated by
standing in a boiling water bath for 15 min).
14. 20% PEG8000/2.5 M NaCl (w/v).
15. 3 M NaOAc.
16. Lamda DNA.
17. dNTPs [a mixture of dATP, dCTP, dGTP, and
dTTP, 10 mM each from Promega, Madison,
WI, USA, stored at −20°C].
18. Thermostable DNA polymerase (e.g., Taq,
and reaction buffer supplied by manufacturer) (see Note 2).
19. Oligonucleotide primers (custom-made from
Eurofins MWG Operon, Huntsville, Alabama,
USA, resuspended to a concentration of
100 mM using sterile distilled water and
stored at −20°C).
20. M13 primer labeled by IRD700 or IRD800 at
the 5¢ end (custom-made from Eurofins
MWG Operon, Huntsville, Alabama, USA,
resuspended to a concentration of 100 mM
using sterile distilled water and stored at
−20°C).
21. TBE buffer (50 mM Tris, 50 mM boric acid,
1 mM EDTA).
22. Ethidium bromide (0.5 mg/mL in stock).
23. Gel loading mixture (40% [w/v] sucrose,
0.1 M EDTA, 0.15 mg/mL bromophenol
blue).
24. RapidGel-XL-6% Liquid Acrylamide (6%
modified acrylamide, 7 M urea, 89 mM
Tris, 89 mM boric acid, and 2 mM EDTA.
USB Corporation, Cleveland, OH, USA)
(see Note 3).
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
Methods
Extraction of High Molecular Weight
DNA from Fungi for Shotgun
Pyrosequencing
See Note 4.
1. Grow the fungus on V8-PDA plates for 7–10
days (see Note 5).
2. Harvest the spores by adding 10 mL of
ddH2O to each plate, scraping the agar surface using a rubber policeman, and pouring
15.
16.
17.
359
the spore suspension through two layers of
cheesecloth into a 250-mL beaker.
Measure the spore concentration using a
hemocytometer.
Inoculate 106–107 spores (see Note 6) into
250 mL of liquid medium (PDB) in a 1-L
flask. Incubate 1–2 days with shaking (150–
200 rpm at RT).
Harvest the mycelia by pouring the fungal
culture through one layer of miracloth into a
1-L beaker.
Collect the mycelia mat retained on the miracloth and place in a 50 mL Falcon tube, and
store at −20°C overnight or until needed.
Lyophilize for 25 h or until the mycelia is dry
and brittle.
Grind the lyophilized mycelia in a mortar
with liquid nitrogen and a pestle to a fine
powder.
Suspend the mycelial powder in disposable
50-mL polypropylene tubes each containing
20 mL DNA extraction buffer. Approximately
1.6 g dry weight is loaded to each tube.
Vortex vigorously for 30–60 s.
Add an equal volume of Tris-saturated phenol
and gently mix well. Centrifuge for 10 min, at
5 K rpm and at 4°C in a rotor. Transfer the
upper (aqueous) phase to a clean tube.
Add an equal volume of 25:24:1 phenol:chloroform:isoamyl alcohol. Centrifuge for 10 min,
at 5 K rpm and at 4°C in a rotor. Transfer the
upper (aqueous) phase to a clean tube.
Repeat step 10 one more time.
Repeat step 11 one more time.
Add 1/10 volume of 3 M NaOAc to the aqueous solution and add two volumes of cold
absolute ethanol slowly into the aqueous
solution to precipitate the DNA. Invert to
mix and leave on ice or−20°C for >10 min.
Spool out the floating DNA from the aqueous
solution with a clean glass rod (or 1-mL
pipette tip) and transfer to a 1.5-mL tube,
wash with 70% ethanol twice, and dry the
DNA for 5–10 min or until dry (see Note 7).
Dissolve the DNA in 200 mL TE (see Note 8).
Add 2 mL of RNase A to each tube and incubate at 37°C for 1 h.
360
18. Run l.5 mL on 0.6% gel to check if the RNA
is still present.
19. Extract the DNA once with 25:24:1
phenol:chloroform:isoamyl alcohol and once
with 24:1 chloroform:isoamyl alcohol.
20. Add 1/10 volume of 3 M NaOAc to the
aqueous solution and add two volumes of
cold absolute ethanol slowly into the aqueous solution to precipitate the DNA. Invert
to mix and leave on ice or−20°C for
>10 min.
21. Spool out the floating DNA from the aqueous
solution with a clean glass rod (or 1-mL
pipette tip) and transfer to a 1.5-mL tube,
wash with 70% ethanol twice, and dry the
DNA for 5–10 min or until dry (see Note 7).
22. Dissolve the DNA in 200 mL TE (see Note 8).
23. Combine two tubes of the DNA solution in
one 2-mL tube. Bring the volume to 1.2 mL
with H2O.
24. Add 800 mL of 20% (w/v) PEG 8000/2.5 M
NaCl. Mix well and incubate on ice for
60 min to precipitate DNA.
25. Spool the DNA out and wash with 70% ethanol (see Note 9). Drain and dry the DNA
pellet.
26. Resuspend the DNA pellet in TE and leave it
in a water bath at 60°C for 10–20 min to dissolve the DNA completely (see Note 10).
27. Check the DNA quality and quantity by electrophoresis using lamda DNA as standards.
Shotgun Pyrosequencing
1. Send the high molecular weight DNA samples
to a DNA facility for library construction and
sequencing using a 454 Genome Sequencer
FLX (see Note 11).
Microsatellite Discovery and Primer
Design
1. Assemble the DNA sequences into contigs and
single reads using the Newbler (see Note 12).
2. Run the software MSATCOMMANDER [16]
to search for all microsatellite loci in the contigs and single reads (see Note 13).
S. Zhong
3. Design primers from the flanking regions
of each microsatellite locus using Primer31
(see Note 14) with the default settings except
for length of expected PCR product set
between 100 and 400 bp, optimal primer pair
annealing temperature of 60°C (range
58–65°C), 50% GC content (range 40–60%),
and optimal primer length of 24 bp (range
21–30 bp).
4. Add an M13 tag (5-CACGACGTTGTAAAACGAC) to the 5¢ end of each forward primer
during primer synthesis so that the fluorescentlabeled M13 primer can be incorporated in
PCR to generate fluorescent-labeled amplicons to be detected with an LI-COR 4300
DNA Analyzer (see Note 15).
Screening of Primer Pairs by PCR
Amplification
1. The designed primers should be tested for
PCR amplification using the DNA sample
extracted from the fungal strain originally
used for the shotgun pyrosequencing.
2. Each PCR amplification contains 1 × PCR
buffer (10 mm of Tris–HCl, 50 mm of KCl),
200 mm of dCTP, dGTP, dTTP, and dATP,
1.5 mm MgCl2, 1 pmol M13 primer labeled
by IRD700 or IRD800 at the 5¢ end, 0.5
pmol 5¢-tagged forward primer, 0.5 pmol
reverse primer, 1 U of Taq polymerase, and
20 ng of genomic DNA in a final volume of
10 ml.
3. PCR is performed in an MJ Research PTC-100
thermal cycler with the following profile: 95°C
for 2 min, 3 cycles at 95°C for 30 s, 56°C for
30 s, 72°C for 60 s, 25 cycles at 94°C for 30 s,
52°C for 30 s, and 72°C for 45 s, and 1 cycle at
72°C for 5 min followed by a 4°C holding step.
4. The PCR products are diluted 10- to 20-fold and
analyzed on an LI-COR 4300 DNA Analyzer
using a 6% polyacrylamide gel (see Note 3).
5. Primer pairs that give clear amplicons are
further used for polymorphism evaluation (see
Note 16).
1
http://frodo.wi.mit.edu/primer3/
32
Development of Microsatellite Markers from Fungal DNA…
Analysis of Polymorphism
of Microsatellite Markers
1. The polymorphism for each microsatellite
marker can be evaluated by PCR using DNA
samples isolated from at least 40 isolates for
haploid fungi (Ascomycota) or at least 20
isolates for diploid or dikaryotic fungi
(Basidiomycota).
2. Each PCR amplification contains 1 × PCR
buffer (10 mm of Tris–HCl, 50 mm of KCl),
200 mm of dCTP, dGTP, dTTP, and dATP,
1.5 mm MgCl2, 1 pmol M13 primer labeled
by IRD700 or IRD800 at the 5¢ end, 0.5
pmol 5¢-tagged forward primer, 0.5 pmol
reverse primer, 1 U of Taq polymerase, and
20 ng of genomic DNA in a final volume of
10 ml.
3. PCR is performed in an MJ Research PTC100 thermal cycler with the following profile:
95°C for 2 min, 3 cycles at 95°C for 30 s, 56°C
for 30 s, 72°C for 60 s, 25 cycles at 94°C for
30 s, 52°C for 30 s, and 72°C for 45 s, and 1
cycle at 72°C for 5 min followed by a 4°C
holding step.
4. The PCR products are diluted 10- to 20-fold
and analyzed on an LI-COR 4300 DNA
sequence using a 6% polyacrylamide gel (see
Note 3).
5. Primers that show polymorphism among the
isolates are chosen for future applications.
Notes
1. A Windows PC that has XP SP2 or Window
7 as its operating system is needed for running MSATCOMMANDER. A Mac computer with OSX 10.6 and greater also works
for MSATCOMMANDER.
2. We typically use Taq from New England
Biolabs (Ipswich, MA, USA), which is supplied with a 10 × reaction buffer containing
20 mM MgSO4. All of these reagents are
stored at −20°C.
3. To make a 6% polyacrylamide gel, add
600 mL 10% ammonium per sulfate and
60 mL TEMED to 100 mL of the
RapidGel-XL-6% Liquid Acrylamide.
361
4. The protocol described here is modified based
on the protocol provided by Drs. Dongliang
Wu and Gillian Turgeon at Department of
Plant Pathology and Plant-Microbe Biology,
Cornell University. It has been used to extract
high molecular weight DNA from Cochliobolus
species for 454 sequencing. Other protocols
may be suitable for isolation of high molecular
weight DNA from fungi for shotgun
pyrosequencing.
5. We use V8-PDA for culturing Cochliobolus
sativus. Other fungi may need a different
medium for spore production.
6. For fungi that do not produce spores, mycelia
can be collected from the agar plates and
blended with a blender before they are used
to inoculate the liquid medium.
7. The DNA pellet should not be too dry.
Otherwise, it is hard to dissolve.
8. Incubating the tube in a water bath at 65°C
can help dissolve the DNA pellet. Check frequently and do not incubate longer than
30 min.
9. The DNA can be recovered by centrifugation
at 10 K rpm and 4°C for 20 min in a
microcentrifuge.
10. If the DNA is still not dissolved, keep the
tube at 4°C for overnight to dissolve the DNA
pellet completely.
11. The 454 GS FLX Titanium system can generate more than ~400 million nucleotides (bases)
per run with average read length of 400 bp. For
a fungal genome with size of 40 million bases,
a full 454 run can generate sequences with 10×
genome coverage, which is enough to develop
hundreds or even thousands of microsatellite
markers. We identified hundreds of microsatellite loci from C. sativus sequences generated
by a ½ 454 run at the Advanced Studies in
Genomics, Proteomics and Bioinformatics
(ASGPB) of University of Hawaii. The
sequencing costs were only $5,250.2
12. Newbler is a software package for de novo
DNA sequence assembly. It is designed
specifically for assembling sequence data
generated by the 454 GS-series of
pyrosequencing platforms.
2
http://asgpb.mhpcc.hawaii.edu/sequence/
362
13. We use MSATCOMMANDER [16] because
it is a free and user-friendly software for
finding microsatellites and for designing
primers. There are other free softwares such
as QDD [17] that can be used for the same
purpose.
14. MSATCOMMANDER has a function for
primer design and for automatically adding tags to the primers, but it only designs
the primer pairs for the first 196 microsatellite loci.
15. If an LI-COR DNA analyzer is not available,
other methods such as vertical polyacrylamide gel electrophoresis with silver staining
[18] can also be used to detect the microsatellite markers.
16. For haploid fungi, primer pairs that generate one allele or amplicon are usually chosen for further characterization. For diploid
or dikaryotic fungi, primer pairs that generate one or two alleles can be chosen for
further evaluation. It depends on if the
fungal isolate used is homozygous or
heterozygous.
Acknowledgements The author thanks Drs. Dongliang
Wu and Gillian Turgeon for sharing the protocol for
DNA extraction from Cochliobolus species and Yueqiang
Leng, Rui Wang, and Krishna D. Puri for technical
assistance.
References
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4. Luikart G, England PR, Tallmon D, Jordan S, Taberlet
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identification of thousands of copperhead snake
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16. Faircloth BC (2008) MSATCOMMANDER: detection of microsatellite repeat arrays and automated,
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Multiplex and Quantifiable
Detection of Infectious Fungi Using
Padlock Probes, General qPCR, and
Suspension Microarray Readout
33
Magnus Jobs, Ronnie Eriksson, and Jonas Blomberg
Abstract
By combining the multiplexing qualities of padlock probes and Luminex™
(Luminex Corporation, Austin, Texas, USA) technology, together with the
well-established quantitative feature of qPCR, a ten-plex fungal detection
protocol that quantitatively reveals ten different fungal species in a single
experiment has been devised. Padlock probes are oligonucleotides designed
to form circular DNA when hybridizing to specific target DNA. The 5¢ and
3¢ regions of the probes meet and ligate only when a specific target
sequence is present in the examined sample. The region of the padlock
probes that separates the target specific 5¢ and 3¢ ends contains general
primer sequences for amplification of circularized probes by means of
rolling circle amplification and qPCR. The interspersed region also contains specific tag sequences for subsequent Luminex recognition.
Keywords
Multiplex • Padlock probes • Rolling circle amplification • Fungal detection
• Suspension microarray
Introduction
M. Jobs (*)
Research Scientist, Dalarna University,
School of Health and Social Studies,
Högskolegatan 2, Falun 79188, Sweden
e-mail: mjb@du.se
R. Eriksson
Livsmedelsverket Sweden,
Box 662, Uppsala 75126, Sweden
J. Blomberg
Department of Medical Sciences Uppsala University,
Uppsala Academic Hospital, Dag Hammarskjölds v. 17,
Uppsala 75237, Sweden
A protocol for simultaneous quantitative detection
of ten different fungal species is described [1]. The
protocol involves padlock probes, qPCR, and
LuminexTM (Luminex Corporation, Austin, Texas,
USA) technology. A padlock probe is a long
oligonucleotide designed to hybridize to a specific
target sequence so that the 5¢ and 3¢ ends of
the probe meet. The DNA nick between the two
ends can be closed via enzymatic ligation, resulting
in a circularized probe [2]. Circularized probes
can be detected via rolling circle amplification
(RCA) and/or PCR followed by amplification
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_33, © Springer Science+Business Media, LLC 2013
363
364
product detection [3]. The amplification of circularized probes is made possible by including targets for PCR primers in the padlock probe region
separating the 5¢ and 3¢ regions. Detection of
amplification products can be done in several ways,
but a neat way is to introduce a specific address tag
sequence in the interspersed region that amplifies
along with the rest of the circularized probe [4].
If one or two fluorescent primers are used,
amplification products can then hybridize to solid
phase-bound anti-tag sequences (i.e., microarray
spots or Luminex beads) and be detectable [5].
Padlock probes have successfully been used in
various multiplex detection systems. The advantage of padlock probes in multiplexing is both the
specificity of the ligation reaction that forms the
circularized probes and the possibility to use a single PCR amplification primer-pair common for all
padlock probes included in the multiplex assay
when amplifying the ligation products [6–8]. The
protocol described here is for a ten-plex fungal
detection panel that quantitatively detects ten clinically important fungi. The technique is built on the
padlock probe concept, but two readouts are used:
(1) a SybrGreen™ real-time PCR for quantification
of circularized padlock probes, and (2) suspension
array technology (i.e., Luminex) for identification
of amplified sequences [1]. In Fig. 33.1 the padlock
probe concept has been outlined, illustrating the
different sections of the padlock probe and how the
probe can hybridize and become circularized when
a target sequence is present (A-B). The RCA and
subsequent real-time qPCR using a single universal primer-pair (one labeled primer) are also illustrated in Fig. 33.1 (C-E). The Luminex suspension
array consists of fluorescence-coded microspheres
with coupled oligonucleotide anti-tag sequences
for capturing of amplimers containing the sequence
tag. Thus, a bead is analogous to a microarray spot
[5]. The combination of two readouts of padlock
probe ligation, first via SybrGreen qPCR and then
by suspension microarray technology, allows the
well-established quantitative aspects of qPCR with
a wide dynamic range to be combined with specific
identification via the high multiplex capacity of
padlock probes. Included in the protocol is a strategy for improved hybridization of PCR amplimers
to the bead bound anti-tag. By letting labeled oligo-
M. Jobs et al.
nucleotides hybridize to the 3¢ and 5¢ end regions of
the amplimers, these regions become blocked for
rehybridization to the complementary PCR
amplimer strand. This reduces competition between
the anti-tag and the competing strand. Moreover,
the fluorophores on the end covering oligonucleotides contribute to an improved detection signal.
Materials
Solutions should be prepared using ultrapure
water. Pre-PCR reagents should be handled in a
clean room well separated from post-PCR activities. We use “PCR-grade water” when diluting
stocks and working solutions of oligonucleotides.
Shelf products in use in this protocol should be
handled and stored according to the manufacturer’s instructions. Oligonucleotides and PCR
reagents should be stored at −20°C and buffers
and microspheres at 4°C.
Materials for DNA Isolation
1. Use 0.5-mm-diameter zirconium-silica beads
(Techtum Lab AB, Umeå, Sweden) for bead
beating in a Tissuelyser (Qiagen AB,
Stockholm, Sweden). Use 2-mL Eppendorf
safe-lock
tubes
(VWR
International,
Stockholm, Sweden) in the bead beating
procedure.
2. Use NucliSENS® easyMAG™ lysis buffer
(BioMerieux Sverige AB, Askim, Sweden) to
lyse clinical samples. For sputum use 2×
Sputolysin® (Calbiochem, San Diego, CA,
USA).
3. Extract DNA on a NucliSENS® easyMAG™
according to manufacturer’s instructions.
Materials for Dilution Series
of Standards
Prepare stock solutions of synthetic targets
100 mM (100 pmol/mL) according to instructions
from the oligonucleotide manufacturer (Note 1).
Prepare two pools of synthetic targets, one pool
33
Multiplex and Quantifiable Detection of Infectious Fungi Using Padlock Probes…
365
Fig. 33.1 Outline of the padlock probe concept. (a) The
3¢ and 5¢ regions of the padlock probe (indicated in green)
are complementary to a specific target sequence. The
internal region contains a sequence designed for a general
primer-pair (indicated in yellow) and an address sequence
(indicated in blue). (b) When the target specific regions
hybridize to the target (gray), the 3¢ and 5¢ ends meet, and
via a ligation reaction the padlock probe becomes circularized. (c) A primer, complementary to one part of the
general primer-pair region, is then hybridized. (d)
A rolling circle amplification (RCA) is performed generating a long repetitive sequence. (e) Finally, using the general primer-pair (one of the primers is the same as that
used for the RCA), PCR is performed on the RCA product. The PCR product contains the address sequence and a
biotin moiety for subsequent detection in the Luminex
instrument. Reprinted with permission from Eriksson [1]
of targets for low efficiency padlock probes
(marked with down arrow in Table 33.1) and one
pool of targets for high efficiency padlock probes
(marked with up arrow in Table 33.1). Prepare
the two pools so that each of the five targets
included in a pool makes up one-fifth of the total
copy number in the pool. Do this by mixing 10 mL
from each stock of synthetic targets belonging to
the same category. Add 251 mL PCR-grade water
to the 50 mL pool stocks so that the pools contain
1013 molecules/mL (all five molecule species
combined). From here, use 0.05× Denhardt’s
solution (Sigma-Aldrich, St. Louis, Missouri,
USA, art. # D2532) instead of pure water when
preparing the standard dilution series. Dilute
1,000 times (1 mL target pool plus 999 mL 0.05×
Denhardt’s solution) and from that solution dilute
five times (50 mL target pool plus 200 mL 0.05
Denhardt’s solution) and from that solution serially dilute with a factor 10 (10 mL target pool plus
90 mL 0.05× Denhardt’s solution) until a series
ranging from 0.2 × 102 molecules/mL to 0.2 * 107
molecules/mL has been established. Make sure to
vortex every new dilution thoroughly before preparing the next dilution in the series.
Materials for Padlock Probe Ligation
and RCA
1. Use padlock probes for Candida albicans,
Candida glabrata, Candida tropicalis,
Candida parapsilosis, Cryptococcus neoformans, Aspergillus fumigatus, Aspergillus
flavus, Aspergillus niger, Aspergillus nidulans,
and Pneumocystis jirovecii (see Table 33.1),
and general forward primer (Biot-5¢AAGATATCGTAAGGAT-3¢) and general
reverse primer (5¢-TTGGATAAGTGGGATA-3¢) synthesized by Thermo Electron
366
Table 33.1 Padlock probe sequences for the ten targeted fungi marked with a star (*)
Fungal species
high eff. probe
¯ low eff. probe
A. flavus
A. fumigatus
A. nidulans
¯
A. niger
C. albicans
C. glabrata
¯
*PO4-TTGCGTTCGGCAAGCGCCATTGGTAAATTGGTAAATGAATTGATCCTTACGATA
TCTTGGATAAGTGGGATATCCAAGGTCAACCTGGAAAAAGATTGAT
**TCCGGCATCGATGAAGAACGCAGCCCCGGCCGGCGCTTGCCGAACGCAA|ATCAATCTTTTTCCAGGTTGACCTTGGATCAGGTAG
CATATCAATAAGCGGAGGA
*PO4-GGGTGTCGGCTGGCGCTTAGATGAATTGTGAAGTATTTAGATCCTTACGATATCT
TGGATAAGTGGGATAATCCGAGGTCAACCTTAGAAAAATAAAGTT
**TCCGGCATCGATGAAGAACGCAGCCCCGGCCGGCGCCAGCCGACACCC|AACTTTATTTTTCTAAGGTTGACCTCGGATCAGGTAGC
ATATCAATAAGCGGAGGA
*PO4-TCGAGCGGGTGACAAAGCCCTGAAATGAATGAATGATGAAATTGATCCTTACGA
TATCTTGGATAAGTGGGATAGCCCGGCCGGCCCTAA
**TCCGGCATCGATGAAGAACGCAGCCGAGCGTATGGGGCTTTGTCACCCGCTCGA|TTAGGGCCGGCCGGGCGCCAGCCGACGCATA
TCAATAAGCGGAGGA
*PO4-AGGCGCCGGCCAATCCTACGTAAAAAGAAAGGTATAAAGGTAAATCCTTACGAT
ATCTTGGATAAGTGGGATAGAAAGAATGGTTGGAAAACGTCGGC
**TCCGGCATCGATGAAGAACGCAGCTGCTCTGTAGGATTGGCCGGCGCCT|GCCGACGTTTTCCAACCATTCTTTCCAGGTTGCATAT
CAATAAGCGGAGGA
*PO4-CGCTACCGCCGCAAGCAATGATTTGAAGATTATTGGTAATGTAATCCTTACGAT
ATCTTGGATAAGTGGGATAAGGTCAAAGTTTGAAGATATACGTGGTAGA
**TCTCGCATCGATGAAGAACGCAGCAACATTGCTTGCGGCGGTAGCG|TCTACCACGTATATCTTCAAACTTTGACCTAAGCATATCAA
TAAGCGGAGGA
*PO4-GTTGGTAAAACCTAATACAGTATTAACCCCGATTGATTATTGTGATTTGAATTG
ATCCTTACGATATCTTGGATAAGTGGGATACTTATCCCTCCCTAGATCAACACCGA
**TCTCGCATCGATGAAGAACGCAGCCGGGGGTTAATACTGTATTAGGTTTTACCAAC|TCGGTGTTGATCTAGGGAGGGATAAGTGG
CATATCAATAAGCGGAGA
*PO4-CCACTAGCAAAATAAGCGTTTTTGGATAAATGATATGAATTGGATTATTGGTAT
ATCCTTACGATATCTTGGATAAGTGGGATAAGGTCAAAGTTATGAAATAAATTGTGGTGG
**TCTCGCATCGATGAAGAACGCAGCTTTATCCAAAAACGCTTATTTTGCTAGTGG|CCACCACAATTTATTTCATAACTTTGACCTGC
ATATCAATAAGCGGAGGA
M. Jobs et al.
C. tropicalis
*Padlock probe sequence (5¢–3¢)
**Synthetic target sequence (5¢–3¢)
33
Multiplex and Quantifiable Detection of Infectious Fungi Using Padlock Probes…
*PO4-GGAGTTTGTACCAATGAGTGGAAAAAAACGTTAGTTAGATTATTGTTAGTTAGA
TCCTTACGATATCTTGGATAAGTGGGATATGATTTGAGGTCGAATTTGGAAGAAGTTTT
¯
**TCTCGCATCGATGAAGAACGCAGCGTTTTTTTCCACTCATTGGTACAAACTCC|AAAACTTCTTCCAAATTCGACCTCAAATCAGCAT
ATCAATAAGCGGAGA
*PO4-GCCGAAGACTACCCCATAGGCCGTAAGATGTTGATATAGAAGATTAATCCTTA
Cr. neoformans
CGATATCTTGGATAAGTGGGATAAAACAAAAAAGAGATGGTTGTTATCAGCAA
¯
**TTCCACATCGATGAAGAACGCAGCTGGGCCTATGGGGTAGTCTTCGGC|TTGCTGATAACAACCATCTCTTTTTTGTTTGAGCATAT
CAATAAGCGGAGGA
*PO4-GAATTTCAGACTAGCATGCATATAATTATTTAATGTTGTGAATAATGTAGAAAG
P. Jiroveci
ATCCTTACGATATCTTGGATAAGTGGGATAGACACTAGGCAAAGAAAAAAAGCTACTTTT
¯
**TCTCGCGTCGATGAAGAACGTGGCAATAATTATATGCATGCTAGTCTGAAATTC|AAAAGTAGCTTTTTTTCTTTGCCTAGTGTCGCA
TATCAATAAGCGGAGGA
The target matching sections are underlined and the address-tags are shown in bold. The remaining unmodified section is the general primer-pair sequence, identical for all
padlock probes. The synthetic target sequences are marked with a double star (**). Here the underlined section represents the padlock probe matching section. The padlock probe
ligation position has been pointed out with a bar ( | ). Extra sequences flanking the target region have been included and represent interspecies conserved sequences (in vivo these
conserved regions may not be immediately adjacent to the padlock probe matching sections)
C. parapsilosis
367
368
M. Jobs et al.
Table 33.2 Outline of amino-modified carbon 12-linked anti-tag sequences coupled to different color-coded
microspheresa
Anti-tag for
A. flavus
A. fumigatus
A. nidulans
A. niger
C. albicans
C. glabrata
C. tropicalis
C. parapsilosis
Cr. Neoformans
P. jiroveci
Anti-tag sequence
N-C12-ATTGGTAAATTGGTAAATGAATTG
N-C12-TTAGATGAATTGTGAAGTATTTAG
N-C12-TGAAATGAATGAATGATGAAATTG
N-C12-GTAAAAAGAAAGGTATAAAGGTAA
N-C12-GATTTGAAGATTATTGGTAATGTA
N-C12-GATTGATTATTGTGATTTGAATTG
N-C12-TGATATGAATTGGATTATTGGTAT
N-C12-GTTAGTTAGATTATTGTTAGTTAG
N-C12-GTAAGATGTTGATATAGAAGATTA
N-C12-TAATGTTGTGAATAATGTAGAAAG
MicroplexTM xTAG
LUA-7
LUA-90
LUA-35
LUA-30
LUA-4
LUA-5
LUA-70
LUA-80
LUA-9
LUA-40
The anti-tags match tags included in padlock probes targeting the different fungal species indicated. Pre-coupled
microspheres, with the same anti-tags, can be ordered directly from the Luminex, and the table points out the corresponding Microplex microspheres
a
GmbH, Ulm, Germany. All padlock probes
must be synthesized 5¢ phosphorylated and the
general forward primer must have a 5¢ biotin
moiety. Dilute padlock probes and primers
in PCR-grade water to 100 mM (100 pmol/mL)
stock solutions (the manufacturer provide
information on what volume to use). Prepare
1 mM work solutions of each padlock probe
by diluting the main stock solutions a hundred times. Then prepare a mixture of all ten
padlock probes with a 10 nM concentration
of each padlock probe (for a 500-mL mixture
take 5 mL of each padlock probe work solution and add 450 mL PCR-grade water).
Prepare two different general forward primer
work solutions, one 1 mM solution for use
when setting up the RCA reaction, and one
10 mM solution for use when setting up the
PCR. Dilute the general reverse primer to a
work solution of 10 mM.
2. Ligation reagents: 10× Ampligase® reaction
buffer, Ampligase thermostable DNA ligase
(Epicentre Biotechnologies, WI, USA), PCRgrade water (Applied Biosystems, Stockholm,
Sweden), and padlock probe mix with 10 nM
of each padlock probe (for padlock probe mix
preparation, see previous).
3. RCA reagents: 10× Phi29 buffer (Fermentas,
Vilnius, Lithuania) dNTP 10 mM, BSA
2 mg/mL, Phi29 DNA polymerase 10 U/mL,
and general forward primer 1 mM.
Materials for PCR
1. PCR reagents: 10× SYBR® Green PCR Buffer,
dNTPs including dUTP 12.5 mM, MgCl2
25 mM, AmpliTaq Gold® DNA Polymerase
5 U/mL, AmpErase® UNG 1 U/mL (Applied
Biosystems, Stockholm, Sweden), and general
forward and reverse primers 10 mM of each.
2. PCR equipment: Carry out PCR on a RotorGene 3000 (Corbett Life Science, Concorde,
New South Wales, Australia) in thin walled
0.2 mL PCR tubes (Qiagen, Stockholm
Sweden). Instrument software: Rotor-gene 6.1.
Materials for Luminex (Coupling of
Anti-Tag Oligonucleotides)
The following material is needed for coupling
of anti-tag oligonucleotides to FlexMap™
microspheres: 5¢C-12 amino-modified oligonucleotide anti-tags 100 mM (Table 33.2)
(Biomers.net GMbH, Ulm, Germany), ten sets
(“microsphere regions”) of carboxylated polystyrene FlexMap™ microspheres (Luminex
Corporation, Austin, Texas, USA), 0.1 M
2-morpholinoethane sulfonic acid (MES, pH
4.5) MES buffer, desiccated 1-ethyl-3-3-dimethylaminopropyl carbodiimide (EDC) (Pierce
Thermo Fisher Scientific), 0.02% Tween-20 and
0.1% SDS, Tris–EDTA (TE, pH 8.0) buffer.
33
Multiplex and Quantifiable Detection of Infectious Fungi Using Padlock Probes…
Coupling of anti-tag oligonucleotides is performed as follows:
1. Bring two 10 mg aliquots of desiccated EDC
powder from the freezer and the stocks of the
ten different FlexMap™ microspheres from
the fridge. Let the reagents adjust to room
temperature (Note 2).
2. Resuspend the microspheres by first vortexing and then sonication for 20 s.
3. Transfer 2.5 million (200 mL from stock) of
each set of microspheres to ten pre-labeled
1.5-mL Eppendorf tubes (Note 3).
4. Pellet the microspheres by microcentrifugation at ³8,000 × g for 1–2 min.
5. Remove the supernatants and resuspend the
microspheres in 25 mL MES buffer (vortex
and sonicate for 20 s).
6. Add 2 mL of the ten anti-tag oligonucleotides
to the corresponding microsphere sets and
vortex.
7. Prepare a fresh solution of 10 mg/mL EDC
by adding 1 mL water to one of the aliquots
of desiccated EDC.
8. Quickly add 2.5 mL of the fresh EDC solution
to all ten microsphere sets and vortex.
9. Incubate at room temperature for 30 min in
darkness.
10. Prepare another fresh solution of EDC by
adding 1 mL water to the second EDC
aliquot.
11. Again quickly add 2.5 mL of the fresh EDC
solution to all ten microsphere sets and vortex.
12. Incubate for another 30 min at room temperature in darkness.
13. Add 0.5 mL 0.02% Tween-20 to each set.
14. Pellet the sets by microcentrifugation at
³8000 × g for 1–2 min.
15. Remove the supernatant and resuspend the
sets in 0.5 mL 1% SDS (vortex).
16. Pellet the sets by microcentrifugation at
³8,000 × g for 1–2 min.
17. Remove the supernatant and resuspend the
sets in 50 mL TE buffer (pH 8.0).
18. Prepare a mixture of all ten sets by mixing
equal volumes of each.
19. Store the sets and the mixture of coupled
microspheres (50,000 spheres/mL) in the
fridge (dark).
369
The coupling procedure is a modification of an
original Luminex protocol (see Notes 4 and 5).
Materials for Luminex (Hybridization
and Detection)
1. Tetramethyl ammonium chloride (TMAC)
hybridization buffer (4.5 M TMAC, 0.15%
sodium lauryl sarcosinate; “sarkosyl”(SigmaAldrich), 75 mM Tris–HCl pH 8 and 6 mM
EDTA pH 8) and Tris–EDTA (TE, pH 8.0)
buffer are used in the hybridization of tagged
PCR products to microsphere-bound anti-tags.
In the hybridization mixture 5¢ Cy3 labeled
end covering oligonucleotides (Cy3-5¢-ATCCTTACGATATCTT-3¢ and Cy3-5¢-TTGGATAAGTGGGATA-3¢) 10 mM (Thermo Electron
GmbH, Ulm, Germany) is included.
Streptavidin-R-Phycoerythrin 0.15 mg/mL
(Qiagen AB, Stockholm, Sweden) is used to
label
the
biotinylated
PCR
products. Wells from non-skirted thin-wall
96 × 0.2 mL plates (Bioplastics, Landgraaf,
The Netherlands) are suitable for the hybridization solution (the plates fit the heater block
part# 67-50066-00-001).
2. The Luminex 200 (Luminex Corporation,
Austin, Texas, USA) system together with
STarStation software (Applied Cytometry
System, Sheffield, UK) is used to analyze the
amount of hybridized products.
Method
DNA Isolation
Clinical samples may be in the form of charcoal
swabs, urine, vaginal swab culture broth, bronchoalveolar lavage, or sputum. Procedures for
initiating DNA isolation vary for the different
types. Steps 1–3 represent the start of the DNA
isolation procedure for the different types of clinical samples.
1. Wash charcoal swabs in 600 mL NucliSENS®
easyMAG™ lysis buffer by inserting the swab
in a 1.5-mL Eppendorf tube containing the
370
2.
3.
4.
5.
6.
buffer and rotate the swab thoroughly. Then
proceed to step 4.
Transfer 1 mL urine, vaginal swab broth, or
bronchoalveolar lavage to 1.5 mL Eppendorf
tube and centrifuge the sample at 13,000 × g for
20 min. Discard the supernatant and dissolve
the pellet in 600 mL NucliSENS® easyMAG™
lysis buffer. Then proceed to step 4.
Dilute sputum samples in an equal volume of
2× Sputolysin® (1× Sputolysin® final concentration), and vortex thoroughly. Incubate for
20 min and then proceed to step 4.
Perform cell lysis by bead beating. First
transfer the suspension to 2-mL Eppendorf
safe-lock tubes containing 600 mL 0.5-mmdiameter zirconium-silica beads. Then shake
the samples in a Tissuelyser for 10 min at
30 Hz.
Transfer the lysates to clean 15-mL Falcon
tubes (leave the beads in the original tube).
Rinse the beads two times in 1 mL NucliSENS®
easyMAG™ lysis buffer and add the rinse
solution to the collected lysate in the Falcon
tube. The final volume of cell lysate will be
approximately 2.5 mL (it is impossible to
recover the entire volume from the bead slur).
Add the full sample volume to a sample vessel
for NucliSENS® easyMAG™ extraction robot
and run the extraction according to manufacturer’s instructions. Set the elution volume to
60 mL. Store extracts at −20°C.
Ligation and RCA
The reaction mixtures of both the ligation reaction and the RCA reaction must be prepared in a
clean room well separated from samples, synthetic targets, and PCR products from previous
experiments. The following protocol is based on
a single reaction and needs to be multiplied to
match the number of samples analyzed. In fact,
preparing a reaction mixture for a single reaction
is not recommended, because it is difficult to
pipette such small volumes accurately. Including
the two standard dilution series, counting five
concentrations each (described above), means
M. Jobs et al.
that at least a volume for ten reactions must be
prepared only to cover the dilution series.
1. Bring out and thaw the material for the ligation reaction (described above). Prepare the
ligation reaction mixture by mixing 1 mL of
10× Ampligase® reaction buffer, 0.2 mL
Ampligase thermostable DNA ligase, 1 mL
Padlock probe mixture, and 4.8 mL PCR-grade
water. Add 3 mL of DNA extract to the reaction mixture, pipette-mix, and place the reaction on a thermal cycler at 95°C for 2 min,
55°C 30 min, and keep at 4°C until the next
step.
2. Bring out and thaw the RCA components
(described previously). Prepare the RCA reaction by mixing 2 mL 10× Phi29 reaction buffer, 2 mL BSA, 0.5 mL forward primer, 0.25 mL
dNTP and 0.3 mL Phi29 DNA polymerase.
Add 4.95 mL PCR-grade water (to reach a volume of 10 mL). Finally, mix the RCA reaction
mixture with the fully incubated ligation reaction (20 mL final volume), and incubate at
37°C for 30 min, 85°C for 4 min, and keep at
4°C until the next step.
PCR
The PCR reaction mixture must be prepared in a
clean room well separated from samples and PCR
products from previous experiments.
1. Prepare a 25 mL PCR mixture by mixing
2.5 mL 10× SYBR® Green PCR Buffer, 0.4 mL
dNTP (including dUTP), 1.5 mL MgCl2,
0.25 mL AmpliTaq Gold® DNA Polymerase,
0.25 mLAmpErase® UNG, and 0.75 mL of each
general padlock probe primer. Add 16.1 mL
PCR-grade water. Finally add 2.5 mL of the
RCA product.
2. Program your real-time PCR instrument (we
use a Rotor-Gene 3000 from Corbett Life
Science) as follows: 50°C for 2 min, 95°C for
10 min, followed by 30 cycles of 95°C for
15 s, 52°C for 15 s and 68°C for 30 s. Monitor
fluorescence during the 68°C step in each temperature cycle. Monitor in the FAM/Sybr
channel (source: 470 nm, detector: 510 nm).
33
Multiplex and Quantifiable Detection of Infectious Fungi Using Padlock Probes…
3. The real-time PCR will reveal any ligation
event in the previous steps in the procedure so
that at this point it is appropriate to review the
real-time PCR data to decide whether to proceed in the protocol or not.
Measurements in the Luminex 200
Instrument
1. Bring all the Luminex components (see previous) to room temperature. Prepare the hybridization reaction by vortexing and sonicating
(for 20 s) the anti-tag coupled microsphere
mixture (containing all ten anti-tag coupled
microspheres). Dilute the microsphere mixture
in TMAC hybridization solution by mixing
1.5 mL microsphere mixture with 33 mL TMAC
(use wells from low non-skirted thin-wall 96
well plates). Then add 1 mL of each end covering oligonucleotide, 8.5 mL TE buffer, and
finally 5 mL PCR product.
2. Pipette-mix the hybridization mixture and
denature it for 2 min at 95°C (preferably in a
96-well PCR instrument). Place the wells in
a heater block on a shaking table heated to
50°C and allow hybridization for 30 min.
Then add 2 mL streptavidin-R-Phycoerythrin,
pipette-mix, and let the incubation continue
for another 15 min.
3. Prepare the Luminex instrument by adjusting the
analysis probe (the suction needle) to the current
sample plate and run the start up scripts. Program
the instrument to analyze median fluorescence
intensity (MFI) from the ten microsphere regions
to which anti-tags have been coupled. Initiate the
assay by setting the instrument to calculate MFI
based on 100 measurements from each microsphere set and allow the Luminex XYP reach
50°C. Place the heater block with the samples in
the instrument and run.
Analysis
PCR amplification data are preferably reviewed
in the dedicated instrument software. Instruct the
PCR software to automatically identify cut off
(Ct) values. MFI from all ten microsphere regions
371
are presented as numbers by the Luminex instrument and can easily be exported to an Excel file
(or be reviewed directly in the instrument software). Figures 33.2a and 33.2b show data from a
titration series of a synthetic Candida glabrata
sequence. The MFIs have been exported to Excel
and converted into a diagram.
When running clinical samples, the use of
standard titration series with known starting copy
numbers run in parallel makes it possible to estimate sample copy numbers. In a ten-plex assay
ten such dilution series of the different targets
would be very impractical and expensive to
include. Instead all ten targets could be pooled so
that each target makes up one-tenth of the total
copy number in the pool. The SybrGreen-based
qPCR would then detect the pool as a single target species and the copy number of the sample
could be calculated based on the amplification of
the pool (given that the sample contains only one
fungal species). However, we have noticed that
the padlock probes matching the different fungal
species amplify with different efficiencies resulting in different Ct-values despite the same target
copy numbers. Therefore, we have categorized
the padlock probes in two categories and made
two pools of targets: one pool containing targets
for the more efficient padlock probes and one for
the less efficient. Both pools contain five target
species. The total copy numbers of the five targets in a pool together make up the standard for
each dilution step. In this way two standard
curves with slightly different slopes will be produced. The two standard curves encompass most
of the differences between padlock probes, and
make it possible to approximate sample copy
numbers once the Luminex analysis has revealed
what fungal species was present in the sample.
Notes
1. Take special care to avoid contamination when
handling concentrated synthetic targets.
Prepare and store the high concentration targets in a room separated from where the main
laboratory work is conducted.
2. Prepare several Eppendorf tubes (1.5 mL)
with 10 mg EDC aliquots in each in advance
372
Fig. 33.2 The figure shows the result of padlock probebased analyses of a titration series of a synthetic fungal
DNA sequence (Candida glabrata). (a) Results from
SybrGreen-based real-time PCR data. (b) Luminex
M. Jobs et al.
median fluorescence intensity (MFI) data. The figure
demonstrates the methods’ ability to quantify a sample
in ten-plex mode. Reprinted with permission from
Eriksson [1]
33
Multiplex and Quantifiable Detection of Infectious Fungi Using Padlock Probes…
and store them in a sealed container together
with desiccant.
3. Label the tubes clearly with information on
microsphere region and what fungus the corresponding anti-tag will detect.
4. Coupled microspheres for 500 reactions are
produced by following this coupling protocol
so that it is not necessary to do this every time
you run an assay.
5. Microspheres with pre-coupled anti-tags
(microplex xTAG microspheres) can be
ordered from Luminex avoiding the coupling
procedure all together. In Table 33.2 the corresponding microplex xTAG microspheres are
indicated.
2.
3.
4.
5.
6.
7.
References
1. Eriksson R, Jobs M, Ekstrand C, Ullberg M, Herrmann
B, Landegren U et al (2009) Multiplex and quantifiable
detection of nucleic acid from pathogenic fungi using
padlock probes, generic real time PCR and specific
8.
373
suspension array readout. J Microbiol Methods
78:95–202
Nilsson M, Malmgren H, Samiotaki M, Kwiatkowski
M, Chowdhary BP, Landegren U (1994) Padlock
probes: circularizing oligonucleotides for localized
DNA detection. Science 265(5181):85–2088
Baner J, Nilsson M, Mendel-Hartvig M, Landegren U
(1998) Signal amplification of padlock probes by rolling
circle replication. Nucleic Acids Res 26:5073–5078
Baner J, Gyarmati P, Yacoub A, Hakhverdyan M,
Stenberg J, Ericsson O et al (2007) Microarray-based
molecular detection of foot-and-mouth disease, vesicular stomatitis and swine vesicular disease viruses, using
padlock probes. J Virol Methods 143:200–206
Fulton RJ, McDade RL, Smith PL, Kienker LJ, Kettman
JR Jr (1997) Advanced multiplexed analysis with the
FlowMetrix system. Clin Chem 43:1749–1756
Nilsson M, Landegren U, Antson DO (2002) Singlenucleotide sequence discrimination in situ using padlock
probes. Curr Protoc Hum Genet. Chapter 4:Unit 4: 11
Baner J, Isaksson A, Waldenstrom E, Jarvius J,
Landegren U, Nilsson M (2003) Parallel gene analysis
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GL (2009) Multiplex single nucleotide polymorphism
(SNP)-based genotyping in allohexaploid wheat using
padlock probes. Plant Biotechnol J 7:375–390
Rapid Deletion Plasmid
Construction Methods
for Protoplast and Agrobacteriumbased Fungal Transformation
Systems
34
María D. García-Pedrajas, Zahi Paz,
David L. Andrews, Lourdes Baeza-Montañez,
and Scott E. Gold
Abstract
The increasing availability of genomic data and the sophistication of analytical
methodology in fungi have increased the need for functional genomics tools
in these organisms. Gene deletion is a critical tool for functional analysis.
The targeted deletion of genes requires both a suitable method for the transfer
of foreign DNA to fungal cells and the generation of deletion constructs.
The deletion constructs should contain the regions flanking the gene of interest, while the ORF is replaced by a DNA fragment harboring a marker that
allows selection of cells transformed with this foreign DNA. Deletion mutants
are produced upon transformation by integration of this construct into the
M.D. García-Pedrajas (*) • L. Baeza-Montañez
Instituto de Hortofruticultura Subtropical y Mediterránea
“La Mayora”, Consejo Superior de Investigaciones
Científicas (IHSM-UMA-CSIC), Estación Experimental
“La Mayora”, Algarrobo-Costa, Málaga, E-29750, Spain
e-mail: mariola@eelm.csic.es; lourdes@eelm.csic.es
Z. Paz • D.L. Andrews
Department of Plant Pathology, Miller Plant Science
Building, University of Georgia, Athens, Georgia
30602-7274, USA
e-mail: zpaz@uga.edu; dandrew@uga.edu
S.E. Gold
Research Plant Pathologist, United States Department of
Agriculture - Agricultural Research Unit (USDA-ARS),
Toxicology & Mycotoxin Research Unit, 950 College
Station Road, Athens, Georgia 30605, USA
e-mail: scott.gold@ars.usda.gov
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_34, © Springer Science+Business Media, LLC 2013
375
376
M.D. García-Pedrajas et al.
fungal genome by homologous recombination. Protoplasts have been widely
used as starting material for genetic transformation in fungal species.
However, a number of fungi have proven to be recalcitrant to protoplastmediated transformation (PMT). Among the alternative methodologies
developed for those species, Agrobacterium tumefaciens-mediated transformation (ATMT) has been particularly successful, becoming the preferred
genetic transformation method for an increasing number of fungi. Here we
describe two methods to rapidly generate plasmid-based gene deletion constructs, namely, DelsGate and OSCAR, which are compatible with PMT and
ATMT, respectively. Both procedures are based on PCR of the target gene
flanks and Gateway cloning technology, allowing generation of deletion constructs in a very simple and robust manner in as little as 2 days. Gateway
vectors have been modified so that a single Gateway cloning step generates
the deletion construct itself. The PCR and transformation steps these methodologies involve should be well suited for high-throughput approaches to gene
deletion construction in fungal species in which either of the two major DNA
transformation methods, PMT or ATMT, is used. We describe here the entire
process, from the generation of the deletion constructs to the analysis of the
fungal transformants for gene replacement confirmation, with the
Basidiomycete fungus Ustilago maydis for DelsGate and PMT, and with the
Ascomycete fungus Verticillium dahliae for OSCAR and ATMT.
Keywords
Gene deletion • Plasmid construction • Protoplast • Agrobacterium
• Fungal transformation • Deletions via Gateway (DelsGate) • One-step
construction of Agrobacterium-recombination-ready plasmids
Introduction
The genomic era is generating large sets of
candidate genes with potential roles that are
worth functional exploration, through either the
sequencing and annotation of fungal genomes or
other large-scale approaches such as transcriptome analysis. Therefore, there is a great demand
for fast and simple methods to generate deletion
constructs
suitable
for
high-throughput
approaches to gene deletion. Here we describe
two methods to rapidly generate gene deletion
constructs, namely DelsGate (Deletions via
Gateway) [1] and OSCAR (one-step construction
of Agrobacterium-recombination-ready plasmids) [2], which are compatible with PMT and
ATMT, respectively. Both approaches to gene
deletion construction combine PCR with Gateway
cloning technology [3]. Gene flanking regions
are amplified by PCR, allowing for precise
deletion of the gene of interest. Although
the frequency of integration by homologous
recombination can vary among fungal species,
and even within species among genes, we regularly use 1 kb of the 5¢ and 3¢ gene flanks in the
deletion construct to promote homologous
recombination. That flank length works well to
promote homologous recombination for both U.
maydis and PMT and for homologous integration
of the T-DNA in the ATMT of V. dahliae, and
should be sufficient in most species. As the
Gateway system developed by Invitrogen is used
for very efficient cloning of the amplified gene
flanks, appropriate recombination sites are
34
Rapid Deletion Plasmid Construction Methods for Protoplast and Agrobacterium-based…
introduced via the primers into the PCR products
during amplification. To speed up the process,
appropriate vectors to be used with Gateway
cloning technology have been developed separately for DelsGate and OSCAR, so that after
PCR of the gene flanks, a single cloning step generates the deletion construct itself.
In DelsGate, during PCR the attB1 and attB2
recombination sites are introduced in the 5¢ flank
and 3¢ flank PCR products, respectively, to
promote in vitro recombination with a donor vector containing the attP1 and attP2 sites. In the
donor vector the attP1 and attP2 sites flank the
ccdB gene and the in vitro recombination by the
BP clonase replaces this gene with the PCR products generating an entry clone. The ccdB gene is
lethal for most Escherichia coli strains including
the commonly used strain employed in this study,
DH5a, thus transformants harboring the PCR
products are highly favored to produce colonies.
The Invitrogen pDONR201 vector was modified
to be used with DelsGate by introduction of suitable selectable markers for fungal transformation.
We also greatly simplified the process by taking
advantage of a report by Suzuki et al. [4] showing
that two independent PCR fragments, each carrying an attB1 or attB2 site on one end, can simultaneously recombine with a single pDONR vector.
This generates a linear construct that is then circularized in vivo via E. coli transformation, provided that there are homologous sequences at the
free ends of the PCR fragments to promote recombination. In the DelsGate method the sequence
added to the 5¢ and 3¢ flanks to promote homologous recombination in vivo is the 18 bp recognition site for the homing endonuclease I-SceI,
absent in most fungal genomes. Thus, in addition
to promoting homologous recombination, this site
is then universally used to generate the linear
DNA for fungal transformation without concern
for inadvertent digestion of the gene flanks.
The OSCAR method is in turn an adaptation
of the available MultiSite Gateway system; however, as with DelsGate a single BP clonase reaction generates the deletion construct. ATMT
requires the generation of deletion constructs in a
binary vector. When using the approach for targeted gene deletion, constructs are designed to
contain, between the T-DNA borders, the flanking
regions of the gene of interest with the intervening
377
ORF replaced by a selectable marker. As above
stated, the OSCAR approach has also been
designed so that a single cloning step is required
to produce the deletion construct in the binary
vector ready for ATMT. With that purpose, two
new vectors were developed. The first vector
developed is a binary vector suitable for ATMT
in fungi that harbors the toxic gene ccdB flanked
by the recombination sites attP2r and attP3,
between the left and right T-DNA borders. The
second vector contains a hygR marker suitable
for selection of transformants in Ascomycete
fungi, flanked by the recombination sites attP1r
and attP4 in pBluescript II KS(+). During PCR,
recombination sites attB2r and attB1r and recombination sites attB4 and attB3 are introduced at
the ends of the 5¢ and 3¢ gene flanks, respectively.
When these PCR products are incubated with the
two vectors in the presence of BP clonase, each
attB site should recombine only with its single
compatible attP site generating the deletion construct. Again, transformation of E. coli DH5a, or
similar strain, prevents selection of the original
binary vector because it contains the ccdB gene,
and selection for spectinomycin resistance prevents selection of the marker vector since this
confers ampicillin and not spectinomycin resistance. Deletion constructs have then only to be
transformed into Agrobacterium tumefaciens in
preparation for fungal transformation.
For user convenience we describe here the
entire DelsGate process from the production of
the deletion construct to transformation of the
fungus and confirmation of gene deletion for the
Basidiomycete U. maydis. Similarly we describe
the entire process of generation of OSCAR constructs and their use in ATMT with the Ascomycete
fungus V. dahliae.
Materials
Culture Media
1. Potato dextrose agar supplemented to 2% agar
(2PDA): 39 g PDA powder (Difco, Franklin
Lakes, NJ), 5 g supplemental agar, 1 L dH2O.
2. Potato dextrose both (PDB): 24 g PDB powder
(Difco), 1 L dH2O. After autoclaving, store at
room temperature (RT) (see Note 1).
378
3. Low Na LB antibiotic plates: 1% bactotryptone, 0.5% yeast extract, 0.1% NaCl, antibiotic of choice. To prepare 1 L: 10 g tryptone,
5 g yeast extract, 1 g NaCl, 20 g agar, dH2O
to 1 L. After autoclaving add 0.5 mL of a
100 mg/mL solution of kanamycin sulfate
(Sigma-Aldrich, Saint Louis, MO), or 0.5 mL
of a 100 mg/mL solution of ampicillin
(Sigma-Aldrich), or 0.5 mL of 100 mg/mL
solution of spectinomycin (Sigma-Aldrich),
according to the plasmid to be selected.
4. YEPS medium: 1% yeast extract, 2% bactopeptone, and 2% sucrose. For 500 mL: dissolve 5 g of yeast extract, 10 g of bactopeptone,
and 10 g sucrose in a final volume of 500 mL
dH2O, dispense in aliquots of 100 mL in 500mL flasks. After autoclaving, store at 4°C.
PCR Amplification of Gene Flanks
and Clean Up of PCR Products
1. Primers: gene-specific primers 1 and 2 to
amplify 5¢ flanks and 3 and 4 to amplify 3¢
flanks. For DelsGate primers 1 and 2 contain at
their 5¢ end the I-SceI recognition sequence (in
the forward orientation) and the attB1 sequence,
respectively, and primers 3 and 4 contain at their
5¢ ends the attB2 sequence and the I-SceI recognition sequence (in the reverse orientation),
respectively (Table 34.1). For OSCAR, primers
1 and 2 contain at their 5¢ ends the attB2r and
attB1r sequences, respectively, and primers 3
and 4 contain at their 5¢ ends the attB4 and attB3
sequences, respectively (see Table 34.1).
2. Taq-polymerases and reaction buffers. Because
the bands to be amplified are only 1 kb long any
taq-polymerase of general use works well for
this step. We regularly use homemade taqpolymerase. For commercial taq-polymerases
the reaction buffer provided with them is used.
For homemade taq-polymerase the 10× buffer
we prepare contains: 0.5 M KCl, 100 mM Tris–
HCl pH 8.3, 0.1% gelatin, 1% Triton X-100.
3. 30% PEG 8000/30 mM MgCl2. To prepare
100 mL dissolve 30 g of polyethylene glycol
(PEG) 8000 (Sigma-Aldrich) in approximately
70 mL dH2O, bring volume to 90 mL, after
autoclaving add 10 mL of sterile 0.3 M MgCl2.
M.D. García-Pedrajas et al.
Store at 4 ºC or dispense in aliquots and freeze
at −20 ºC.
4. QIAquick PCR purification kit (QIAGEN,
Valencia, CA).
5. QIAquick gel extraction kit (QIAGEN).
BP Clonase Reaction
1. Gateway BP clonase II enzyme mix
(Invitrogen, Carlsbad, CA).
2. Modified Gateway donor vectors for DelsGate:
For PMT of U. maydis: pDONR-Cbx and
pDONR-Hyg; for PMT of Ascomycete fungi:
pDONR-A-Hyg (Fig. 34.1a). Note that these
vectors are freely available from the Fungal
Genetics Stock Center.
3. Modified Gateway binary and marker vectors
for OSCAR: pOSCAR (binary vector) and
pA-Hyg-OSCAR (marker vector) (see
Fig. 34.1b). Note that these vectors are also
freely available upon request from the Fungal
Genetics Stock Center.
4. Donor vectors and the binary vector are maintained in E. coli strain DB3.1 (Invitrogen)
since they contain the ccdB gene which is
toxic to most other E. coli strains used in
molecular biology.
Transformation of Bacterial Strains
1. E. coli strain DH5a (Bethesda Research
Laboratories).
2. To increase transformation frequencies, commercial One Shot® MAX Efficiency™ DH5aT1R, One Shot® Mach1™ T1R, or One Shot®
OmniMAX™ 2-T1R E. coli competent cells
(Invitrogen) can be used.
3. A. tumefaciens strain AGL-1 [5].
Verification of Deletion Constructs
1. Primers to verify DelsGate constructs: SceI-F
and SceI-R (see Table 34.1) combined with
gene-specific primers from the sect. PCR
Amplification of Gene Flanks and Clean Up
of PCR Products or alternatively with vector
34
Rapid Deletion Plasmid Construction Methods for Protoplast and Agrobacterium-based…
379
Table 34.1 Primers used for DelsGate and OSCAR deletion construct generation, verification of deletion constructs,
and testing of deletion mutants
Primer
Primer 1-(I-SceIF)
Method
DelsGate
Primer 2-(attB1)
DelsGate
Primer 3-(attB2)
DelsGate
Primer 4-(I-SceIR)
DelsGate
Primer 1-(attB2r)
OSCAR
Primer 2-(attB1r)
OSCAR
Primer 3-(attB4)
OSCAR
Primer 4-(attB3)
OSCAR
SceI-F
DelsGate
SceI-R
DelsGate
DonrF-C
DelsGate
DonrF-H
DelsGate
DonrR
DelsGate
OSC-F
OSCAR
Hyg-R(210)
OSCAR
Hyg-F(850)
OSCAR
OSC-R
OSCAR
CbxF-DG
DelsGate
Hyg-DG
DelsGate
Use
Amplification of 5¢ flank,
primer forward
Amplification of 5¢ flank,
primer reverse
Amplification of 3¢ flank,
primer forward
Amplification of 3¢ flank,
primer reverse
Amplification of 5¢ flank,
primer forward
Amplification of 5¢ flank,
primer reverse
Amplification of 3¢ flank,
primer forward
Amplification of 3¢ flank,
primer reverse
Verification of deletion
construct
Verification of deletion
construct
Verification of deletion
construct when using
pDONR-Cbx
Verification of deletion
construct when using
pDONR-A-Hyg and
pDONR-Hyg
Verification of deletion
construct for all vectors
Verification of presence
of 5¢ flank in deletion
construct
Verification of presence
of 5¢ flank in deletion
construct and verification
of gene deletion
Verification of presence
of 3¢ flank in deletion
construct
Verification of presence
of 3¢ flank in deletion
construct
Verification of gene
deletion when using
pDONR-Cbx
Verification of gene
deletion when using
pDONR-Hyg
Sequence
5¢- TAGGGATAACAGGGTAAT-(gene-specific
sequence, N20-25)-3¢
5¢-GGGGACAAGTTTGTACAAAAAAGCAGGC
TAA-(gene-specific sequence N20-25)-3¢
5¢-GGGGACCACTTTGTACAAGAAAGCTGGG
TA-(gene-specific sequence, N20-25)-3¢
5¢-ATTACCCTGTTATCCCTA-(gene-specific
sequence, N20-25)-3¢
5¢-GGGGACAGCTTTCTTGTACAAAGTGGAA(gene-specific sequence, N20-25)-3¢
5¢-GGGGACTGCTTTTTTGTACAAACTTGT(gene-specific sequence, N20-25)-3¢
5¢-GGGGACAACTTTGTATAGAAAAGTTGTT(gene-specific sequence, N20-25)-3¢
5¢-GGGGACAACTTTGTATAATAAAGTTGT(gene-specific sequence, N20-25)-3¢
5¢- TAGGGATAACAGGGTAAT-3¢
5¢-ATTACCCTGTTATCCCTA-3¢
5¢-TCGCGTTAACGCTAGCATGGATCTC-3¢
5¢-ATCAGTTAACGCTAGCATGGATCTC-3¢
5¢-GTAACATCAGATTTTGAGACAC-3¢
5¢-CTAGAGGCGCGCCGATATCCT-3¢
5¢-GCCGATGCAAAGTGCCGATAAACA-3¢
5¢-AGAGCTTGGTTGACGGCAATTTCG-3¢
5¢-CGCCAATATATCCTGTCAAACACT-3¢
5¢-GACAGCCTATTGTGGCAGCC- 3¢
5¢-AGAGCTTGGTTGACGGCAATTTCG-3¢
380
M.D. García-Pedrajas et al.
Fig. 34.1 Maps of selected plasmids for transformation of Ascomycota. (a) Map of DelsGate vector pDONR-A-Hyg.
(b) Map of OSCAR binary (pOSCAR) and selection (pA-Hyg-OSCAR) plasmids
primers DonrF-C or DonrF-H (for donor vectors harboring carboxin and hygromycin as
selectable markers, respectively) and DonrR
(see Table 34.1).
2. Primers to verify OSCAR constructs: OSC-F
and Hyg-R(210), and Hyg-F(850) and OSC-R
(see Table 34.1).
3. Restriction enzymes KpnI and HindIII (New
England Biolabs, Ipswich, MA).
Preparation of DelsGate Deletion
Constructs for Fungal ProtoplastMediated Transformation
1. QIAprep spin miniprep kit (QIAGEN).
2. Restriction enzyme I-SceI (New England
Biolabs).
Protoplast-Mediated Fungal
Transformation
1. SCS buffer: 20 mM sodium citrate pH 5.8,
1 M sorbitol. For 200 mL: dissolve 1.18 g
Na3-citrate and 36.44 g sorbitol (SigmaAldrich) in approximately 180 mL dH2O,
bring volume to 200 mL and autoclave. Store
at 4°C.
2. STC buffer: 10 mM Tris–HCl pH 7.5, 100 mM
CaCl2, 1 M sorbitol. For 200 mL: dissolve
36.44 g sorbitol in approximately 160 mL
dH2O, bring volume to 178 mL and autoclave,
then add 2 mL sterile 1 M Tris–HCl (pH 7.5)
and 20 mL sterile 1 M CaC12. Store at 4°C.
3. Buffer II: 5 mM Tris–HCl, pH 7.5, 25 mM
CaC12, 1 M sorbitol. For 100 mL: dissolve
18.22 g sorbitol in approximately 80 mL dH2O,
34
4.
5.
6.
7.
Rapid Deletion Plasmid Construction Methods for Protoplast and Agrobacterium-based…
bring volume to 95 mL and autoclave. Then
add 2.5 mL sterile 1 M Tris–HCl (pH 7.5) and
2.5 mL sterile 1 M CaC12. Store at 4°C.
Lallzyme MMX solution: 500 mg/mL in
Buffer II. For 10 mL: dissolve 5 g of Lallzyme
MMX (standard activities: 1840 poly-galacturonase units/g, 24 pectin lyase units/g and 545
pectin esterase units/g) (Lallemand) in a final
volume of 10 mL Buffer II by gently pipetting
up and down, use fresh or dispense in aliquots,
and store at −80°C. Thaw at RT upon use.
Alternatively Vinoflow® FCE (Novozyme)
can be used instead of Lallzyme MMX to
digest cell walls. Vinoflow solution: 384 mg/
mL in Buffer II. For 10 mL: dissolve 3.84 g of
Vinoflow in a final volume of 10 mL Buffer II,
use fresh or dispense in aliquots, and store at
−80 ºC. Thaw at RT upon use.
40% PEG in STC. For 10 mL: autoclave 4 g
PEG 4000 (Sigma-Aldrich) and 1.82 g sorbitol with 3 mL dH2O (see Note 2), then add
0.1 mL sterile 1 M Tris–HCl (pH 7.5), 1 mL
sterile 1 M CaC12, and sterile dH2O to 10 mL.
Store at 4 ºC and keep on ice when in use.
YEPS with sorbitol (YEPS-S). For 1 L: dissolve 10 g yeast extract, 20 g bactopeptone,
20 g sucrose, and 182.2 g sorbitol in approximately 800 mL of dH2O, bring volume to 1 L,
add 20 g of agar, and autoclave. After autoclaving add 3 mg/mL carboxin (cbx) or 150 mg/
mL hygromycin (hyg) depending on the vector used.
4.
5.
6.
7.
8.
9.
10.
11.
Agrobacterium tumefaciens-mediated
Fungal Transformation
1. Potassium buffer (K-buffer) pH 7.0. For
100 mL: dissolve 20 g K2HPO4 and 14.5 g
KH2PO4 in 80 mL of dH20, bring volume to
100 mL with dH2O. Adjust pH with 10 N
NaOH. Filter sterilize and store at 4 ºC.
2. M-N solution: 3% (w/v) MgSO4.7H2O, 1.5%
(w/v) NaCl. For 100 mL: dissolve 3 g
MgSO4.7H2O and 1.5 g NaCl in 80 mL dH2O,
bring volume to 100 mL with dH2O. Filter
sterilize and store at 4 ºC.
3. Spore elements. For 500 mL: dissolve 50 mg
ZnSO4·7H2O, 50 mg CuSO4·5H2O, 50 mg
12.
13.
381
H3BO3, 50 mg MnSO4·H2O, and 50 mg
Na2MoO4·2H2O in 500 mL dH2O. Filter sterilize, dispense in aliquots, and store at
−20°C.
1 M 2-[N-Morpholino]ethanesulfonic acid
(MES) pH 5.3. For 100 mL: dissolve 21.33 g
of MES in 80 mL dH2O, adjust pH with 10 N
NaOH (see Note 3), bring volume to 100 mL.
Sterilize by autoclaving and store at −20 ºC.
2 M glucose. For 100 mL: dissolve 36.03 g
of glucose in 80 mL dH2O, bring volume to
100 mL. Filter sterilize and store at RT.
50% glycerol. Mix 50 mL of dH2O with
50 mL glycerol. Autoclave and store at RT.
20 mM 3¢,5¢-Dimethoxy-3¢-hydroxyacetophenone acetosyringone. For 10 mL: dissolve 0.039 g of acetosyringone in 10 mL
ethanol. Store at −20 ºC.
200 mM Cefotaxime. Dissolve 0.191 g of
cefotaxime in 2 mL of dH2O. Filter sterilize.
100 mg/mL Moxalactam. Dissolve 0.1 g of
moxalactam in 1 mL of dH20. Filter sterilize.
Minimal Medium (MM). For 100 mL: to
94.15 mL of sterilized dH2O add
1 mL K-buffer (pH 7.0), 2 mL of M-N solution, 0.1 mL of 1% CaCl2.H2O (w/v), 1 mL of
0.01% FeSO4 (w/v), 1 mL of 20% glucose
(w/v), 0.5 mL of spore elements, and 0.25 mL
of 20% NH4NO2 (w/v). Prepare from stock
solutions upon use.
Induction Medium (IM). For 100 mL: to
89.87 mL of sterilized dH2O add
1 mL K-buffer (pH 7.0), 2 mL M-N solution,
0.1 mL of 1% CaCl2.H2O (w/v), 1 mL of
0.01% FeSO4 (w/v), 0.5 mL of spore elements and 0.25 mL of 20% NH4NO2 (w/v),
1 mL 50% glycerol, 4 mL 1 M MES, and
0.5 mL 2 M glucose. Prepare from stock
solutions upon use.
Cocultivation medium plates. This medium
is the same as MM but with 100 mM final
concentration of acetosyringone and 1.5%
agar. For 100 mL: mix 89.87 mL of dH2O
with 1.5 g of agar, autoclave. Then add the
same components as for MM plus 1 mL of
20 mM acetosyringone. Store plates at 4°C.
Selection medium. For 500 mL: mix 19.5 g
Potato Dextrose Agar (PDA, Difco) and 2.5 g
of agar with 500 mL of dH2O and autoclave.
382
M.D. García-Pedrajas et al.
After autoclaving add 50 mg/mL of hygromycin B, 0.5 mL of 200 mM cefotaxime and
0.5 mL of 100 mg/mL moxalactam. Store
plates at 4°C.
Analysis of Fungal Transformants
to Confirm Gene Deletion
1. For DelsGate, gene-specific primer 5 and
CbxF-DG or Hyg-DG (see Table 34.1 and
Fig. 34.2c) for cbx and hyg vectors, respectively. For OSCAR, gene-specific primer 5
and Hyg-R(210) (see Table 34.1 and
Fig. 34.2d). Additionally, it is useful to use
ORF-specific primers to confirm deletion.
2. Alternatively use gene-specific primers 2-O
and 3-O (Fig. 34.2c, d).
Methods
Figures 34.2a and 34.2b outlines the DelsGate
and OSCAR construction steps, respectively.
Figure 34.2c shows a schematic representation of
the entire procedure to generate deletion mutants
using DelsGate: production of deletion constructs
by DelsGate, manipulation of the deletion construct for fungal transformation, and finally analysis of transformants to test for gene replacement.
Figure 34.2d shows a schematic representation of
the entire procedure to generate deletion mutants
using OSCAR: production of deletion constructs
by OSCAR, introduction of the deletion construct
into A. tumefaciens, and analysis of fungal transformants for gene replacement.
DelsGate deletion construction involves the
following three primary steps: (1) simultaneous
independent PCRs of the 5¢ and 3¢ ORF flanks;
(2) Gateway BP cloning; (3) E. coli transformation. A number of features have been included in
the design of our DelsGate method to accelerate
generation of deletion constructs. For the
amplification of the 5¢ flank the recognition
sequence for the homing endonuclease I-SceI is
included at the 5¢ end of the forward primer
(primer 1), while an attB1 sequence is included
at the 5¢ end of the reverse primer (primer 2).
For the 3¢ flank, an attB2 sequence is included at
the 5¢ end of the forward primer (primer 3) and
the I-SceI recognition sequence in the reverse
orientation is included at the 5¢ end of the reverse
primer (primer 4). PCR products are then
inserted into a deletion plasmid vector via the
Invitrogen Gateway BP clonase system. During
the BP clonase reaction the co-purified 5¢ and 3¢
gene flank PCR products recombine with the
attP1 and attP2 sequences of the donor vector,
respectively. This reaction generates a linear
molecule harboring 18 bp of homologous
sequences in opposite orientation at the free
ends (the I-SceI recognition site). After E. coli
transformation, this homologous sequence
recombines in vivo, as reported by Suzuki et al.
[4], generating a circular construct. Selection of
E. coli transformants containing the original
donor vector is prevented by the presence on
these plasmids of the ccdB gene, which is lethal
to DH5a and most other E. coli strains used for
cloning. DelsGate modified donor vectors
pDONR-Cbx and pDONR-Hyg for use with U.
maydis and pDONR-A-Hyg for use in
Ascomycete fungi have been produced by addition of appropriate selectable markers. Therefore,
the entry clone resulting from the BP clonase
reaction and in vivo recombination is the final
deletion construct itself, which has the gene precisely replaced by the vector containing the
selectable marker for fungal transformation and
both flanks separated by the I-SceI recognition
site. Thus, in addition to promoting homologous
recombination to generate the circular molecule
in vivo, this 18 bp sequence is then used to generate the linear DNA for fungal transformation.
The I-SceI site is extremely rare; it does not
exist in the U. maydis genome and is likely completely absent from most fungal genomes. The
deletion constructs generated with this method
are compatible with PMT; we describe here the
formation and transformation of protoplasts of
the basidiomycete fungus U. maydis with
DelsGate deletion constructs, and finally the
analysis of fungal transformants for gene deletion. However, in addition to its extensive use
with U. maydis DelsGate has to our knowledge
been successfully applied to Cryptococcus
34
Rapid Deletion Plasmid Construction Methods for Protoplast and Agrobacterium-based…
383
Fig. 34.2 DelsGate and OSCAR deletion methodologies. (a) Flowchart of steps in the DelsGate method. (b)
Flowchart of steps in the OSCAR method. (c) Schematic
representation of DelsGate deletion construction method,
and generation of deletion mutants using DelsGate con-
structs and PMT. (d) Schematic representation of OSCAR
deletion construction method, and generation of deletion
mutants using OSCAR constructs and A. tumefaciensmediated transformation
neoformans [6], Fusarium verticillioides,1 and
Colletotrichum graminicola.2
OSCAR deletion construction is also based
on Gateway technology and similarly to DelsGate
involves three primary steps: (1) simultaneous
independent PCRs of the 5¢ and 3¢ ORF flanks;
(2) a single Gateway BP cloning step; (3) E. coli
transformation. For OSCAR we have specifically
adopted the attB sites as described in the
MultiSite Gateway system. Thus, during PCR,
recombination sites attB2r and attB1r and
recombination sites attB4 and attB3 are introduced at the ends of the 5¢ and 3¢ gene flanks,
respectively. To simplify the method so that only
a single cloning step is required to generate a
structure in which the 5¢ and 3¢ target gene flanks
are separated by the hygR marker all placed
between the right and left T-DNA borders of the
1
2
A. Glenn, personal communication.
S. Sukno, personal communication.
384
M.D. García-Pedrajas et al.
Fig. 34.2 (continued)
binary plasmids, two new vectors were developed. The first vector developed is a binary vector suitable for ATMT in fungi that harbors the
toxic gene ccdB flanked by the recombination
sites attP2r and attP3, between the left and right
T-DNA borders. We named this vector pOSCAR.
The second vector contains a hygR marker driven
by the Aspergillus nidulans trpC promoter, suitable for selection of transformants in Ascomycete
fungi, flanked by the recombination sites attP1r
and attP4 in pBluescript II KS(+). This is the
marker vector, here named pA-Hyg-OSCAR.
When the PCR products are incubated with
pOSCAR and pA-Hyg-OSCAR in the presence
of BP clonase, each attB site should recombine
only with its single compatible attP site
(e.g., attB2r with attP2r) such that the favored
plasmid structure generated is the deletion construct itself. Transformation of E. coli DH5a, or
similar strain, and selection for spectinomycin
resistance prevent selection of the original
pOSCAR and pA-Hyg-OSCAR vectors since
the former contains the ccdB gene and the latter
confers ampicillin and not spectinomycin
resistance. Deletion constructs produced are then
transformed into A. tumefaciens in preparation
34
Rapid Deletion Plasmid Construction Methods for Protoplast and Agrobacterium-based…
385
Fig. 34.2 (continued)
for fungal transformation. We describe here the
ATMT of Verticillium dahliae and analysis of
fungal transformants for gene deletion.
Primer Design for the Amplification
of Gene Flanks
1. Primers are designed to separately amplify
1 kb of the 5¢ and 3¢ sequences flanking the
ORF of the gene of interest. Specific sequences
are added to the 5¢ end of each primer.
2. Primers 1 and 2 are designed to amplify the 5¢
flank and for DelsGate their 5¢ ends contain
the I-SceI recognition sequence (in the forward orientation) and the attB1 sequence,
respectively (see Table 34.1). For OSCAR, on
the other hand, the 5¢ end of primer 1 contains
the attB2r sequence while the 5¢ end of primer
2 contains the attB1r (see Table 34.1).
The gene-specific sequence of primer 1 (forward primer) is designed for the primer to
anneal approximately 1 kb before the start
codon while primer 2 (reverse primer) is gen-
386
M.D. García-Pedrajas et al.
Fig. 34.2 (continued)
erally designed to anneal as close as practical
before the start codon (see Fig. 34.2c, d).
3. Primers 3 and 4 are designed to amplify the
3¢ flank. For DelsGate their 5¢ ends contain
the attB2 sequence and the I-SceI recognition
sequence (in the reverse orientation), respectively (see Table 34.1). For OSCAR the 5¢
ends of primers 3 and 4 contain the attB4
and attB3 sequences, respectively (see
Table 34.1). The gene-specific sequence of
primer 3 (primer forward) is designed for the
primer to anneal as close as practical after the
termination codon, while the gene-specific
sequence of primer 4 (primer reverse) is generally designed for the primer to anneal
approximately 1 kb distal to primer 3 (see
Fig. 34.2c, d).
4. Gene-specific sequences between 20 and 25
nucleotides work well for these primers.
PCR Amplification of 1 kb Gene Flanks
and Clean Up of PCR Products
1. The 1 kb 5¢ and 3¢ gene flanks are amplified
separately. PCRs are performed in a total volume
34
2.
3.
4.
5.
6.
7.
Rapid Deletion Plasmid Construction Methods for Protoplast and Agrobacterium-based…
of 50 mL. For each of the two reactions combine
in a PCR tube: 32.0 mL PCR-grade ddH2O,
5.0 mL 10× taq-polymerase reaction buffer,
5.0 mL MgCl2 25 mM (see Note 4), 1.0 mL 50×
dNTP mix (10 mM each), 2.5 mL forward primer
20 mM, 2.5 mL reverse primer 20 mM, 1.0 mL
taq-polymerase, 1.0 mL (10–100 ng) fungal
genomic DNA.
Amplification is performed under the
following conditions: an initial denaturation
of 1 min at 94 ºC, 30 cycles of 30 s at 94 ºC,
30 s at 60 ºC, and 1 min at 72 ºC, and completed with a final extension of 5 min at 72 ºC
(see Note 5).
After amplification, load 5 mL of each PCR
reaction on a 0.8% agarose gel to confirm
amplification of the desired bands.
To eliminate primers from the sample prior to
BP clonase reaction, the 5¢ and 3¢ PCR products are co-purified either by PEG/MgCl2 precipitation or by using the QIAquick PCR
purification system (QIAGEN) (see Notes
6 and 7).
For PEG precipitation 90 mL of the combined
PCR products (see Note 8) are mixed with
270 mL of TE buffer (pH 7.5–8) and 180 mL of
30% PEG 8000/30 mM MgCl2, vortexed thoroughly and centrifuged 15 min at 14,000 rpm.
Finally, the pellet is resuspended in 15 mL of
TE buffer or ddH2O.
Purification of the combined 5¢ and 3¢ flanks
with QIAquick columns is performed according to the manufacturer’s instructions (see
Note 8). In the final step DNA is eluted in
30 mL of buffer EB (10 mM Tris-Cl, pH 8.5)
or ddH2O.
After purification of the combined 5¢ and 3¢
flanks by either method DNA concentration is
determined by A260. One mL of the purified
DNA can be directly used for this measurement when using a NanoDrop spectrophotometer. When using a standard spectrophotometer
measure concentration of co-purified PCR
products by mixing 2 mL DNA with 58 mL TE
buffer or ddH2O (Dilution Factor (DF) = 30)
and measuring A260. To calculate concentration: A × DF × 50/1,000 = concentration (mg/mL).
387
BP Clonase Reaction
1. For each DelsGate deletion construct set up
a 5 mL BP clonase reaction, adding the following components in a 0.2-mL microcentrifuge tube at RT: 3 mL (75–125 ng) of
combined purified 5¢ and 3¢ gene flanks, 1 mL
(75 ng) of pDONR vector (see Note 9).
Proceed to step 3.
2. For each OSCAR deletion construct set up a
5 mL BP clonase reaction, adding the following components in a 0.2-mL microcentrifuge
tube at RT: 2 mL (15–20 ng) of combined
purified 5¢ and 3¢ gene flanks, 1 mL (60 ng) of
pA-Hyg-OSCAR vector, 1 mL (60 ng) of
pOSCAR (see Note 9). Proceed to step 3.
3. Thaw BP Clonase II enzyme mix on ice for
about 2 min and vortex briefly twice (2 s each
time).
4. Add 1 mL of BP Clonase II enzyme mix to the
reaction and vortex briefly twice. Spin down
briefly.
5. Incubate overnight (o/n) at 25°C in a PCR
thermocycler with a heated lid (see Note 10).
6. To terminate reaction add 0.5 mL of proteinase
K, vortex briefly, and incubate sample for
10 min at 37°C.
Transformation of E. coli
1. 5 mL of BP clonase DelsGate or OSCAR
reaction is used to transform E. coli competent cells. Generally, transforming homemade
DH5a competent cells with standard heat
shock methods is sufficient to produce a
number of colonies to analyze. However, the
use of commercial One Shot® MAX
Efficiency™ DH5a-T1R, One Shot® Mach1™
T1R, or One Shot® OmniMAX™ 2-T1R E. coli
competent cells (Invitrogen) can be an option
to increase transformation frequencies (see
Note 11).
2. After transformation plate aliquots of 200 mL
on low Na kanamycin plates for DelsGate and
low Na spectinomycin plates for OSCAR (see
Note 12), incubate plates o/n at 37 ºC.
388
Verification of Deletion Constructs
1. Prepare a replica LB kanamycin or LB
spectinomycin plate with the transformants to
be analyzed for the presence of the correct
construct. Between 10 and 20 transformants
are routinely analyzed.
2. Plasmid DNA is then purified to confirm construct structure by PCR and/or restriction
enzyme digestion.
3. Inoculate cells from colonies in the replica
plate in 5 mL of LB amended with 50 mg/mL
kanamycin or 50 mg/mL spectinomycin and
grow o/n at 37 ºC and 200 rpm.
4. Purify plasmid DNA using standard plasmid
miniprep alkaline lysis methods (see Note 13).
5. For OSCAR constructs the standardized test
to confirm correct deletion structure initially
involves a double restriction digestion with
KpnI and HindIII.
6. Each reaction is performed in a total volume
of 20 mL that contains: 1–2 mg of plasmid
DNA (see Note 14), 2 mL of reaction buffer,
1 mL (10 units) of KpnI and 1 mL (10 units) of
HindIII, ddH2O to 20 mL. Incubate reactions
for 2 h at 37 ºC.
7. After digestion load 20 mL of each reaction on
a 0.8% agarose gel to test for the presence of a
band of the expected size.
8. A PCR analysis standardized to test for the
presence of the 5¢ and 3¢ gene flanks is performed to confirm construct structure with
both DelsGate and OSCAR methodology. For
DelsGate the two primer combinations used
are: SceI-F and gene-specific primer 2, and
SceI-R and gene-specific primer 3 (see
Fig. 34.2c). Primers SceI-F and SceI-R are
designed from the I-SceI recognition site in
the forward and reverse orientation, respectively (see Table 34.1). Alternatively, this
analysis can be performed using primers from
the vector in substitution for the gene-specific
primers. In this case the primer combinations
used are DonrF-C or DonrF-H (for donor vectors harboring carboxin and hygromycin and
selectable markers, respectively) and DonrR
(see Table 34.1) in combination with the
SceI-F and SceI-R primers, respectively.
M.D. García-Pedrajas et al.
For OSCAR the two primer combinations are:
OSC-F and Hyg-R(210), and Hyg-F(850) and
OSC-R (see Table 34.1 and Fig. 34.2d).
9. Each reaction is performed in a total volume
of 25 mL that contains: 2.5 mL 10× taq-polymerase reaction buffer, 2.5 mL MgCl2 25 mM
(see Note 4), 0.5 mL 50× dNTP mix (10 mM
each), 1.25 mL forward primer 20 mM,
1.25 mL reverse primer 20 mM, 0.5 mL taqpolymerase, approximately 10 ng of plasmid
DNA, PCR-grade ddH2O to 25 mL.
10. For each of the two primer combinations
prepare a master mix containing all the components except the DNA, for all the reactions
you need to perform plus one. For each reaction mix 24 mL of the master mix with 1 mL
of plasmid DNA sample.
11. Amplification is then performed under the
following conditions: an initial denaturation
of 1 min at 94 ºC, 30 cycles of 30 s at 94 ºC,
30 s at 60 ºC, and 1 min at 72 ºC, and completed with a final extension of 5 min at
72 ºC.
12. After PCR load 10 mL of each PCR product
on a 0.8% agarose gel to test for the presence
of a band of the expected size.
Preparation of DelsGate Deletion
Construct for Fungal Transformation
1. After confirmation of proper deletion construct structure, plasmids are digested with
I-SceI for fungal transformation. This generates a linear vector molecule containing the
selectable marker flanked by the 5¢ and 3¢
flanks of the ORF to be deleted.
2. Select one of the colonies confirmed to have
the desirable construct and grow again in 5 mL
of LB amended with 50 mg/mL kanamycin.
3. Purify plasmid DNA using a QIAGEN column following the manufacturer’s manual.
Measure DNA concentration as above
described.
4. For I-SceI digestion, combine in a microcentrifuge tube, 3–5 mg of plasmid DNA, 5 mL of
10× I-SceI buffer, 1 mL (5 units) of I-SceI and
ddH2O to 50 mL. Incubate 2 h at 37 ºC.
34
Rapid Deletion Plasmid Construction Methods for Protoplast and Agrobacterium-based…
5. Generally, this incubation is sufficient to fully
digest the vector DNA; however, it is a good
practice to test if the digestion is completed by
loading 5 mL of the digestion reaction on a
0.8% agarose gel. A single band of about
between 6.8 and 5.5 kb should be visible
depending on the pDONR vector that was
used to make the deletion construct.
6. After digestion is complete precipitate DNA
by adding 5 mL of sodium acetate 3 M (pH
5.2) and 100 mL 95% ethanol. Vortex briefly
and incubate on ice for 15 min. Centrifuge
15 min at 14,000 rpm. Discard supernatant
and wash pellet with 70% ethanol. Dry pellet
and resuspend in 12 mL of ddH2O.
7. Concentration of transforming DNA can then
be measured by A260 as described previously
(see Note 15).
Transformation of A. tumefaciens with
OSCAR Deletion Constructs
1. After confirming correct OSCAR deletion
constructs, these are transformed into A.
tumefaciens AGL-1, following the same
standard heat shock methods used to transform E. coli. Deletion construct plasmid
DNA purified as described in the
sect. Verification of Deletion Constructs by
standard plasmid miniprep alkaline lysis
methods can be used directly for A. tumefaciens transformation. Alternatively, plasmid
DNA can be purified with a QIAGEN column for this purpose.
2. Transformants are selected on low Na LB
spectinomycin plates.
3. The presence of the deletion construct in the
strain resistant to spectinomycin selected to
be used in the fungal transformation may be
confirmed by colony PCR with primer combinations OSC-F and Hyg-R(210), and/or
Hyg-F(850) and OSC-R (see Table 34.1),
used to confirm deletion constructs as
described in the sect. Verification of Deletion
Constructs (see Note 16).
4. For colony PCR, strains to be tested are
replicated on an LB spectinomycin plate and
5.
6.
7.
8.
9.
10.
389
grown o/n to have enough bacterial
biomass.
Take approximately half of the bacterial
biomass from each replicated colony with a
toothpick or a yellow pipette tip and mix it
with 20 mL of water by vortexing in a microcentrifuge tube.
Incubate 10 min at 100 ºC to lyze cells and
then place samples on ice for 2 min.
Spin cells down for a few seconds at maximum speed and take 5 mL of the supernatant
for PCR.
Each reaction is performed in a total volume of
25 mL which contains: 5 mL of plasmid DNA
solution, 2.5 mL 10× taq-polymerase reaction
buffer, 2.5 mL MgCl2 25 mM (see Note 4),
0.5 mL 50× dNTP mix (10 mM each), 1.25 mL
forward primer 20 mM, 1.25 mL reverse primer
20 mM, 0.5 mL taq-polymerase, and PCRgrade ddH2O to 25 mL.
Amplification is performed under the
following conditions: an initial denaturation
of 1 min at 94 ºC, 30 cycles of 30 s at 94 ºC,
30 s at 60 ºC, and 1 min at 72 ºC, and completed with a final extension of 5 min at
72 ºC.
After PCR, load 15 mL of each PCR reaction
on a 0.8% agarose gel to test for the presence
of a band of the expected size.
Protoplast-Mediated Fungal
Transformation
1. For PMT of U. maydis we use a variation of
the method described by Tsukuda et al. [7]
(see Note 17).
2. Inoculate 5 mL of PDB with one isolated colony of U. maydis grown on 2PDA, incubate
o/n at 28 ºC and 250 rpm.
3. Use approximately 100 mL of the o/n culture
to inoculate 100 mL YEPS medium in 500mL flasks.
4. Incubate o/n at 28 ºC and 250 rpm until an
OD600 between 0.6 and 0.8.
5. Spin cells down in two 50-mL conical tubes,
fairly gently approximately 1,100 × g, discard
supernatant (s/n).
390
6. Add 10 mL of SCS to each tube and
resuspend with gentle vortexing. Combine
the two tubes from each strain into one. Bring
the volume to 30 mL with SCS, and spin as
in step 5.
7. Resuspend cells in 1 mL SCS and add 200 mL
Lallzyme MMX solution (500 mg/mL in
Buffer II). Alternatively add 200 mL Vinoflow
solution (384 mg/mL in Buffer II).
8. Incubate with gentle mixing at RT checking
protoplast formation on a microscope every
15 min. It takes around 45 min for a majority
of cells to generate protoplasts when using
Lallzyme solution. With Vinoflow incubation times are shorter, ranging from 10 to
30 min (see Note 18).
9. When a majority of cells have formed protoplasts spin them down for 10 min at 1,000 × g,
discard s/n, resuspend pellet in 1 mL SCS,
and transfer this suspension to a 1.5-mL
microfuge tube.
10. Spin protoplasts down in a microcentrifuge
for 5 min at 1,000 × g at RT (see Note 19),
carefully remove s/n, and resuspend protoplasts in 1 mL SCS.
11. Spin down as above, resuspend in 1 mL STC,
and spin down in the same conditions.
12. Resuspend protoplasts, on ice, in 1 mL of ice
cold STC. Protoplasts are now ready for
transformation; they can be either used fresh
or stored at −80 ºC for future transformations. For freezing protoplasts add filter sterilized DMSO to 7% (70 mL for 1 mL of
protoplast suspension) and dispense in
aliquots.
13. For transformation, in a 1.5-mL microcentrifuge tube combine approximately 1 mg of
transforming linear DNA in a maximum volume of 5 mL (from step 4, in the
sect. Preparation of DelsGate Deletion
Construct for Fungal Transformation), 1 mL
heparin (15 mg/mL in STC; stored at 4 ºC),
and 50 mL of protoplast suspension, mix gently, and incubate on ice for 20 min.
14. Add 500 mL of PEG 4000 solution, mix by
inverting the tube several times or by very
gentle pipetting, and incubate on ice for
30 min.
M.D. García-Pedrajas et al.
15. Add 500 mL STC, mix by inverting the
tubes several times and pellet protoplasts
by centrifuging at 1,000 × g for 5 min (see
Note 20).
16. Aspirate off the s/n carefully and resuspend
protoplasts in 200 mL of STC.
17. Plate cells on YEPS-S plates containing the
appropriate selection (carboxin or hygromycin) and incubate plates at 30 ºC, checking
them every 24 h. Usually transformants start
appearing after 4–5 days.
18. When transformants start to appear transfer
them to 2PDA with the appropriate
selection.
Agrobacterium tumefaciens-Mediated
Fungal Transformation
1. For A. tumefaciens-mediated transformation
of V. dahliae we use the method described by
Mullins et al. [8] adapted for V. dahliae by
Dobinson et al. [9] with minor variations.
2. On day 1, streak the A. tumefaciens strain
harboring the deletion construct on LB supplemented with 50 mg/mL of spectinomycin,
and grow for two days at 28 ºC.
3. On day 2, inoculate 5 mL of YEPS (see Note
21) with fragments of V. dahliae mycelium
from the edge of an actively growing colony,
and incubate culture for 3–4 days at 24 ºC
and 250 rpm, to produce conidia.
4. On day 3, inoculate 5 mL of MM supplemented with 50 mg/mL spectinomycin with a
colony of the A. tumefaciens strain containing the deletion construct. Grow at 28 ºC and
250 rpm for 2 days.
5. On day 5, measure O.D. of the A. tumefaciens
culture at 600 nm, harvest cells by centrifugation at 5,500 × g for 10 min, discard supernatant, and resuspend cells in 500 mL of IM.
6. Dilute the A. tumefaciens suspension in 5 mL
IM supplemented with 50 mg/mL
spectinomycin, to a final O.D. of 0.15, and
grow for 6–7 h to an optical density, at
600 nm, of 0.6–0.8 (see Note 22).
7. On day 6, harvest V. dahliae conidia by
filtration through Miracloth (Calbiochem)
34
8.
9.
10.
11.
12.
Rapid Deletion Plasmid Construction Methods for Protoplast and Agrobacterium-based…
and then centrifugation at 5,500 × g for
10 min. Finally, resuspend conidia in sterilized dH2O to a final concentration of 106–107
cells/mL.
In a 1.5-mL microcentrifuge tube mix 100 mL
of spore suspension and 100 mL of A. tumefaciens culture.
Spread mixture on a plate of cocultivation
medium overlaid with a sterile 0.45 mm pore
nitrocellulose membrane. Incubate for 2 days
at RT.
Cut nitrocellulose membrane in strips and
distribute these strips on 2 plates of selection
medium.
When fungal transformants start to appear
(see Note 23) transfer them individually to
2PDA supplemented with 50 mg/mL
hygromycin.
Prepare single spore cultures of each transformant in preparation for analysis to determine absence of the gene of interest.
Analysis of Fungal Transformants
to Confirm Gene Deletion
1. Analysis of fungal transformants for gene deletion has been standardized by performing PCR
with a gene-specific primer located approximately 1.1 kb upstream of the start codon (i.e.,
outside the deletion construct) (gene-specific
primer 5), combined with a primer that anneals
to the selectable marker (see Fig. 34.2c, d). The
marker primers used are: CbxF-DG when using
the U. maydis pDONR-Cbx vector, Hyg-DG
when using DelsGate vectors containing hygR
as selectable marker, and Hyg-R(210) for
OSCAR (see Table 34.1).
2. The reaction for each transformant to be analyzed is performed in a total volume of 25 mL
that contains: 2.5 mL 10× taq-polymerase
reaction buffer, 2.5 mL 25 mM MgCl2 (see
Note 3), 0.5 mL 50× dNTP mix (10 mM each),
1.25 mL 20 mM gene-specific primer, 1.25 mL
selectable marker primer 20 mM, 0.5 mL taqpolymerase, 1.0 mL genomic DNA (10–
100 ng), PCR-grade ddH2O to 25 mL.
391
3. Prepare a master mix containing all the components except the DNA, for all the reactions
you need to perform plus one. For each reaction mix 24 mL of the master mix with 1 mL
of DNA.
4. Amplification is then performed under the
following conditions: an initial denaturation
of 1 min at 94 ºC, 30 cycles of 30 s at 94 ºC,
30 s at 60 ºC, and 90 s at 72 ºC, and completed with a final extension of 5 min at
72 ºC.
5. After PCR, load 10 mL of each PCR product
on a 0.8% agarose gel; only transformants in
which the gene has been replaced by the
deletion construct through homologous
recombination are expected to produce a
band with this primer combination.
6. When using this approach to test for gene
deletion it is good practice to confirm absence
of the gene of interest in those transformants
that generated a band in the above PCR by
designing primers that anneal within the ORF.
Perform this PCR in the manner described in
steps 2, 3, and 4. Use a wild type strain as
positive control in the amplification.
7. Replacement of the gene ORF by the deletion construct through homologous recombination can also be tested using primers with
the same sequence as the gene-specific primers 2 and 3 but in the opposite orientation, we
name these primers 2-O and 3-O (see
Fig. 34.2c, d) (see Note 24).
8. Prepare reactions like in step 2 but substituting gene-specific and marker primers by
primers 2-O and 3-O. Proceed as in step 3.
9. Amplification is then performed under the
following conditions: an initial denaturation
of 1 min at 94 ºC, 40 cycles of 30 s at 94 ºC,
30 s at 60 ºC, and 2–5 min at 72 ºC (see Note
25), and completed with a final extension of
7 min at 72 ºC.
10. After PCR load 18 mL of each PCR product
on a 0.8% agarose gel. In transformants in
which gene replacement has taken place you
should observe absence of the wild type band
and presence only of the deletion construct
band, which ranges in size from about
392
3.4–4.4 kb in DelsGate, depending of which
pDONR vector was used, and approximately
1.5 kb in OSCAR.
11. Further confirmation of gene of interest deletion is carried out by Southern blot
hybridization.
M.D. García-Pedrajas et al.
9.
Notes
1. While the rest of media are generally stored
at 4 ºC, PDB tends to precipitate in cold and
it is stored at RT.
2. This is a very dense solution; to prepare it
mix the PEG 4000, sorbitol, and dH2O and
directly autoclave. After autoclaving a clear
viscous solution is obtained to which the rest
of components are added.
3. Addition of drops of NaOH to increase the
pH helps bring the MES into solution.
4. If the10× taq-polymerase reaction buffer
used already contains the MgCl2, do not add
any extra to the reaction and increase volume
of ddH2O accordingly.
5. As with any PCR, the annealing temperature
may have to be adjusted for your particular
primer combination. However, 60 ºC generally works well for the primer lengths
proposed.
6. Although PCR products can be used directly
for BP clonase reactions, if not eliminated
from the sample, primers can also recombine
with vectors carrying the attP sites and
greatly increase the background of colonies
that do not harbor the desired construct. We
therefore strongly recommend performance
of this clean up step.
7. The PEG/MgCl2 precipitation removes primerdimers and DNA molecules smaller than
300 bp. Purification through QIAquick columns eliminates primers and DNA molecules
smaller than 100 bp. We favor the use of PEG/
MgCl2 precipitation since it is an inexpensive
method and produces very clean DNA.
8. It is important that both flanks are roughly at
the same concentration during the BP clonase reaction; therefore if one of the flanks
was amplified more efficiently it is advised
to mix appropriate volumes of each PCR
10.
11.
12.
13.
14.
15.
16.
17.
product to result in similar molar ratios in the
combined purified sample.
Vector DNA suitable for the BP clonase reaction can be purified using standard methods.
This includes alkaline lysis; however this
DNA cannot be quantified by A260 due to
contamination with RNA and we therefore
prefer to use the QIAprep miniprep method
from QIAGEN or any other “clean” plasmid
purification method.
This incubation time can be reduced to as
short as 1 h. However, we favor overnight
incubations which increase the number of
colonies by five- to tenfold.
Transformation can also be performed by
electroporation; however in our experience
this method increases the background
of colonies that do not have the right
construct.
We have consistently found that selection
works much better on low Na LB than on
standard LB. In standard LB sometimes colonies appear that contain no plasmid.
Any method for purification of plasmid DNA
compatible with digestion with restriction
enzymes can be used. At this stage we do not
use QIAGEN columns since of the plasmids
analyzed, only one with the correct structure
will be selected as the deletion construct for
further work.
Standard plasmid miniprep alkaline lysis
methods do not allow quantification of plasmid DNA by measuring O.D. as the samples
contain RNA. In that case, after extracting
DNA, load an aliquot on an agarose gel and
quantify
approximate
concentration
visually.
We start with 3–5 mg of plasmid DNA to
make sure that after the precipitation step we
have enough DNA for transformation. We
regularly use ³1 mg of linear DNA for each
transformation.
Protocols to extract plasmid DNA do not
work well with A. tumefaciens; thus we have
found that the best way to confirm the presence of deletion construct is by colony PCR.
The DelsGate method can be used with any
organism with an efficient recombination
system. We have used deletion constructs
34
18.
19.
20.
21.
22.
23.
24.
Rapid Deletion Plasmid Construction Methods for Protoplast and Agrobacterium-based…
generated by DelsGate for PMT to produce
deletion mutants in U. maydis and this is the
methodology we describe here. For other
fungi, aspects such as formation of protoplasts
or amount of linear DNA that give efficient
rates of fungal transformation may vary.
Since the sale of Novozyme, previously used
for the formation of protoplasts, was discontinued we have tried several commercial
enzyme mixtures; we have found that Vinoflow
generates protoplasts very efficiently and that
Lallzyme MMX, although requiring longer
incubation times, also generates protoplasts
with high transformation frequencies.
Be certain to use a variable speed centrifuge
and that you have it set at the appropriate setting since centrifugation at higher speed at
this stage will burst protoplasts.
STC is added to decrease the density of the
solution to allow protoplasts to pellet.
Other authors use Complete Medium (CM)
[9]. We have found that both media, YEPS
and CM, are suitable to produce conidia for
ATMT.
Sometimes longer incubations, such as to
grow o/n, are required to reach this O.D.
Transformants are visible growing from the
edges of the strips of nitrocellulose membrane into the agar.
This approach can only be used for DelsGate
when the length of the deletion is different
from that of the vector in the deletion construct. Therefore it is not suitable for genes
of about 4.4 kb when using pDONR-Cbx to
generate the deletion construct, for genes of
about 4.8 kb when using pDONR-Hyg, and
for genes of about 3.5 kb when using
pDONR-A-Hyg. For OSCAR this approach
can only be used when the length of the ORF
is different from the hygR marker, approximately 1.5 kb. However, when it can be
applied we consider this approach desirable
because whether there was homologous or
ectopic integration a positive PCR result is
expected, allowing these events to be distinguished by amplicon length. We therefore
find this approach very accurate in testing for
393
gene deletion by PCR. Its disadvantage is
that larger bands need to be amplified; however, except for large ORFs the expected size
of bands is still within the range that can be
amplified with standard taq-polymerasebased PCR reactions.
25. Because larger bands need to be amplified
with this approach, longer extension times
are used.
References
1. García-Pedrajas MD, Nadal M, Kapa LB, Perlin MH,
Andrews DL, Gold SE (2008) DelsGate, a robust and
rapid gene deletion construction method. Fungal Gen
Biol 45:379–388
2. Paz Z, García-Pedrajas MD, Andrews DL, Klosterman
SJ, Baeza-Montañez L, Gold SE (2011) One Step
Construction of Agrobacterium-Recombination-ready
plasmids (OSCAR), an efficient and robust tool for
ATMT based gene deletion construction in fungi.
Fungal Gen Biol 48:677–684
3. Walhout AJ, Temple GF, Brasch MA, Hartley JL,
Lorson MA, van den Heuvel S et al (2000) GATEWAY
recombination cloning: application to the cloning of
large numbers of open reading frames or ORFeomes.
Methods Enzymol 328:575–592
4. Suzuki Y, Kagawa N, Fujino T, Sumiya T, Andoh T,
Ishikawa K et al (2005) A novel high-throughput (HTP)
cloning strategy for site-directed designed chimeragenesis and mutation using the Gateway cloning system.
Nucleic Acids Res 33:e109
5. Hellens R, Mullineaux P, Klee H (2000) Technical
focus: a guide to Agrobacterium binary Ti vectors.
Trends Plant Sci 5:446–451
6. Kmetzsch L, Staats CC, Simon E, Fonseca FL, Oliveira
DL, Joffe LS et al (2011) The GATA-type transcriptional activator Gat1 regulates nitrogen uptake and
metabolism in the human pathogen Cryptococcus neoformans. Fungal Gen Biol 48:192–199
7. Tsukuda T, Carleton S, Fotheringham S, Holloman
WK (1988) Isolation and characterization of an autonomously replicating sequence from Ustilago maydis.
Mol Cell Biol 8:3703–3709
8. Mullins ED, Chen X, Romaine P, Raina R, Geiser DM,
Kang S (2001) Agrobacterium-mediated transformation of Fusarium oxysporum: an efficient tool for
insertional mutagenesis and gene transfer. Phytopathology 91:173–180
9. Dobinson KF, Grant SJ, Kang S (2004) Cloning and
targeted disruption, via Agrobacterium tumefaciensmediated transformation, of a trypsin protease gene
from the vascular wilt fungus Verticillium dahliae.
Curr Genet 45:104–110
Improved Transformation Method
for Alternaria Brassicicola and Its
Applications
35
Yangrae Cho, Akhil Srivastava, and
Christopher Nguyen
Abstract
Alternaria brassicicola is a filamentous fungus that causes black spot
disease on most plants in the Brassicaceae, including cultivated Brassica
species and weedy Arabidopsis. Since the concept of transformation constructs of linear minimal elements was developed [1], we have optimized
protoplast production by reducing melanin accumulation in fungal mycelium. We have used this method either to delete targeted genes or to insert
exogenous genetic information in almost any location of research interest.
Key Words
Alternaria brassicicola • Transformation • Melanin • Protoplast production
Introduction
The imperfect filamentous fungus, Alternaria
brassicicola (Schwein, Wiltshire), causes black
spot disease on a broad range of cultivated and
weedy members of the Brassicaceae. Of note,
A. brassicicola has been used as an example of a
true necrotrophic fungus in studies with
Arabidopsis. Since genome sequences and functional methodologies have been developed for
Y. Cho (*) • A. Srivastava • C. Nguyen
Department of Plant and Environmental Protection
Sciences, University of Hawaii at Manoa,
3190 Maile Way, Honolulu, HI 96822, USA
e-mail: yangrae@hawaii.edu
both the plant1 and its pathogen,2 this has become
an attractive system for the discovering events
that occur at the host–pathogen interface. These
events ultimately determine the outcome of the
interaction.
Knocking out gene functions either by targeted gene disruption or gene replacement [1–5]
has been very helpful for targeted mutational
analysis. The targeted gene approach will be
especially powerful when combined with large
numbers of expressed sequence tags using rapidly evolving technologies. They include serial
analysis of gene expression (SAGE) [6], cap
analysis of gene expression (CAGE) [7], massively parallel signature sequencing (MPSS) [8],
1
2
http://www.arabidopsis.org/
http://genome.jgi-psf.org/Altbr1/Altbr1.home.html
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_35, © Springer Science+Business Media, LLC 2013
395
396
and RNA-seq [9]. Targeted gene knockout methods have been widely used within the fungal
research community. Investigators have adopted
a similar approach for other filamentous fungi,
including plant pathogens such as Magnaporthe
grisea, Fusarium oxysporum, and Colletotrichum
lagenarium [10–12]. Although there were previous reports on the transformation of Alternaria
species, including A. brassicicola [13–15], studying gene functions by targeted gene knockout has
remained difficult owing to the low efficiency of
both transformation and targeted integration. To
improve targeted gene disruption efficiency and
expedite production of gene disruption constructs, a short linear construct with minimal elements that contains an antibiotic resistance
selectable marker gene and a 250–600 bp-long
partial target gene, were developed and successfully used with an almost 100 % targeted gene
efficiency [1]. The production of protoplasts,
however, continued to be a limiting factor in high
throughput transformation.
During the last several years we have optimized a method of producing large numbers of
protoplasts. We used the protoplasts to detect
pathogenicity associated genes among 200 transcription factor-coding genes. We modified protoplast production by inoculating an artificial
medium with hyphae instead of conidia and then
growing the mycelium under conditions that
reduced melanin production.
Melanin is a ubiquitous pigment that plays an
important role in protecting fungi from the damaging effects of environmental stress. It is produced by and accumulates in the cell walls of
hyphae and conidia during the late stationary
phase of mycelial growth. Melanin also forms
under stressful conditions, including ultraviolet
irradiation, a hyperosmotic environment, nutrient
deficiency, or an accumulation of toxic wastes
during in vitro culture. Melanin increases the tolerance of fungi to UV irradiation [16–18], enzymatic lysis [19], and extreme temperatures
[17–20]. Previously, we grew mycelia for 7 days
to acquire an adequate biomass. After 7 days the
mycelia were moderately melanized and digesting them for 4 h produced about ten million protoplasts. With our new method, we inoculate a
nutrient broth with hyphae instead of conidia.
Y. Cho et al.
After only 2 days in culture and one change of
medium at 24 h, melanin accumulation in the
A. brassicicola mycelium is dramatically reduced.
We then digest the mycelium with the same
amount of enzyme for 1–1.5 h. With this method,
we can produce up to 200 million protoplasts and
use them to transform up to 20 constructs.
We have observed an increase in transformation and targeted gene efficiency using protoplasts from mycelia with reduced melanin. We
speculate that reducing melanin accumulation
during mycelial growth was an important factor
in this increased efficiency. We use this method
of protoplast production not only for the transformation of linear minimal element constructs for
gene disruption, but also for linear gene deletion
constructs with a selectable marker gene flanked
on both sides by a targeted gene sequence. We
also use the protoplasts to transform various constructs that are designed to tag targeted promoters
or targeted coding sequences. They can be used
to investigate the spatial and temporal expression
patterns of the gene or the localization of the
tagged proteins, or to purify tagged proteins
(Fig. 35.1).
Materials
The following solutions need to be prepared several days before the transformation procedures.
1. STC buffer: 1 M Sorbitol (54.615 g), 50 mM
Tris (15 mL of 1 M Tris, pH 8.0), 50 mM
CaCl2 (1.66485 g) in 300 mL distilled water.
Stored at 4 °C, it can be used for more than
1 year.
2. 40 % PEG in STC: Dissolve 40 g of PEG 3500
in STC with a final volume of 100 mL. Stored
at 4 °C, it can be used for up to 1 year.
3. Molten regeneration medium: 0.5 % yeast
extract (1.5 g), 0.5 % casamino acid (1.5 g),
1 M sucrose (102.6 g), 0.8 % agar (2.4 g) in
300 mL distilled water. Autoclave to liquefy
the medium. Cool to about 40–45 °C to pour.
Medium stored at room temperature can be
used for several months.
4. 0.7 M NaCl: 140 mL of 5 M NaCl + 860 mL
distilled water. Autoclave. Stored at 4 °C, it
can be used for more than 1 year.
35
Improved Transformation Method for Alternaria Brassicicola and Its Applications
397
Fig. 35.1 Confocal microscopy showing expression of a
targeted gene tagged with green fluorescent protein (GFP).
The green fluorescence appears white in this image. (a)
Hyphae (upper right corner) invading a host plant leaf.
The GFP gene was inserted at the end of a targeted promoter by replacing the targeted gene. The expression of
the GFP gene is controlled by the targeted gene promoter.
(b) Bright dots represent the nuclei in hyphal cells. The
GFP protein gene was fused to the end of the targeted
gene coding sequence. The targeted gene was expressed
as a fusion protein along with the GFP protein. The targeted gene expression and its protein localization were
examined in real time
5. PDA with 30 ng/mL Hygromycin B: Add
19.5 g of Potato Dextrose Agar powder to
500 mL distilled water and autoclave.
6. 1 % Glucose and 0.5 % yeast extract broth in
250 mL flasks.
7. DNA constructs to transform. 5–10 mg of
DNA constructs in 10 mL water in each 1.5 mL
aliquot. Stored constructs at −20 °C and put
on ice before the procedure.
The following equipment and materials are needed:
1. Kitalase (Wako cat #114-00373)
2. Shaker/incubator
3. Centrifuge (Eppendorf 5180R or equivalent)
4. Water bath
5. An incubator or a second water bath
6. Hemocytometer
7. Sterile 50-mL tubes and 12 or 15 mL tubes
with screw caps
8. Tube holder(s) for 1.5, 12, and 50 mL-sized
tubes
9. Miracloth (Calbiochem cat #475855)
10. 0.2 mm Millex-FG syringe-driven filter
(Millipore, Carritowhill, Ireland)
11. 10 mL Syringe
12. Funnel
13. Sterilized 250-mL flasks with aluminum foil
covers
14. Scalpel
15. Ice box and ice
16. Petri plates
Methods
The following method is written for people with
little laboratory experience. It may be used to
train new students, but it may be too detailed for
experienced researchers. This method can be used
by the latter, however, as a guideline for making
simplified protocol. This method was also tested
with Alternaria alternata and Maganporthe grisea to increase the efficiency of protoplast production and transformation. Note: Contamination
of the environment should be avoided. Do all
work with conidia in a biological safety cabinet.
Always perform experiments using proper sterilization techniques. For example, avoid touching
the openings of flasks, tubes, and caps with your
fingers; flame the rims of flask or tube openings
before opening and closing them.
398
Fungal Culture
1. Sterilize the scalpel blade by dipping it in
ethanol and then igniting it with a flame. Put
the scalpel to the side and allow it to cool.
2. Remove the lid of a Petri plate containing
sporulating A. brassicicola. Before actually
touching the fungus, insert the scalpel blade
into the agar near the edge of the plate. This
will further cool the blade.
3. Gently scrape the conidia toward the edges
of the plate, being careful not to disturb
the agar. You want the hyphae growing in
the agar for protoplast production, not the
conidia.
4. After clearing conidia to the side, collect
only hyphae by gently scraping, not slicing
the hyphae-containing agar from the surface.
Note: Each piece of agar will serve as the
core of a mycelia ball. If the agar pieces are
small, there will be many small mycelia
balls; and if the agar pieces are large, there
will be only a few large mycelia balls. Harvest
enough mycelium-impregnated agar to produce 3–5 mL of fungal biomass after washing with NaCl (see Transformation, step 8).
5. Transfer the agar pieces containing mycelium into a 250-mL flask containing 50 mL
of 1 % glucose and 0.5 % yeast extract broth
(GYEB). Shake the scalpel to release the
agar into the broth. Flame the scalpel before
each transfer of agar and hyphae to the flask.
6. Lightly swirl the flask, flame the rim of the
opening, and then seal it with foil. Flaming
the rim of the flask opening and aluminum
cover will help prevent contamination.
7. Label the foil cover with the sample name,
type of medium, date, and time.
8. Incubate the flasks in the shaker/incubator at
25 °C and 100 rpm in the dark.
9. Let the fungal mycelium grow for about
36–48 h.
10. Optional: the day before the transformation
procedure, replace the old GYEB with fresh
GYEB and continue to incubate the culture.
11. After incubation the broth should contain
separate, spiky, milky white mycelia balls.
A grey color to the mycelia or conidia is a
Y. Cho et al.
sign of melanin accumulation, which inhibits
enzyme digestion, as described in the following section.
Transformation
1. Turn on the water bath and set it at 42 °C.
Gather the necessary materials and turn on
the biosafety cabinet.
2. Place the molten regeneration medium in the
40–45 °C water bath or incubator.
3. Put the prepared 0.7 M NaCl in the ice box to
keep it cold.
4. Remove the incubating mycelium from the
shaker/incubator and pour it into a sterilized
50 mL falcon tube. Centrifuge at 3,600 rpm
for 5 min.
5. While the mycelium is being centrifuged,
prepare 10 mg/mL Kitalase. Measure 100 mg
of the Kitalase powder and pour it into a sterilized 12 mL polypropylene tube; add 0.7 M
NaCl to make 10 mL. Place the tube in the
shaker/incubator until the contents are completely dissolved.
6. For the mycelium harvested from the 50 mL
falcon tube, use 10 mL of Kitalase solution.
7. Pour off the GYEB medium and fill the tube
with 0.7 M NaCl. Centrifuge at 3,600 rpm
for 5 min and then decant the NaCl solution.
8. Continue washing the mycelium by repeating step 6. We prefer to have 3–5 mL of fungal biomass when the two washings with
NaCl are finished.
9. Get a sterile 250-mL flask, 20-mm pore size
filter, and a syringe kit and prepare to filtersterilize the Kitalase solution as it is added to
the culture sample. Do not touch the filter
with your hands or allow anything else to
touch it. Grip it through the plastic wrap.
10. After washing the culture sample with NaCl
(step 8), open the syringe and pull out the
plunger, attach the filter to the front of the
syringe, and pour the Kitalase into the
syringe. Reassemble the syringe, then apply
pressure to the plunger to force the Kitalase
through the filter and into the washed
mycelium.
35
Improved Transformation Method for Alternaria Brassicicola and Its Applications
399
Fig. 35.2 Protoplasts produced by enzyme digestion.
(a) Protoplasts before purification. Digestion is complete
and the protoplasts in the digestion solution are clean
enough to be used for transformation after precipitation
by centrifugation. Floating cell wall debris (arrow) does
not inhibit transformation. (b) Protoplasts after
purification. The size of protoplasts varies from less than
10 to almost 50 mm in diameter
11. Pour the washed mycelium and Kitalase into
a sterile 250-mL flask. Flame the mouth of the
flask and the foil, then cover. Place the flask in
the shaker/incubator and set the temperature
at 28 °C. Digestion of the cell walls of the
mycelium will be complete within 90 min.
Check the progress of the digestion after 1 h
by removing a 10-mL sample from the flask
and viewing it under a compound microscope.
Undigested hyphae look like long, smooth
hairs. Partially digested hyphae look like
strings of sausages. Completely digested
hyphae will normally float like cotton balls
and the protoplasts are well separated from
the floating cell-wall debris (Fig. 35.2).
12. During the digestion process, pour 50 mL of
molten regeneration medium into each 50 mL
tube and incubate in the 40–45 °C water bath
or incubator. The number of 50-mL tubes
needed depends on the number of samples
being prepared for transformation.
13. Once the fungal mycelium has been properly
digested, filter the protoplasts through a
miracloth-lined funnel and collect them in a
sterile 12-mL polypropylene tube. Gently
swirl the miracloth funnel while filtering.
Wash the miracloth funnel with a small volume of 0.7 M NaCl to recover uncollected
protoplasts. Add 0.7 NaCl until the 10 mL
line of the polypropylene tube is reached.
14. Centrifuge the polypropylene tube at 4 °C
and 700 rpm for 10 min. Decant the liquid,
add 3 mL of cold 0.7 M NaCl and resuspend
the protoplasts by gently inverting the tube
back and forth. Ideally, the pellet of protoplasts will remain attached to the bottom of
the tube during the inverting. If the pellet
breaks loose, it may be difficult to completely
resuspend the protoplasts. Gentle inversion
will slowly release the pellet into the liquid.
Add the remaining 0.7 M NaCl until the volume reaches 10 mL.
15. Centrifuge again at 4 °C and 700 rpm for
10 min. Decant the liquid and add 3 mL of
STC buffer and gently invert to mix. When
the pellet is completely resuspended, add
STC to make 10 mL and mix well. Collect
10 mL of the sample and place it in a hemocytometer. Centrifuge again at the same
settings to precipitate the protoplasts.
400
16. Count the number of protoplasts with the
hemocytometer. Five to ten million protoplasts will be needed for each transformation
construct; more is better than less.
17. When this centrifugation is done, decant the
liquid and add STC to make a concentration
of ~10 million protoplasts in 70 mL.
18. Add the 70 mL of protoplasts using 1-mL
pipette to the DNA transformation constructs
in 1.5-mL aliquots. Pipetting with a smaller
pipette may damage protoplasts. Smaller
pipettes can be used after cutting off the
pipette tips to make the opening wider.
Place the aliquots on ice for 10 min. Move
the ice box next to the water bath so aliquots
can be quickly transferred from the ice to the
water bath. Place the aliquot holder in the ice
box and insert the aliquots as they are filled.
Move the aliquot holder and aliquots to the
42 °C water bath and incubate for 2–10 min.
19. Remove the aliquots from the water bath and
quickly put them back on ice for 10 min.
Collect the Petri plates that will be needed.
20. Add 40 % PEG in STC at room temperature
to the aliquots. Mix by gently inverting
each tube. Incubate for 15 min at room
temperature.
21. Transfer 400 mL of the sample to a 50-mL
tube of molten regeneration medium.
Thoroughly mix the protoplasts and PEG
into the molten regeneration medium.
22. Pour each 50-mL tube of molten regeneration medium equally into two 100 × 15 mm
Petri plates.
23. Incubate these plates in an incubator at 25 °C
for 16–24 h.
24. Overlay the plates with 25 mL of PDA with
Hygromycin B (30 ng/mL).
25. Transformants will begin to appear in 5–7
days and continue to emerge for about 2
weeks (Fig. 35.3). They are ready for singlespore purifications as they emerge from the
selective medium. Cross-contamination of
the transformation plates is rare as long as
the plates are kept undisturbed.
Y. Cho et al.
Fig. 35.3 Transformation plate 4 days after overlaying it
with the selective medium. The arrowhead, arrows, and
circles mark emerged transformants, growing transformants in the selective medium, and putative transformants,
respectively, that may grow within a few days
Acknowledgements We thank Lindsay Oxalis for assisting with the research and Dr. Fred Brooks for his critical
review of the manuscript. This research was supported by
USDA-TSTAR 2009-34135-20197 and HATCH funds to
Yangrae Cho, administered by the College of Tropical
Agriculture and Human Resources, University of Hawaii
at Manoa, Honolulu, HI.
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2. Adachi K, Nelson GH, Peoples KA, Frank SA,
Montenegro-Chamorro MV, DeZwaan TM et al (2002)
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fungi. Proc Natl Acad Sci U S A 98:5110–5115
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Improved Transformation Method for Alternaria Brassicicola and Its Applications
4. Lo C, Adachi K, Shuster JR, Hamer JE, Hamer L
(2003) The bacterial transposon Tn7 causes premature
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TAGKO mutagenesis in filamentous fungi. Nucleic
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Souza CP et al (2004) Rapid production of gene
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fluorescent protein-tagged centromeric marker in
Aspergillus nidulans. Eukaryot Cell 3:1359–1362
6. Velculescu VE, Zhang L, Vogelstein B, Kinzler KW
(1995) Serial analysis of gene expression. Science
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8. Brenner S, Johnson M, Bridgham J, Golda G, Lloyd
DH, Johnson D et al (2000) Gene expression analysis
by massively parallel signature sequencing (MPSS) on
microbead arrays. Nat Biotechnol 18:630–634
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revolutionary tool for transcriptomics. Nat Rev Genet
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Methods for High-Quality DNA
Extraction from Fungi
36
Vijai Kumar Gupta, Maria G. Tuohy, and Rajeeva Gaur
Abstract
Because of breakage of rigid fungal cell walls, the major challenge for
DNA isolation from fungi is to obtain samples of good quality and quantity. We developed a fast and efficient DNA isolation method from fungi
that was successfully applied to Fusarium spp., Colletotrichum spp.,
Trichoderma spp., Gliocladium roseum, and Lasiodiplodia theobromae.
Keywords
DNA isolation • Fungi • Fusarium spp. • Colletotrichum spp. • Trichoderma
spp. • Gliocladium roseum • Lasiodiplodia theobromae
Introduction
V.K. Gupta (*)
Molecular Glycobiotechnology Group,
Department of Biochemistry,
School of Natural Sciences,
National University of Ireland Galway,
University Road, Galway, Ireland
Assistant Professor of Biotechnology, Department
of Science, Faculty of Arts, Science & Commerce,
MITS University, Rajasthan, India
e-mail: vijai.gupta@nuigalway.ie; vijaifzd@gmail.com
M.G. Tuohy
Molecular Glycobiotechnology Group, Department
of Biochemistry, School of Natural Sciences,
National University of Ireland Galway,
University Road, Galway, Ireland
R. Gaur
Department of Microbiology, Dr. R.M.L. Avadh
University, Faizabad, Uttar Pradesh, 224001, India
Fungal biology has become important in relation to
the study of evolutionary biology and in the industrial process. Consequently, there is a need to investigate the molecular genetics of this fungus. A wide
range of molecular manipulation techniques of
fungi are currently available, including gene disruption, various PCR applications (random
amplified polymorphic DNA analysis, microsatellite typing, etc.), and DNA-based epidemiological
studies (restriction fragment length polymorphism
analysis, fingerprinting, etc.). [1–3] Each of these
techniques requires the recovery of good-quality
genomic DNA. Most DNA extraction protocols for
fungi are based on mechanical isolation methods
that employ grinding mycelia after freezing them
in liquid nitrogen or glass bead disruption, followed
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_36, © Springer Science+Business Media, LLC 2013
403
404
V.K. Gupta et al.
by additional purification steps. Although the
quantity and quality of DNA obtained by these
methods are generally satisfactory, the techniques
are time-consuming and, therefore, not suitable for
analysis of a large number of samples. Consequently,
new protocols have been developed in order to
reduce the time required for DNA isolation. [4, 5]
However, most of these protocols involve extractions using hazardous chemicals such as phenol,
chloroform, and isoamyl alcohol and are, therefore,
not a preferred option.
In this chapter, we investigate a simple method
for isolation of good-quality DNA from fungi.
This method is convenient and easy to use, does
not require physical methods, and works directly
with mycelia.
Materials
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
1 M Tris HCl
0.5 M Na2 EDTA (pH-8.0)
1 M NaCl
0.5 % SDS
25 mM EDTA
200 mM Tris HCl
2.5 M Sodium acetate (pH 5.2)
10x TAE buffer
Extraction buffer: Prepared using 200 mM
Tris HCl, 20 mM Nacl, 25 mM ethylenediaminetetraacetic acid (EDTA) and 1 %
sodium dodecyl sulfate (SDS) (pH 8.5)
Saturated isopropanol
70 % alcohol
Tris-EDTA (TE): Prepared by using 10 mM
Tris and 1 mM EDTA, pH 8.0
6 × gel loading dye: Prepared using 25 mg
bromophenol blue, 25 mg xylene cyanol,
3 mL glycerol in 6.75 mL DW (for 10 mL of
6 × loading dye). The dye was then autoclaved and stored at 4 °C
Ethidium bromide: 10 mg/mL stock
Methods
The methods given as follows describe general
procedures for obtaining DNA from different
fungal sources. The volumes and number of tubes
used per sample may need to be varied, depending
on the type of sample and size. This method was
employed to prepare DNA from Fusarium spp.,
including those of Colletotrichum spp.,
Trichoderma spp., Gliocladium roseum, and
Lasiodiplodia theobromae. [6–8]
1. Grow fungal cultures in suitable broth for
72 h at 28 ± 2 °C on a shaker at 120 rpm.
2. Decant the sample containing growth media
including fungal growth in small centrifugal
tubes as per size of the experiment. Pellet the
sample by centrifugation at high speed
(13,000 rpm) for 5 min. Wash with sterile
distilled water for 5 min for two times at
same speed (see Note 1).
3. Overdry the pelleted sample at 65 °C for
30 min to remove excess liquid (see Note 2).
4. Homogenize the required amount of the pellet in an Eppendorf tube/sterile tube for 5 min
in a suitable ratio of fungal mycelia sample
and extraction buffer (1:6), respectively (i.e.,
for our experiments we used 50 mg of mycelia with 300 mL of extraction buffer in an
Eppendorf tube) (see Note 3).
5. Add 200 mL of 2.5 M sodium acetate, vortex
the mixture, and cool the mixture at −20 °C
for 10–15 min.
6. Centrifuge for 5 min at 13,000 rpm and transfer the supernatant in fresh Eppendorf tubes
and then add equal volume of chilled saturated isopropanol. If the DNA is in good
quantity, spool out the DNA into a fresh tube
and follow step 8 (see Notes 4 and 5).
7. Centrifuge for 5 min at 13,000 rpm to pellet
the DNA.
8. Supernatant was discarded without interfering with the DNA pellet.
9. Rinse the DNA pellet twice with 500 mL of
70 % ethanol for 5 min at 13,000 rpm (see
Note 5).
10. DNA pellet was air dried in an aseptic condition for 1.5 h to remove traces of ethanol and
was subsequently dissolved in 100 mL TE
buffer.
11. The Eppendorf tubes were kept in water bath
at 55 °C for 4 h to inhibit DNAse activity and
to resuspend the pellet completely in TE buffer or allow the pellets to resuspend overnight at 4 °C.
36
Methods for High-Quality DNA Extraction from Fungi
12. After 4 h of resuspension of DNA in TE buffer
at 55 °C in water bath or overnight resuspension at 4 °C, the TE-dissolved DNA was
cooled and stored at −20 °C for further use.
13. Purify the DNA using DNA purification kits.
We found PowerClean® DNA Clean-Up Kit
(Mo Bio Laboratories, USA) to be very
effective for purifying DNA (see Note 6).
14. To determine the approximate concentration
of the DNA, take 1 mL of DNA solution and
dilute it by adding 999 mL of Milli-Q water.
Mix well and measure absorbance values at
260 nm and 280 nm in a UV spectrophotometer. For good-quality DNA the ratio is 1–1.8,
but OD is not useful in the presence of any
contaminants. Therefore, run 10 mL genomic
(uncut) DNA using 6 × gel loading dye on a
0.8 % agarose gel with ethidium bromide.
The amount of fluorescence compared to the
l(lambda) DNA markers will provide a
rough estimate of the DNA concentration
(see Notes 7 and 8).
405
7. The ethidium bromide acts as an intercalating
agent. It has four cyclic rings similar to purine
and pyrimidine nitrogen bases of the nucleotides. After the addition of ethidium bromide
to the gel and loading of the gel was over, the
ethidium bromide slipped in between the adjacent base pairs of the DNA double helix. This
intercalated ethidium bromide showed
fluorescence in the presence of the UV light.
The density of ethidium bromide is directly
proportional to the concentration of DNA present in the given volume of sample loaded.
Concentration of ethidium bromide prepared is
10 mg/mL, and it is autoclaved after preparation. Ethidium bromide helps us to analyze the
position and concentration of DNA in the gel).
8. Note: Because l(lambda) markers are the
known molecular weight strands from
l(lambda) phage DNA, by using these markers one can quantitate the molecular weight of
the DNA to be quantified. This is a very
sophisticated and accurate method for quantitation of a genomic DNA as compared to UV
spectroscopy.
Notes
1. Washing fungal mycelia with sterile distilled
water is necessary to remove excess metabolites attached with mycelia.
2. DNA pellets should not be overdried because
this may cause a problem in dissolving the
DNA in TE buffer. Undissolved residues
should be removed using sterile pipette tips.
3. Completely mixing the pelleted mycelia sample with extraction buffer into a homogenate
increases the efficiency of DNA extraction.
4. If the DNA does not spool out, do not discard;
centrifugation helps to recover the DNA.
5. 70 % alcohol should be freshly prepared. Both
saturated isopropanol and 70 % alcohol should
be used after being chilled.
6. If the extracted DNA is not sufficiently pure
(i.e., not suitable to restriction enzyme digestion and/or PCR amplification), repeat the
saturated isopropanol extraction. Alternatively,
use any standard DNA purification kit to
purify DNA, following the manufacturer’s
instructions.
Acknowledgements The authors are very thankful to
Head, Department of Microbiology, R. M. L. Avadh
University, Faizabad, UP, India, and Prof. Shakti Baijal,
Ex-Dean, FASC, MITS University, Rajasthan, for providing the necessary research grants.
References
1. Raeder U, Broda P (1985) Rapid preparation of DNA
from filamentous fungi. Appl Microbiol 1:17–20
2. Cubeta MA, Echandi E, Abernethy T, Vilgalys R
(1991) Characterization of anastomosis groups of
binucleate Rhizoctonia species using restriction analysis of an amplified ribosomal RNA. Phytopathology
81:1395–1400
3. Clarke DL, Woodlee GL, McClelland CM, Seymour
TS, Wickes BL (2001) The Cryptococcus neoformans
STE11alpha gene is similar to other fungal mitogenactivated protein kinase kinase kinase (MAPKKK)
genes but is mating type specific. Mol Microbiol
40:200–213
4. Graham GC, Mayers P, Henry RJ (1994) A simplified
method for the preparation of fungal genomic DNA for
PCR and RAPD analysis. Biotechniques 16:48–50
5. Min J, Arganoza MT, Ohrenberger J, Xu C, Akins RA
(1995) Alternative methods of preparing whole-cell
406
DNA from fungi for dot-blot, restriction analysis, and
colony filter hybridization. Anal Biochem 225:94–100
6. Gupta VK (2009) Molecular characterization of
Fusarium wilt pathogens of guava (Psidium guajava
L.) using RAPD and microsatellite maker. Ph.D.
Thesis. Dr. R.M.L. Avadh University, Faizabad, Uttar
Pradesh, India
V.K. Gupta et al.
7. Gupta VK (2012) Genetic Diversity of Fusarium wilt
pathogens of Guava in India. J Environ Sci Health [B]
47:315–325
8. Gupta VK (2012) Characterization of virulence gene
loci in the genome of Fusarium spp. associated with
wilt disease of guava in India. Arch Phytopathol Plant
Protect (accepted), APPP ID: 45(2):244–259
Production of Recombinant Proteins
from Pichia pastoris: Interfacing
Fermentation and Immobilized
Metal Ion Affinity Chromatography
37
Berend Tolner, Gaurav Bhavsar, Bride Foster,
Kim Vigor, and Kerry Chester
Abstract
The methods describe a Pichia pastoris fermentation system for generation
and purification of recombinant proteins. The proteins are secreted with
hexahistidine tags and purified from feedstock by immobilized metal ion
affinity chromatography (IMAC) using either radial flow or expanded bed
adsorption. IMAC allows for an initial fast capture and isolation step that
omits the need for filtration or centrifugation as primary procedures.
The methods are applicable to production of recombinant protein in the
laboratory and can be adapted to good manufacturing practice (GMP)
compliant processes.
Keywords
P. pastoris • Recombinant • Protein • Purification • His-tag • IMAC
• Radial flow bed • Expanded bed
Introduction
The use of Pichia pastoris as a production
platform for recombinant proteins has been
highly successful on both the laboratory and good
manufacturing practice (GMP) bioprocess scale.
The system can produce heterologous recombinant protein at yields of grams per liter [1–8]. To
date some 600 genes have been cloned and
expressed in P. pastoris [9], and the overall
B. Tolner (*) • G. Bhavsar • B. Foster
K. Vigor • K. Chester
Department of Oncology, University College London
Cancer Institute, Paul O’Gorman Building, 72 Huntley
Street, London, WC1E 6BTUK
e-mail: b.tolner@ucl.ac.uk
production of proteins from this yeast is likely to
increase [10, 11].
Depending on application, yeast production
can be an attractive alternative to mammalian cells
or the commonly used Escherichia coli platform.
Relative to production of recombinant proteins in
mammalian cell lines, yeast expression has a
shorter fermentation time, shows fast progression
from gene synthesis to first-time protein production, makes use of simple chemically defined
media without animal-derived products (exclusion of viral contaminants), and has an overall
reduced operation cost [12]. P. pastoris performs
posttranslational modifications such as disulphide
bonds and glycosylation [3, 4, 5]. However, Pichia
glycoforms are of the high mannose type and
therefore differ from mammalian structures. This
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_37, © Springer Science+Business Media, LLC 2013
407
408
can be problematic for some in vivo therapeutic
purposes because mannosylation facilitates accelerated clearance from circulation; in some cases,
such accelerated clearance is a desirable design
feature [13, 14]. Pichia glycoforms may also be
disadvantageous if the protein required depends on
specific glycosylation for optimal activity. For
example, antibody functions such as antibodydependent cell cytotoxicity (ADCC) depend on
glycosylation [15, 16], and differences relative to
human glycosylation may cause wild-type P. pastoris recombinant proteins not to engage effector
functions. However, advances in development of
glyco-engineered strains [17–20] allow production
of recombinant proteins with mammalian-like glycosylation. The availability of the complete
sequence of the P. pastoris genome [21, 22], as
well as public access to this dataset [23–25], support further strain development and are likely to
result in a toolbox of strains for future applications
with designer glycoforms.
In E. coli, the expression constructs usually
have an episomal location and therefore must be
maintained under selective pressure to ensure
stable plasmid copy number and counteract segregational plasmid instability. Furthermore, structural plasmid stability is critical. In P. pastoris an
expression cassette can be stably integrated into
the chromosome, although this could lead to
clonal variation.
However, relative to E. coli the major advantage of yeast is its capability to perform posttranslational modifications and secretion of
product directly into the media. The latter
simplifies downstream processing because P.
pastoris secretes only very low concentrations of
host cell proteins into the media. Therefore, P.
pastoris fermentation with secretion of recombinant His-tagged protein can be readily interfaced
with expanded bed adsorption–immobilized
metal ion affinity chromatography (EBA-IMAC)
[26, 27] or radial flow bed adsorption IMAC
(RBA-IMAC) and could be advantageous in
bringing new products into the clinic [28]. Indeed,
several His-tagged recombinant proteins are in
clinical applications (Table 37.1). For a listing of
other P. pastoris produced therapeutic proteins
see, for example, Refs. [11, 12].
B. Tolner et al.
The methods described in this chapter have
been used in our laboratory for production of a
range of proteins including single chain Fv antibodies and diabodies [29–32] and N-A1 domains
of carcinoembryonic antigen [33]. The yields
ranged from 4 to 200 mg/L of supernatant. The
quality of proteins generated with this protocol is
sufficient for applications where purity is critical—for example, affinity studies using surface
plasmon resonance [33], biodistribution [13, 14,
34], flow cytometry analysis [13], crystallization
(Fig. 37.1), and nanoparticle functionalization
[28]. Furthermore, the protocol can be readily
adapted to GMP-compliant manufacture [13, 14,
28, 35].
Materials
See Notes 1–3.
Chemicals
1. 70% Ethanol
2. CuSO4·(H20)5
3. H2O: sterile deionized or water for irrigation
(Baxter, UKF7114)
4. Imidazole
5. Na-EDTA
6. NaCl
7. NaOH
8. PBS
9. Virkon
10. pH indicator strips pH 1–14 (VWR
315082P)
11. Solutions: Prepare as per Table 37.2.
Equipment
1. Radial flow column: Proxcys, model AXCIS
MD 62 MKIII; depth 6 cm and volume
125 mL
2. IDA chelating matrix (Sterogen, Cellthru
BigBead resin; 300 mL, catalog no. 37311)
3. Peristaltic pump, capacity up to 300 mL/min
(e.g., Watson Marlow 520S)
37
Production of Recombinant Proteins from Pichia pastoris...
409
Table 37.1 His-tagged proteins in clinical trials
Product names
NY-ESO-1
Company/University
Cornell University
Description/trial status
Early trial
OFA/iLRP &
Quantum
Immunologics
rCR2
University of South
Alabama
Phase I/II
Preclinical testing
Vicinium
Proxinium
MFECP1
Medical Research
Council UK (MRC)
Viventia Biotechnology
Viventia Biotechnology
Royal Free Hospital/
University College
London
OprF-OprI
University of Hannover At least 4 Phase I/II studies
IdioVax
CellGenix
rMAGE-A3
Endostar (or
Endu)
ASCI/GSK
Simcere
Pharmaceutical Group
Phase I/II
Phase I/II/III
Phase I
Phase II & has Orphan Drug
status in EU granted 2004
Phase II/III
Phase I, II, III, IV
Reference
http://www.news.cornell.edu/stories/
Aug09/cancerVaccine.htmL
http://www.ncbi.nlm.nih.gov/
pubmed/21365782
http://quantumimmunologics.us/
About-Quantum-Immunologics.htmL
http://www.ncbi.nlm.nih.gov/pmc/
articles/PMC3071809/
http://www.viventia.com/
http://www.viventia.com/
http://www.ucl.ac.uk/cancer/researchgroups/recombinant-therapeutics/index.
htm
http://clincancerres.aacrjournals.org/
content/12/21/6509.full
http://www.ncbi.nlm.nih.gov/
pubmed/17683588
http://www.old-herborn-university.de/
literature/books/OHUni_book_17_article_7.pdf
http://www.cellgenix.com/en/news/
EN-news-press-2004-Orphan.htmL
http://www.ascitrials.com/
http://www.simcere.com/english2009/
news/News_show.asp?gongs_id=44
Fig. 37.1 Crystals of a single chain Fv (scfv). Protein was isolated using P. pastoris fermentation interfaced with EBAIMAC. Crystals were isolated and images prepared by Dr. Noelia Sainz-Pastor
4. Pump head tubing, ID 9.5 mm (Cole-Parmer
HV-06508-73)
5. Tee connectors, PFA, ½-inch sanitary clamp
fitting (Cole-Parmer, KH-31530-10)
6. Four sanitary to hose barb connection
(Cole-Parmer WZ-30560-10)
7. Four sanitary clamps (Cole-Parmer WZ-3056201)
410
B. Tolner et al.
Table 37.2 Buffers and solutions required
Buffer/solution
Packing buffera
Composition
0.5 M NaCl
0.5× PBS
Storage solution
0.1 M NaOH
0.5 M NaCl
4
29.22
Copper sulphate solutiona
0.1 M CuSO4·(H2O)5
25
Equilibration buffera
0.5 M NaCl
0.5× PBS
10 mM Imidazole
29.22
4.8
0.68
Biomass dilution buffera,b
2 M NaCl
2× PBS
40 mM Imidazole
116.88
19.2
2.7
Matrix wash buffera
0.5 M NaCl
0.5× PBS
40 mM Imidazole
29.22
4.8
2.7
Elution buffera
0.5 M NaCl
0.5× PBS
200 mM Imidazole
29.22
4.8
13.6
EDTA solutiona
50 mM Na-EDTA
18.75
Strip solution
1 M NaCl
1 M NaOH
58.44
40
a
Per liter (g)
29.22
4.8
This protocol (g)
58.44
9.6
2 L H20
4
29.22
1 L H20
50
2 L H20
146.1
24
3.4
5 L H2O
467.52
76.8
10.8
4 L H2O
292.2
48
27
10 L H20
58.44
9.6
27.2
2 L H2O
37.5
2 L H2O
58.44
40
1 L H20
pH of this buffer is not adjusted
Assume harvesting a 10-L bioreactor
b
8. Sanitary gaskets 10/pk (Cole-Parmer
EK-30548-00)
9. Sanitary pressure gauge 0–30 psi (ColeParmer WZ-68056-72)
10. Clear tubing, 1/4″ ID × 3/8″ OD (0.6 cm
ID × 1 cm OD), Tygon ACG00017-CP (ColeParmer RZ-95635-53)
11. Male-pipe-thread-CPC nonvalved (Proxcys
APC24B)
12. Female-in line-CPC barbed fitting nonvalved
(Proxcys APC17)
13. Male-pipe-thread-CPC valved (Proxcys
APCD24B)
14. Female-in line-CPC barbed fitting valved
(Proxcys APCD17)
Spares:
15. 40-mm inlet and outlet frits (Proxcys; PX-FS1AXCIS-MD62-MKII)
16. Inlet and outlet O-rings (Proxcys; PX-UP1AXCIS- MD62-MKII)
17. Five autoclaveable waste containers (e.g.,
Nalgene carboy, 10 L)
18. Five Duran 1-L bottles
19. Five Duran 2-L bottles
20. Stirrer platform plus stirrer bar (suitable for
1- to 2-L bottles)
21. One mixer vessel: 2-L aspirator bottle with
bottom sidearm (e.g., Corning PYREX®
Product #1220-2L)
37
Production of Recombinant Proteins from Pichia pastoris...
Methods
In this protocol we describe our methods for
methanol inducible recombinant protein production in P. pastoris with secretion of the target protein into the media, followed by IMAC as primary
capture step. Subsequent steps comprise concentration and dialysis (using a tangential flow
filtration (TFF) system) and size exclusion chromatography. We describe the complete process
flow (Table 37.3). However, to be concise, we
refer to our published procedures where applicable but outline in detail our recently developed
RBA-IMAC procedure. Suggestions for alternatives and optimization of the protocol are given
or referred to (see also Refs. [26, 27]).
Seed Lot Production and Fermentation
Genetic engineering of P. pastoris is straightforward using standard molecular biology techniques
[36], with a range of vectors and strains available
[37, 38]. The vector used in this method is
pPICZa(alpha) [37], which carries the methanol
inducible alcohol oxidase I (AOXI, [39]) promoter. As an alternative the constitutive glyceraldehyde-3-phosphate dehydrogenase promoter
(GAP) [9, 37, 40–42] or formaldehyde dehydrogenase promoter (FLD1) [37, 43] from P. pastoris
can be used. Considerations to choose either promoter can be related to, for example, (1) the need
to produce instable proteins or those that are toxic
to the host cell (both require inducible expression); or (2) health and safety risks and cost of
bulk amounts of highly flammable methanol on
site. Vector pPICZa(alpha) carries the a(alpha)factor secretion signal, which directs expression
of the recombinant protein into the broth. This
simplifies downstream processing because P. pastoris secretes only a small amount of host cell proteins (Fig. 37.2); for example, on glucose media
there are only some 20 proteins secreted at detectable levels [22]. Upon initial construction of the
expression vector in E. coli, the construct is stably
integrated in the P. pastoris chromosome, and following expression testing a clone is taken forward
411
to produce a seed lot. Subsequently, this clone is
tested in the bioreactor.
Vector Construction and Seed Lot
Preparation
For details on plasmid construction and P. pastoris transformation, see Ref. [37].
See Notes 4 through 6 for additional considerations regarding vector and seed lot preparation.
Fermentation
For details on recombinant protein production
using P. pastoris by fermentation, see Ref. [26].
Primary Capture: Expanded
and Radial Flow Bed Adsorption IMAC
Packed bed columns can be divided in axial and
radial flow devices (Fig. 37.3; Table 37.4). Axial
flow columns have a long bed length and consequently contain a large number of theoretical
plates, which gives the column a potential for
high resolution. Only clarified feed can be used
and therefore the major application of axial columns will be for secondary purification and
polishing steps in downstream processing.
Preconditioning of the feed can be performed by
(continuous) centrifugation or (cross-flow or
dead-end) filtration. For larger volumes this is a
challenge due to the high biomass of P. pastoris
at harvest time.
In contrast, an axial column where the bed is
allowed to expand (see Fig. 37.3B1, B2) can be
fed with feedstock almost “as is” directly from
the bioreactor. The only preconditioning would
involve dilution of the biomass, buffer additions, (e.g., NaCl), and/or pH changing (acid or
base addition). The extent to which the bed
expands is related to particle size of the matrix,
flow rate, and viscosity of the applied sample
(and buffer). Unclarified feed may cause deposition of biomass onto the matrix that limits
product binding; furthermore, cell-to-cell
aggregation can restrict the flow rate. Both
potentially can be counteracted by the addition
of chemical additives [44]. Amongst others,
suppliers for laboratory and process scale
412
B. Tolner et al.
Table 37.3 Process flow scheme and timeline
Process stage
Clone selection and seed lot production
Primary seed
Secondary seed
Start bioreactor
AOXI: generate biomass (glycerol)
GAP: generate product (e.g., glucose)
Product generation
AOXI: generate product (methanol)
GAP: generate product (e.g., glucose)
Set up downstream processing
(EBA- or RBA-IMAC; TFF; FPLC)
Protein isolation
Timing
1–4 months
Day 1
Day 2
Day 3
Day 4–6
Day 7
The flow process and timelines of the production processes using either AOXI or GAP promoter to drive generation of
recombinant protein are indicated. TFF tangential flow filtration; FPLC fast protein liquid chromatography
Fig. 37.2 Relative to the target protein (a single chain
Fv), the abundancy of secreted native P. pastoris proteins
is low at 40 h post methanol feed start (preharvest sample). The bioreactor conditions were as outlined in Ref.
[26]. Gel: 12% poly acryl amide, tris–glycine buffer;
staining with Coomassie brilliant blue R250. Lane 1,
molecular weight markers (see Blue® Plus2 Pre-Stained
Standard, Invitrogen); lane 2, preharvest sample. c1–5,
contaminating proteins; arrow, target protein
expanded bed equipment and matrices are GE
Healthcare [45] and Upfront [46]. We have used
EBA-IMAC (GE Healthcare StreamLine 50
column with chelating matrix) successfully for
laboratory- and clinical-grade recombinant protein production [13, 14, 28–33, 35].
The disadvantages of EBA-IMAC, such as
high buffer consumption, slow flow, and overall
long process time, can be overcome by implementation of a radial flow column. Radial flow
columns are available from small laboratory to
full process scale (for review, see Ref. [47]). The
two main radial flow column suppliers are
Sepragen [48] and Proxcys [49]. A radial flow
column is cylindrical at process scale, and the
chromatography step takes place by passing the
feedstock through the matrix from the outside to
the inside of the column (see Fig. 37.3C).
Consequently the feed enters through a large surface area and exits through a much smaller area.
The distance between entry and exit is the bed
height or column length and is short relative to
axial columns. This short length combined with
large matrix beads makes it very suitable for primary capture and gradient elution. Also, the relatively low height of the radial column relative to
the axial column allows for a much lower pressure differential between the highest and lowest
point of the column. This pressure differential is
prohibitive to build large tall axial columns; consequently, large axial columns tend to be pancake-shaped. (For a performance comparison
between similar-sized axial and radial columns,
see Ref. [50].)
In this protocol we use radial flow adsorption
in IMAC mode (RBA-IMAC) with a chelating
matrix consisting of large-size beads (Sterogene
37
Production of Recombinant Proteins from Pichia pastoris...
Fig. 37.3 Axial and radial flow chromatography. (a)
Axial flow column (packed bed). (b) Axial flow expanded
bed column, collapsed (B1), and expanded in up flow
(B2). (c) Radial flow column (packed bed). A, adapter; M,
413
matrix; arrows, flow direction. Dimensions are not to
scale: smaller axial flow columns are thin and long, and
larger axial flow columns tend to be pancake-shaped
Table 37.4 Characteristics of radial and axial flow columns
Bed type
Relative flow rate
Feed
Usage
Axial flow bed
Fixed size
Slow-medium
Preconditioned
Secondary and
polishing steps
Medium
Medium
Buffer consumption
Total process time (including
setup and cleaning)
High
Resolutiona
Application surface
Small
Scalable
Low
a
This is application-dependent; see Refs. [47,50]
Expanded bed
Expanding
High
Bioreactor content as is; or
crude part purified
Primary capture step
(secondary steps)
High
Slow
Radial flow bed
Fixed size
High
Bioreactor content as is; or
crude part purified
Primary capture step
(secondary steps)
Medium
Fast
Low
Small
Medium
low
Large
High
Cellthru BigBead) that allow capture of Histagged proteins directly from an unclarified feed.
The column used is a small process/laboratory
scale wedge-shaped column. In essence this
wedge is a “slice” from the larger cylindrical columns with the same characteristics and therefore
is inherently scaleable. Relative to EBA [27],
RBA has several advantages: the buffer consumption is lower, the flow rate is faster, and the overall process time is reduced (see also Table 37.4).
However, RBA-IMAC is more susceptible to
clumping of cells, which, depending on severity,
could restrict flow and demand a further dilution
of the feed.
The following section presents guidelines for
installing and implementing a Proxcys radial
flow column (AXCIS MD 62 MKIII) in the process flow (see Note 7).
Installation in the Process Flow
and Packing of the Column
Use the Proxcys AXCIS MD 62 MKIII manual to
assemble the column and use the drawings to
familiarize yourself with all connections.
414
1. Verify the flow rate of the Watson Marlow
pump: Place the outlet line into the measuring cylinder and measure the actual flow rate
at a specific pump setting. Determine the
pump setting for flow rates of 10, 50, 100,
and 200 mL/min (see Note 8).
2. Prepare the following radial flow column
setup (Fig. 37.4a):
• Connect the feed line from the feed container to the Watson Marlow pump head
tubing.
• Use connecting tubing, to connect pump
head tubing and pressure gauge.
• Use connecting tubing, to connect pressure gauge to the column inlet port using
the push fitting.
• Connect the column via the outlet line to
the waste container using the push fitting
on the outlet port. (see Note 9).
3. Fill the feed container with packing buffer
(0.5 M NaCl/0.5× PBS) and flush the empty
column with packing buffer to remove any
debris or dust particles attached to the inside
walls and frits.
4. Mix the IDA chelating Cellthru BigBead
resin bottle and transfer approximately
150 mL resin into a 1-L Duran bottle; add
500 mL of packing buffer and place on a stirrer platform (see Note 10).
5. Disconnect the column outlet line from the
outlet port; this closes the outlet port (see
Fig. 37.4b; see Note 9).
6. Place the feed line into a feed bottle with
packing buffer.
7. Remove the tubing located between the
gauge and the inlet port; put a new section of
tubing between the gauge and the MPC connector on the packing port; and open the column packing port.
8. Place the column upside down to facilitate
entry of the beads into the column while
packing (see Note 11).
9. Connect tubing to the column inlet port using a
push fitting, and place the other end of the tubing into the feed bottle to allow recirculation.
10. Start the Watson Marlow pump slowly at
10 mL/min to ensure that flow into the column
is unrestricted and that there is no leakage.
B. Tolner et al.
11. Increase the flow rate to 50 mL/min, again to
ensure that flow into the column is unrestricted and that there is no leakage.
12. Stop the pump. The system is now ready for
packing.
13. Transfer the feed line into the resin bottle
(leave the stirrer on).
14. Transfer the column inlet line into the resin
bottle to allow recirculation while packing.
15. Start the Watson Marlow pump slowly at
10 mL/min to ensure that flow of the beads
into the column is unrestricted.
16. Increase the speed to 50 mL/min. While
packing is in progress the gauge will show
that there is (virtually) no pressure (reading
about 0) in the system.
17. When the bed is almost full (15 min), the
pressure will increase sharply to about 20 psi.
Decrease the flow rate to 10–20 mL/min and
continue packing under low pressure.
18. The bed is packed when the pressure again
increases sharply at a slow flow rate.
19. Close the column packing port and remove
the tubing from this port by disconnecting the
MPC connector and replace it with blank
MPC connector. Disconnect the line on the
inlet port using the push fitting. Reconnect the
tubing section between inlet port and gauge.
20. Connect the outlet line back to the outlet port
with the push fitting.
At this point the process can be paused and the
system prepared for storage by alkali treatment
(see “Treating the Column with NaOH”). Upon
storage the column has to be neutralized (see
“Neutralizing the Column with Deionized
Water”). Alternatively, the column can be primed
with copper (see “Copper Priming and
Equilibrating with Equilibration Buffer”).
Treating the Column with NaOH
1. Transfer the feed line into storage solution
(0.1 M NaOH and 0.5 M NaCl)
2. Transfer the outlet line into the waste
container
3. Feed five column volumes of storage solution
at 50 mL/min through the column; subsequently
disconnect the inlet and outlet lines and store
the column at 2–8 °C until further use.
37
Production of Recombinant Proteins from Pichia pastoris...
415
Fig. 37.4 Configuration of a
radial flow column in the
process flow. (a)
Configuration for system
flushing and loading of feed
stock. (b) Configuration for
packing of the column
Neutralizing the Column with Deionized
Water
1. Transfer the feed line into the feed container
and fill this with sterile deionized water (or
water for irrigation).
2. Transfer the outlet line into the waste container
3. Start the pump at 200 mL/min and flush the
column with sterile water until the pH of the
waste matches the pH of the applied water
(see Note 12).
Copper Priming and Equilibrating
with Equilibration Buffer
1. Transfer the feed line into 2 L 0.1 M copper
sulphate solution.
2. Start the pump and apply all copper sulphate
to the column at 200 mL/min.
3. Stop the pump once the copper has been
applied.
4. Transfer the feed line into sterile water.
5. Wash the column with sterile water to remove
any unbound copper from the column. Check
the OD600 of the waste outlet and continue
washing the column until it matches the OD of
the feed.
6. Transfer the feed line into 2 L sterile equilibration buffer (0.5 M NaCl/0.5× PBS/10 mM
Imidazole).
7. Equilibrate the column with minimum of ten
column volume of equilibration buffer.
Adding the Biomass to the Column
1. Leave the feed line in the equilibration buffer
container.
2. Transfer the outlet line into the equilibration
buffer container.
3. Start the Watson Marlow pump slowly at
10 mL/min and allow the air to be displaced
in both lines.
4. Apply a short pulse of the feed pump to
shock remove air from the frits and the
column.
5. Perform the same procedure in reverse direction
to remove the air from the column.
416
6. Repeat steps 3–5, if needed (see Note 13).
7. Connect the feed line to the side arm of a 2-L
“side arm aspirator” bottle (mixer vessel).
8. Place a sterile magnetic stirrer bar inside the
mixer vessel.
9. Place the mixer vessel on magnetic stirrer
platform.
10. Connect the bioreactor harvest line to the
inlet of the mixer vessel.
11. Connect the dilution buffer container outlet
line to the inlet of the mixer vessel.
12. Add 500 mL of biomass dilution buffer (2 M
NaCl/2× PBS/40 mM Imidazole) into the
mixer vessel and transfer 1,500 mL of bioreactor culture into the mixer vessel.
13. Allow to mix for 1 min and subsequently apply
to the radial flow column at 200 mL/min.
14. After each liter, collect a 1-mL aliquot of the
flow through, spin down the cells, and store
the supernatant of each fraction for later
analysis (column breakthrough analysis).
15. Repeat steps 12–14 until the complete bioreactor content has been applied (see Note 14).
16. Stop the pump and transfer the feed line into
a container with wash buffer.
17. Restart the pump to wash any remaining cells
and cell debris from the matrix.
18. After each liter, collect a 1-mL aliquot of the
wash flow through, spin down the cells, and
store the supernatant of each fraction for later
analysis (leakage from matrix).
19. Apply wash buffer until the OD280 is steady.
Eluting of Nonspecifically Bound
Contaminants from the Matrix
1. Apply 2 L matrix wash buffer (0.5 M
NaCl/0.5× PBS/40 mM Imidazole) to the column, at 200 mL/min; collect 50-mL fractions.
2. Measure OD280 of each fraction and store an
aliquot of each fraction for later analysis. (see
Note 15) (Fig. 37.5).
Eluting Target Protein from the Matrix
1. Apply 2 L elution buffer (0.5 M NaCl/0.5×
PBS/200 mM Imidazole) to the column, at
200 mL/min; collect 50-mL fractions.
2. Measure OD280 of each fraction and store an
aliquot of each fraction for later analysis.
3. See Fig. 37.5.
B. Tolner et al.
4. Pool the fractions with the highest OD280 for
further downstream processing
5. Continue the elution buffer feed to remove
any remaining cells and cell debris from the
matrix by applying equilibration buffer until
the OD280 is steady.
Regeneration of the Column: Removing
Copper and Washing with Sodium
Hydroxide
1. Apply 2 L EDTA solution to the column, at
200 mL/min; collect the first 10 fractions of
50 mL. Store an aliquot of each fraction for
later analysis (detection of protein still bound
to the column).
2. Apply all EDTA to the column.
3. Transfer the feed line into the strip solution.
4. Apply sodium hydroxide until the A280 of the
waste is the same as that of the applied
solution.
5. Neutralize the column as per instructions in
“Neutralizing the Column with Deionized
Water”.
The column can now be prepared for storage
(see “Treating the Column with NaOH”) or
recharged with copper for the next round of
purification (see “Copper Priming and
Equilibrating with Equilibration Buffer”).
Downstream Processing
At this stage many options are available for further downstream processing, depending on the
nature of the protein and/or final application in
question. These are beyond the scope of this
chapter (see Note 16).
Notes
1. Health and safety considerations: before starting practical work, it should be verified that
the procedures given in this method are in
alignment with local rules and regulations.
Risk assessments, waste disposal, and decontamination procedures should be in place.
Also, any hazard sheets (Material Safety Data
Sheet, MSDS) should be read and first-aid
37
Production of Recombinant Proteins from Pichia pastoris...
417
Fig. 37.5 Anticipated purification profile during radial
flow bed elution, illustrated with a single chain Fv (scfv).
(a) OD280 profile of eluted fractions. Fractions 1–20,
40 mM elution; fractions 21–43, 200 mM elution. Each
fraction was 50 mL. (b) 12% polyacrylamide Coomassie
(brilliant blue R250) stained gel of eluted fractions. Lane
1, molecular weight marker; lane 2, fraction 4 of 40 mM
elution; lanes 3–9, fraction 23–29 of 200 mM elution; lane
10, pool of fractions 23–30. Arrow, target protein
measures should be known and available.
Where applicable, personal protection equipment (PPE) should be used.
• Ensure acid and base are stored separately
in suitable storage cupboards. Special care
should be taken with ammonia, which
develops pressure on standing, particularly
when warm. Release pressure in an appropriate ventilated hood!
• Methanol is highly flammable, and suitable
containers should be used.
2. The equipment, chemicals, disposables, and
so on described in this method are currently in
use in the authors’ laboratory; however, these
should be easily exchangeable with equipment/materials of equivalent specification.
3. Pichia pastoris belongs to the group of microorganisms that are “generally regarded as
safe” (GRAS).
• Decontamination: Spray work area and
other surfaces with 70% (v/v) ethanol and
leave for 5 min.
• Spills: Decontaminate the area by sprinkling Virkon granules.1 For larger spills
adsorbent containment booms2 can be used;
1
e.g., http://www2.dupont.com/DAHS_EMEA/en_GB/
products/disinfectants/virkon_s/index.htmL.
2
e.g., http://www.newpig.com.
418
4.
5.
6.
7.
B. Tolner et al.
these effectively adsorb the liquid waste and
subsequently need to be autoclaved (center
of the load should be 121 ºC for 30 min).
Regarding the DNA/protein sequence to be
expressed and vector to be used, consider the
following:
• Codon optimization for expression in
Pichia.
• Presence of amber codons, e.g., E. coli
TG1 (often used in phage display) is an
amber suppressor and Pichia is not.
• Restriction sites PmeI, SacI, or BstXI;
these enzymes are used to linearize plasmid pPICZa(alpha) before insertion into
Pichia and consequently should not be
located in the gene.
• Presence of LysArg encoding sequences;
this site can be prone to cleavage by the
native Pichia Kex2 protease and cause
protein breakdown.
• Presence of consensus sequence for
N-glycosylation (Asn-Xaa-Ser/Thr).
• Addition of a stop codon: ligation in frame
at the 3¢ prime causes the addition from
the vector of a sequence that encodes an
Myc and His-tag. Alternatively, a stop
codon or any tag of choice followed by a
stop coden could be placed directly following the inserted gene.
• 5¢ modifications to the vector sequence
encoding KREAEA: this sequence is cleaved
by the Pichia enzymes Kex2 (KR) and Ste13.
However, Ste13 cleavage can be partial (e.g.,
Refs. [51–53]) and removal of the Ste13 recognition sequence could be considered.
Expression of clones can vary substantially;
therefore screening should be performed on
at least 10 or more clones; of each clone the
supernatant and cell pellet should be examined for the target protein.
The seed lot can be very valuable; therefore,
it should be stored in two locations at −80 °C
and should be subjected to minimal characterization like viability, production level sterility (monosepsis), and stability.
Before implementing the RBA-IMAC process, pilot studies can be performed to establish binding conditions for the target protein.
At least optimal conditions for ionic strength,
8.
9.
10.
11.
12.
13.
14.
15.
16.
imidazole concentration, and pH should be
established. For a detailed practical approach,
see appendix 3 in Ref. [27].
Pump performance and tubing can deteriorate in time and “as good practice” should be
measured at a regular interval.
Disconnecting the push fitting will close the
port and reconnecting will open the port.
Stir moderately to avoid damaging the resin.
Turning the column upside down at this stage
allows a more even spread of the beads and
reduces the pressure while filling column.
Follow neutralization using pH strips or a pH
meter.
Before adding the biomass it is important to
ensure that all air is removed from the column; ensure even application of biomass to
the column; and maximize access of the protein to the column matrix. Also, at this point
each connection should be verified and
checked for leakage.
During the biomass application the pressure
should not or only minimally rise. A rise could
suggest clocking up of frits and or matrix and
should be investigated immediately.
It is important to establish that the protein
remains bound during this washing step. If an
excessive amount of protein elutes, then other
concentrations of imidazole should be explored
(e.g., 20 or 30 mM in the wash buffer).
A simple procedure which can be easily
adapted and is in use in the author’s laboratory is the following.
• Sterile filter the protein fraction on a 0.2
mm filter, for example 0.5 L Nalgene,3
especially PES (polyethersulfone) filters
are suitable since they show very low protein binding.
• Concentrate the protein fraction on a TFF
system, e.g., a Labscale system from
Millipore with Pellicon XL membrane4 or
a Pall system.5
• Using the TFF system, dialyze the protein
fraction into a suitable buffer, e.g., 1× PBS.
http://www.nalgenelabware.com/techdata/technical/
FiltrationIndex.asp.
4
http://www.millipore.com/catalogue/item/xx42lss11.
5
http://www.pall.com/laboratory_53238.asp.
3
37
Production of Recombinant Proteins from Pichia pastoris...
• Apply the sample to an FPLC, fitted with a
size exclusion column (e.g., a sample can be
loaded using a superloop to a 500-mL
Superdex [GEHealthcare]) and collect the
fractions of interest.
• If required, remove endotoxins by successive passaging over an endotoxin removal
gel (Thermo Scientific). Progress can be
monitored using a Limulus Amebocyte
Lysate (LAL) test (Pyrogent-Plus 64 Gel
Clot Assay, Lonza Biologics).
Acknowledgements This work was supported by
Cancer Research UK; Department of Health (ECMC,
Experimental Cancer Medicine Network Centre);
Engineering and Physical Sciences Research Council
(EPSRC); The Breast Cancer Campaign; and UCL Cancer
Institute Research Trust.
11.
12.
13.
14.
15.
16.
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Development of a Real-Time
Quantitative PCR Assay
for the Assessment of Uncultured
Zoosporic Fungi
38
Télesphore Sime-Ngando and Marlène Jobard
Abstract
Recently, molecular environmental surveys of the eukaryotic microbial
community in lakes have revealed a high diversity of sequences belonging
to uncultured zoosporic fungi. Although they are known as saprobes and
algal parasites in freshwater systems, zoosporic fungi have been neglected
in microbial food web studies. Recently, it has been suggested that zoosporic fungi, via the consumption of their zoospores by zooplankters,
could transfer energy from large inedible algae and particulate organic
material to higher trophic levels. However, because of their small size and
their lack of distinctive morphological features, traditional microscopy
does not allow the detection of fungal zoospores in the field. Hence, quantitative data on fungal zoospores in natural environments are missing. We
provide a simplified step-by-step real-time quantitative PCR laboratory
protocol, for the assessment of uncultured zoosporic fungi and other zoosporic microbial eukaryotes in natural samples.
Keywords
Fungi • Spores • DNA extraction • Real-time qPCR • Environmental
samples
T. Sime-Ngando (*)
UMR CNRS 6023, Université Blaise Pascal, Clermont
II, 24 Avenue des Landais, BP 80026, Aubière 63171,
Cedex, France
e-mail: telesphore.sime-ngando@univ-bpclermont.fr
M. Jobard
LMGE UMR CNRS, U.F.R. Sciences et Technologies,
24 avenue des Landais, BP 80026, Aubière 63171,
Cedex, France
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_38, © Springer Science+Business Media, LLC 2013
421
422
Introduction
Recent molecular surveys of microbial eukaryotes have revealed overlooked uncultured environmental fungi with novel putative functions
[1–3], among which zoosporic forms (i.e.,
chytrids) are the most important in terms of diversity, abundance, and functional roles, primarily
as infective parasites of phytoplankton [4, 5] and
as valuable food sources for zooplankton via
massive zoospore production, particularly in
freshwater lakes [6–8]. However, due to their
small size (2–5 mm), their lack of distinctive morphological features, and their phylogenetic position, traditional microscopic methods are not
sensitive enough to detect fungal zoospores
among a mixed assemblage of microorganisms.
Most chytrids occupy the most basal branch of
the kingdom Fungi, a finding consistent with
choanoflagellate-like ancestors [9]. These reasons may help explain why both infective (i.e.,
sporanges) and disseminating (i.e., zoospores)
life stages of chytrids have been misidentified in
previous studies as phagotrophic sessile flagellates
(e.g., choanoflagellates) and “small undertermined” cells, respectively. These cells often
dominate the abundance of free-living heterotrophic nanoflagellates (HNFs), and are considered the main bacterivores in aquatic microbial
foodwebs [2, 10]. Their contribution ranges from
10 to 90% of the total abundance of HNFs in
pelagic systems (see review in reference [11]).
Preliminary data have provided that up to 60% of
these unidentified HNFs can correspond to fungal zoospores [12], establishing HNF compartment as a black box in the context of microbial
food web dynamics [4]. A recent simulation analysis based on Lake Biwa (Japan) inverse model
indicated that the presence of zoosporic fungi
leads to (1) an enhancement of the trophic
efficiency index, (2) a decrease of the ratio detritivory/herbivory, (3) a decrease of the percentage
of carbon flowing in cyclic pathways, and (4) an
increase in the relative ascendancy (indicates
trophic pathways more specialized and less
redundant) of the system [13]. Unfortunately,
because specific methodology for their detection
T. Sime-Ngando and M. Jobard
is not available, quantitative data on zoosporic
fungi are missing.
Molecular approaches have profoundly
changed our view of eukaryotic microbial diversity, providing new perspectives for future ecological studies [3]. Among these perspectives,
linking cell identity to abundance and biomass
estimates is highly important for studies on carbon flows and the related biogeochemical cycles
in natural ecosystems [11]. Historically, taxonomic identification and estimation of in situ
abundances of small microorganisms have been
difficult. In this context, our inability to identify
and count many of these small species in the natural environment limits our understanding of
their ecological significance. Thus, new tools that
combine both identification and quantification
need to be developed. Fluorescent in situ hybridization (FISH) has been an assay of choice for
simultaneous identification and quantification of
specific microbial populations in natural environments [14, 15]. However, this technique is limited because of (1) the relatively low number of
samples that can be processed at a time, and (2)
its relatively low sensitivity due to background
noise and the potentially low number of target
rRNA molecules per cell in natural environments
[16]. In contrast, real-time quantitative PCR
(qPCR), which has been widely used to estimate
prokaryotic and eukaryotic population abundances in natural ecosystems, allows the simultaneous analysis of a high number of samples with
a high degree of sensitivity [15].
The main objective of this chapter is to provide, in a simplified step-by-step format, a qPCR
assay for the quantitative assessment of uncultured zoosporic fungi and other zoosporic microbial eukaryotes in natural environments [15],
together with practical advice on how to apply
the method. QPCR was recently used to estimate
fungal biomass in a stream during leaf decomposition [17] and in biological soil crusts [18]. The
interpretation of the semi-quantitative data
obtained in these studies was relatively difficult
because the whole fungal community was targeted (including unicellular, multicellular, and
multinuclear fungal species). Thus, an estimation
of fungal density or even fungal biomass was not
38
Development of a Real-Time Quantitative PCR Assay for the Assessment…
possible. In the following protocol, the primary
targets are zoospores in liquid suspensions.
Because zoospores are unicellular, qPCR data
could be directly converted into cell density estimates (i.e., by multiplying semi-quantitative data
by a number of rDNA copies per cells). Moreover,
we designed primers targeting Rhizophidiales
taxon to limit quantification bias generated by the
variability in the number of rDNA copies within
eukaryotic ribosomal operon.
Materials
1. Gloves (should be worn when manipulating
most of the following materials).
2. Sterile distilled water.
3. 0.6 mm pore size polycarbonate filters.
4. Filtration columns.
5. Sodium dodecyl sulfate (SDS).
6. Proteinase K.
7. TE buffer—10 mM Tris–HCl pH 7.5 (25 °C),
1 mM EDTA.
8. NucleoSpin Plant kit® (Macherey-Nagel,
Bethlehem, PA) with silica-membrane columns and the materials for running the provided protocol from the manufacturer.
9. Molecular-biology-grade agarose.
10. Ethidium bromide—because suspected as a
mutagen, particular care should be taken
when handling (consult safety data sheet).
11. Calf thymus DNA (Sigma-Aldrich, St. Louis,
MO).
12. Oligonucleotidic primers resuspended in
sterile distilled water and stored at −20 °C
(see Note 1).
13. SYBR Green (Sigma-Aldrich).
14. dNTPs—a mixture of dATP, dCTP, dGTP,
and dTTP (10 mM of each), stored at
−20 °C.
15. Thermostable DNA polymerase and reaction
buffer supplied by manufacturer. To avoid
nonspecific amplicon, use hot-start (e.g.,
HotStarTaq, Qiagen, Valencia, CA).
16. Vortexer.
17. Centrifuge.
18. Water bath.
19. Horizontal electrophoresis machine.
423
20. TBE buffer: 50 mM Tris, 50 mM boric acid,
1 mM EDTA, diluted when needed from a
50× stock solution.
21. Spectrophotometer—the
authors
use
NanoDrop (NanoDrop Technologies, Inc,
Wilmington, DE).
22. Disposable conical tubes (1.5 mL); PCR tube
strips or plate with adhesive film and cap
adapted for real-time quantitative PCR
assay.
23. Thermal cycler—we use Mastercycler ep
realplex detection system (Eppendorf).
24. UV transilluminator equipped with a camera
suitable for photographing agarose gels
Methods
DNA Extraction and Purification
Collect zoosporic organisms onto 0.6 mm pore
size polycarbonate filters (after removal of the
algal host by prefiltrations when only zoospores
are targeted) (see Note 2).
1. For cell disruption, incubate the filters in
560 mL of a buffer containing 1% SDS and
1 mg/mL proteinase K in TE buffer for 1 h at
37 °C in a water bath (see Notes 3 and 4).
2. For DNA purification, use the silica-membrane
columns provided with the NucleoSpin Plant
kit® (Macherey-Nagel), following the instructions from the manufacturer (see Note 4).
3. Visualize the integrity and yield of the
extracted genomic DNA in a 1% agarose gel
stained with 0.3 mg/mL of ethidium bromide
solution (Sigma-Aldrich), using UV transilluminator and photograph. For this (1) heat (45 s
using a microwave oven) a mixture of agarose
in 1× TBE buffer; (2) leave it to cool on the
bench for 5 min down to about 60 °C before
adding ethidium bromide (i.e., to avoid vapor
formation); (3) mix and pour into suitable gel
gray with comb and leave to set for at least
30 min; (4) remove the comb and submerge
the gel to 2–5 mm depth in electrophoresis
tank containing 1× TBE buffer; (5) transfer
DNA sample aliquots (i.e., 2 volumes of sample and 1 volume of loading buffer), marker,
424
and the serial dilution of 5–10 ng of calf thymus DNA (Sigma-Aldrich-Aldrich) to the
wells of the agarose gel; and (6) start the electrophoresis migration for about 30 min at
100 V.
4. Calculate DNA extract concentrations from
dilutions of calf thymus DNA (SigmaAldrich), using a standard curve of calf thymus DNA vs. band intensity.
Real-Time qPCR Assays
1. PCR mix contained SYBR Green (SigmaAldrich), 200 mm of each dNTPs, 10 pM of
each primer, 2.5 units of Taq DNA polymerase,
the PCR buffer supplied with the enzyme, and
1.5 mM MgCl2. Vortex briefly (less than 10 s)
and centrifuge the mix before distributing aliquots in suitable PCR tubes (strips or plates)
and place on ice.
2. Add variable quantity of DNA (we used 5 ng
for our environmental freshwater samples, and
10 ng for DNA from appropriate PCR negative control strains) used as template in a final
volume of 25 mL (see Note 5).
3. Standard curve of Ct (see Note 4) vs. DNA
copy number required to calculate target copy
numbers (see Note 6) in each reaction is generated using triplicates of PCR reactions of
tenfold dilutions of linear plasmid (containing
Rhizophidiales
18S
rDNA
insert;
PFB11AU2004) ranging from 100 to 1 × 108
copy/mL (see Note 7). This number of copies
was calculated using the equation: molecules/
mL = a/(b × 660) × 6.022 × 1023, where a is the
plasmid DNA concentration (g/mL), b the
plasmid length in bp, including the vector and
the inserted 18S rDNA fragment, 660 the average molecular weight of one base pair, and
6.022 × 1023 the Avogadro constant [15, 19].
4. Place all tubes (i.e., samples, controls, and
standards) in the real-time qPCR cycler and
run the appropriate cycling program: initial
HotStarTaq activation at 95 °C for 15 min, 35
cycles with denaturation at 95 °C for 1 min,
annealing at 63.3 °C for 30 s with Fchyt/Rchyt
primers pair (see Note 1), elongation at 72 °C
T. Sime-Ngando and M. Jobard
for 1 min, and a final additional elongation
step at 72 °C for 10 min.
5. Using SYBR Green molecule, melting curves
analysis can be performed immediately following each qPCR assay to check specificity
of amplification products (to confirm the
absence of primer dimers or unspecific PCR
products) by increasing the incubation temperature from 50 to 95 °C for 20 min.
6. Analyze the real-time PCR result with the
suitable software. Check to see if there is any
bimodal dissociation curve or abnormal
amplification plot (see Note 5) before calculating the initial concentration of the targeted uncultured fungal 18S rDNA (copies/
mL) in the environmental genomic DNA (see
Note 6).
Notes
1. Consensus (universal) primers can be used to
amplify regions of fungal ribosomal RNA
gene. For natural waters, we have designed
primers specific to chytrids using a database
containing about a hundred 18S rDNA environmental sequences recovered from surveys
conducted in different lakes and sequences
belonging to described fungi [15]. Sequences
were aligned using BioEdit software (http://
www.mbio.ncsu.edu/BioEdit/bioedit.html )
and the resulting alignment was corrected
manually. A great proportion of the environmental chytrid sequences recovered from
lakes was closely affiliated to the
Rhizophidiales. Thus, Rhizophidiales-specific
primers F-Chyt (sequence 5¢ > 3¢: GCAGGCTT
ACGCTTGAATAC) and R-Chyt (sequence
5¢ > 3¢:
CATAAGGTGCCGAACAAGTC)
were designed in order to fulfill three requirements: (1) a GC content between 40 and 70%,
(2) a melting temperature (Tm) similar for both
primers and close to 60 °C, and (3) PCR products below 500 bp (i.e., between 304 and
313 bp depending on the species considered).
The absence of potential complementarities
(hairpins and dimers) was checked using
Netprimer (http://www.premierbiosoft.com/
38
2.
3.
4.
5.
Development of a Real-Time Quantitative PCR Assay for the Assessment…
netprimer/netprlaunch/netprlaunch.html), and
confirmed by inspection of the melting curve
following the qPCR assay.
For targeting uncultured zoosporic fungi, zoospores are discarded from other environmental microorganisms by successive prefiltrations
through 150, 80, 50, 25, 10, and 5 mm filters
before collected them onto 0.6-mm polycarbonate filters. Filters can be conserved at
−80 °C until DNA extraction in appropriate
tubes (2 mL).
Other enzymes such as lyticase can be used
for cell disruption, with no significant difference compared to proteinase K. However, the
one-step proteinase K yields higher amount of
genomic DNA than the lyticase method and
has a better reproducibility. Physical disruption procedures such as sonication or thermal
shocks (i.e., freezing in liquid nitrogen and
thawing) are to be avoided because of the possible degradation of DNA.
A standard phenol–chloroform purification
procedure can also be used but when the
genomic DNA extracts are used as template in
PCR reactions, the DNA purification method
using the commercial kit gave significantly
better results (based on Ct, the threshold cycle
during PCR when the level of fluorescence
gives signal over the background and is in the
linear portion of the amplified curve) than the
phenol–chloroform method. Consequently,
the DNA extraction method using Proteinase
K and the commercial kit was selected and
considered the best overall.
In the case of novel designed primers (see
Note 1) for uncultured fungi, DNA from both
positive and negative plasmids and different
mixtures (e.g., 5, 10, 25, and 50% of the positive plasmids) will be used for the optimization of the conditions (annealing temperature,
cycling), cross-reactivity, the detection limit
(using serial tenfold dilutions of the positive
plasmids; see Note 7), and the amplification
efficiency of the qPCR essays, which should
be at least 90%. Poor primer quality is the
leading cause for poor PCR efficiency. In this
case, the PCR amplification curve usually
reaches plateau early and the final fluorescence
425
intensity is significantly lower than that of
most other PCRs. This problem may be solved
with re-synthesized primers.
6. The initial concentration of target 18S rDNA
(copies/mL) in environmental samples can be
calculated using the formula [(a/b) × c]/d,
where a is the 18S rDNA copy number estimated by qPCR, b is the volume of environmental genomic DNA added in the qPCR
reaction, c is the volume into which the environmental genomic DNA was resuspended at
the end of the DNA extraction, and d is the
volume of sample filtered from which environmental DNA was extracted.
7. In the absence of cultures, plasmids used in
qPCR to construct standard curves and to optimize qPCR reactions come from genetic libraries constructed during previous environmental
surveys [2]. Briefly, the complete 18S rRNA
gene was amplified from environmental
genomic DNA extracts using the universal
eukaryote primers 1f and 1520r. An aliquot of
PCR products was cloned using the TOPO-TA
cloning kit (Invitrogen, Carlsbad, CA) following the manufacturer’s recommendations.
Plasmid containing the insert of interest was
extracted with NucleoSpin® plasmid extraction kit (Macherey-Nagel) following the manufacturer’s recommendations. The 18S rRNA
gene was sequenced from plasmid products by
the MWG Biotech services using M13 universal primers (M13rev (−29) and M13uni (−21)).
Phylogenetic affiliation of sequences acquired
was established using Neighbor-Joining and
Bayesian methods. In our case, positive plasmids contain insert affiliated to target chytrid
(i.e. Rhizophidiales species) and displaying
less than two mismatches with primers F-Chyt
and R-Chyt sequences (see Note 1). The plasmid PFB11AU2004 (Genbank accession number DQ244014) was selected to construct the
standard curve required for qPCR. Linearized
plasmids were produced from supercoiled plasmids by digestion with restriction endonuclease
one-time cutting into the vector sequence.
Linear plasmid DNA concentration can be
determined by measuring the absorbance at
260 nm (A260) in a spectrophotometer.
426
Acknowledgements MJ was supported by a PhD
Fellowship from the Grand Duché du Luxembourg
(Ministry of Culture, High School, and Research). This
study receives grant-aided support from the French ANR
Programme Blanc #ANR 07 BLAN 0370 titled DREP:
Diversity and Roles of Eumyctes in the Pelagos.
T. Sime-Ngando and M. Jobard
10.
11.
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Nucleic and Protein Extraction
Methods for Fungal
Exopolysaccharide Producers
39
Jochen Schmid, Dirk Mueller-Hagen, Volker Sieber,
and Vera Meyer
Abstract
Exopolysaccharides (EPS) produced by filamentous fungi often interfere
with common methods for the extraction of nucleic acids or proteins.
Similar precipitation behavior (e.g., by formation of hetero-triple helixes
between the EPS and nucleic acid strands) severely minimizes yields of
DNA or RNA extraction. In fungal strains that produce very high amounts
of EPS, the fraction of mycelium per volume of fermentation broth is low,
which requires removal of EPS to generate sufficient concentration of proteins or nucleic acids for downstream applications.
In this chapter we provide adapted methods for the extraction of nucleic
acids and proteins of strong EPS-producing fungal strains using Sclerotium
rolfsii as an example. The concentration and purity achieved are suitable
for applications such as sequencing, RT-PCR, or 2D-PAGE.
Keywords
Exopolysaccharides • DNA • RNA • Protein • Extraction • Fungi •
Scleroglucan • Sclerotium rolfsii
Introduction
J. Schmid (*) • V. Sieber
Chair of Chemistry of Biogenic Resources,
Technische Universität München, Schulgasse 16,
Straubing, 94315, Germany
e-mail: J.schmid@tum.de
D. Mueller-Hagen • V. Meyer
Department of Applied and Molecular Microbiology,
Berlin University of Technology, Gustav-MeyerAllee 25, Berlin 13355, Germany
Traditional methods for RNA, DNA, and protein
extraction do not account for the presence of
exopolysaccharides (EPS) at higher concentrations. A similar precipitation behavior of polysaccharides and other biological polymers such
as RNA, DNA, and proteins, as well as an adverse
volumetric ratio between mycelia and secreted
EPS, lead to a low yield, if any at all. Alcohol
precipitation is commonly used for concentrating, desalting, and recovering nucleic acids.
Using alcohols such as ethanol or isopropanol as
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_39, © Springer Science+Business Media, LLC 2013
427
428
a precipitant, DNA or RNA as well as polysaccharides lose their hydration hull and are thereby
co-extracted. Precipitation of DNA using isopropanol at 4 °C is very effective; however, it is
accompanied by the co-precipitation of salts.
This negative effect is further increased in the
presence of polysaccharides. As a result, with
organisms that secrete large amounts of extracellular polysaccharides, these methods result in
less concentrated DNA that is salt-enriched and
highly contaminated with polysaccharides and
therefore not suitable for downstream applications such as PCR or restriction analysis. One
solution to counter this problem involves CTAB,
which either precipitates the DNA or the contaminants [1]. At concentrations of 0.7–0.8 M NaCl,
polysaccharides precipitate while nucleic acids
remain in solution. This method is also suitable
for purification of DNA and was also modified
for high EPS-producing organisms [2] and for
other highly EPS-contaminated products such as
banana [3]. As with nucleic acids, proteins may
interact with EPS, which results in a loss of certain proteins for downstream applications [4, 5].
Product yield is also hampered by an unfavorable ratio between the amount of mycelia and
EPS, which are intensively swollen due to incorporation of water. This has to be opposed either
by removal of EPS or the incorporated water.
Commercially available DNA and RNA
extraction kits focus on samples such as plant
leaves and blood, which show only a low contamination with polysaccharides. However, in
our lab, extensive tests of a spectrum of these kits
showed that they are not applicable to high EPS
levels, which are achieved in cultivation of
Sclerotium rolfsii, for example, or the bacterial
EPS producer Xanthomonas campestris.
Another approach to get a grip on nucleic acid
and protein extraction from high EPS-producing
fungi is the development of cultivation conditions
that impede EPS production. However, this
approach alone is not suitable for sophisticated
methods such as comparative transcriptome analysis [6].
In this chapter, methods optimized for the
extraction of DNA, RNA, and proteins from high
EPS-producing fungal strains are presented. We
J. Schmid et al.
developed these methods for transcriptomic and
proteomic analyses of S. rolfsii, an industrially
important EPS-producing filamentous fungus of
the genus basidiomycota. DNA, RNA, and proteins extracted were of high purity and yield and
are applicable for many methods, such as microarray analysis, cDNA synthesis, RT-PCR, cloning,
and 2D-PAGE. These protocolls can also act as a
starting point for other high-level polysaccharideproducing organisms, such as bacteria, microalgae,
and yeast, and also for metagenomic approaches,
which are very often limited by the presence of
polymeric substances such as polysaccharides.
Materials
See Note 1.
1. Cultivation media (see Note 2).
2. Filter gauze (for S. rolfsii with a pore size of
70 mm, see Note 3).
3. Sterile double distilled water
4. Liquid nitrogen (see Note 4)
5. Dismembrator (e.g., FastPrep FP220A
Instrument and Braun Mikro Dismembrator
2) (see Notes 5 and 6).
6. DNA extraction buffer (10 mM Tris/HCl pH
8.0, 0.1 M EDTA pH 8.0, 0.5% SDS) (see
Note 7).
7. RNA extraction buffer (4 M GuSCN, 0.1 M
Tris/HCl pH 7.5, 1% b-Mercaptoethanol
(0.14 M). Solve 50 g of GuSCN in 10 mL of
1 M Tris/HCl pH 7.5. Add to 100 mL with
H2O. Solution is filtered sterile and stored at
RT. Directly before usage, sterilized and
stored 1% of Mecaptoethanol is added).
8. RNAse (10 mg/mL).
9. Proteinase K (20 mg/mL).
10. DEPC-H2O (see Note 8).
11. Chloroform/isoamylalcohol (24:1).
12. Isopropanol (see Note 9).
13. Phenol (Te-buffered) (see Note 1).
14. Ice.
15. Vortexer.
16. Magnetic stirrer.
17. Water bath.
18. Microcentrifuge
(e.g.,
Hettich
Lab
Technology, Tuttlingen Germany).
39
Nucleic and Protein Extraction Methods for Fungal Exopolysaccharide Producers
19. Disposable polypropylene microcentrifuge
tubes, 1.5 and 2 mL conical (Greiner,
Germany).
20. Ultracentrifuge (Sorvall, Dupont, Bad
Homburg).
21. Single-use ultracentrifugation tubes (suitable
to the rotor).
22. Centrifuge tubes (suitable for 45,000×g).
23. Rotation incubator.
24. 10% Na-Laurylsarcosine.
25. AN-Prot-Ex buffer (25 mM Hepes pH 7.5,
50 mM KCl, 5 mM MgCl2, 0.1 mM EDTA
pH 8.0, 10% glycerol, 0.7 mL mercaptoethanol/1 mL, 35 mg/mL PMSF, 0.7 mg/mL
Pepstatin A, 0.5 mg/mL Leupeptin).
26. Membrane-lysis buffer (5 mM Tris/HCl pH
7.4, 2 mM EDTA, 0.7 mg/m. Pepstatin A,
35 mg/mL PMSF, 0.5 mg/mL Leupeptin).
27. Membrane-suspension buffer (75 mM Tris/
HCl pH 7.4, 12.5 mM MgCl2, 5 mM EDTA).
28. Impact-resistant 2-mL tubes pre-filled with
Lysing Matrix C particles.
29. PE-buffer (0.1 M Tris/HCl pH 8.8, 10 mM
EDTA, 0.4% 2-mercapto-ethanol, 0.9 M
sucrose).
30. IEF-buffer (7 M urea, 2 M thiourea, 4% (w/v)
CHAPS, 20 mM DTT, 20 mM Tris, 1%
Zwittergent pH 3–10, 0.5% Pharmalyte pH
3–10, 0.002% bromophenol blue).
Methods
Here, we present methods suitable for the extraction of DNA, RNA, and proteins from fungi that
produce high amounts of exopolysaccharides.
Obtained nucleic acids or proteins can be used
for many applications, such as transcriptomic or
proteomic analyses. With minor modifications all
methods are applicable to other fungi that produce low amounts of EPS or no EPS at all as well
as to bacteria.
Small-Scale Preparation of Genomic
DNA from S. rolfsii
The method describes a fast procedure for
obtaining DNA from fungal exopolysaccharide
429
producers with sufficient purity for subsequent
PCR assays and is based on a method described
earlier [7]. The number of tubes can be adapted
if needed.
1. Inoculate 1 mL of cultivation media in a
2-mL reaction tube with a piece of mycelia
(1 or 2 mm in diameter of a freshly overgrown plate) and incubate overnight at 28 °C
with shaking.
2. Harvest mycelia in a tabletop centrifuge for
5 min with 12,000 rpm at RT.
3. Wash once with 1 mL ddH2O.
4. Resuspend the pellet subsequently in 600 mL
ddH2O.
5. Disrupt cells by three times repeated shock
freezing in liquid nitrogen following thawing
for 5 min at 75 °C.
6. Purify DNA by adding 500 mL phenol and
thoroughly shaking the tube for 2 min.
7. Separate phases in a tabletop centrifuge for
10 min with 12,000 rpm at 4 °C.
8. Transfer the top phase to a new tube.
9. Add 400 mL of chloroform and homogenize
thoroughly by shaking the tube for 2 min.
10. Separate phases in a tabletop centrifuge for
10 min with 12,000 rpm at 4 °C.
11. Transfer the top phase to a new tube.
12. Add two volumes of ethanol (96%) and 1/25
volume 3 M Na-acetate.
13. Precipitate DNA for 1 h at −20 °C or overnight (see Note 10).
14. Centrifuge for 15 min with 12,000 rpm at
4 °C.
15. Wash the pellet once with 70% ethanol.
16. Air-dry the pellet for 15 min (see Note 11).
17. Resuspend DNA pellet in 20 mL ddH2O.
18. Use 1 mL for PCR.
Large-Scale Preparation of Genomic
DNA from S. rolfsii
This method describes a general procedure for
obtaining high concentrations of high-quality
DNA from fungal exopolysaccharide producers.
1. Inoculate 100 mL of respective medium with
mycelium of S. rolfsii (1/8th of a freshly overgrown Petri dish cut into stripes with a scalpel) and add a sterile magnetic stirrer bar.
430
2. Incubate at 28 °C at 250 rpm on a magnetic
stirrer (see Note 12).
3. Harvest mycelia after 48 h of cultivation by
filtration through a piece of gauze.
4. Wash mycelia twice with hot water (85 °C)
to remove scleroglucan.
5. Shock-freeze mycelium in liquid nitrogen
(mycelia can be stored for several weeks at
−80 °C).
6. Transfer 500 mg of mycelium to a pre-frozen
Teflon cup (see Note 5).
7. Add 700 mL DNA extraction buffer.
8. Disrupt mycelia with a dismembrator (e.g.,
Braun Mikro-Dismembrator 2) for 2 min
(see Note 6). Use of FastPrep FP220A will
result in heavily sheared DNA.
9. Thaw the mixture and transfer to a 2-mL tube.
10. Add DNA extraction buffer to final volume
of 1.8 mL.
11. Add 10 mL of RNAse (10 mg/mL).
12. Incubate for 10 min at 37 °C.
13. Add 10 mL of proteinase K (20 mg/mL).
14. Incubate for 30 min at 50 °C.
15. Divide the mixture in two equal parts.
16. Mix each with 900 mL of phenol.
17. Homogenize the two resulting phases by
gentle shaking for 2 min.
18. Centrifuge for 10 min with 12,000 rpm at RT
in a tabletop centrifuge.
19. Carefully transfer the top phase to a new
2-mL tube (see Note 13).
20. Add again 900 mL of phenol.
21. Repeat steps 17 and 18.
22. Transfer the top phase carefully to a new
2-mL tube (see Note 13).
23. Add 900 mL of chloroform/isoamylalcohol
(24:1).
24. Repeat steps 17 and 18.
25. Combine both top phases and add an equal
amount of isopropanol.
26. Do not mix for the first 5 min (see Note 14).
27. After 5 min mix by inverting the tube
carefully.
28. Precipitate DNA for 30 min at −20 °C or
overnight (see Note 10).
29. Centrifuge the sample for 5 min with
8,000 rpm at 4 °C in a tabletop centrifuge.
J. Schmid et al.
30. Rinse the DNA pellet once with 70%
ethanol.
31. Centrifuge for 2 min with 8,000 rpm at
4 °C.
32. Air-dry the pellet for 15 min (see Note 11).
33. Re-dissolve the DNA pellet in 50 mL ddH2O
and incubate for 10 min at 65 °C.
34. DNA is now ready to use for PCR, restriction
analysis, or Southern blot etc. (see Note 15).
Isolation of RNA from S. rolfsii using
CsCl-Pads
The method given is based on a method by
Chirgwin et al [8] and describes an efficient procedure for obtaining RNA from fungal exopolysaccharide producers. In our hands, RNA
purification based on CsCl is the method of
choice for EPS-producing microorganisms.
1. Inoculate 100 mL of respective medium with
mycelium of S. rolfsii (1/8th of a freshly overgrown Petri dish cut into stripes with a scalpel) and add a sterile magnetic stirrer bar.
2. Incubate the flask at 28 °C with 250 rpm on a
magnetic stirrer.
3. Harvest mycelia after 48 h of cultivation by
filtration through a piece of gauze (70 mM)
(see notes 3, 16, and 17).
4. Wash mycelia with sterile ddH2O.
5. Transfer approximately 1 g mycelium to a
pre-chilled (liquid N2) Teflon cup, filled with
two steel balls.
6. Disintegrate the mycelium for 2.5 min with a
dismembrator (see Note 6).
7. Freeze again in liquid N2.
8. Add 5 mL of RNA-extraction buffer to the
Teflon cup.
9. Resuspend the pulverized mycelium and
transfer to an SS-34 centrifuge tube, seal
with parafilm and agitate.
10. Add 250 mL of 10% Na-Laurylsarcosine and
gently turn the tube.
11. Centrifuge for 10 min with 9,000 rpm at RT.
12. Pre-fill an ultracentrifuge tube with 5 mL of
the CsCl-pad (3.5 mL 5.7 M CsCl + 0.01 M
EDTA).
39
Nucleic and Protein Extraction Methods for Fungal Exopolysaccharide Producers
13. Carefully overlay the CsCl-pad with 1.5 mL
of the supernatant (keep ultracentrifuge tube
angular for adding the supernatant gently at
the boundary).
14. Centrifuge for 19 h with 125,000 ´ g at 4 °C
using a swing bucket rotor (e.g., AH650).
15. Carefully remove supernatant leaving
approximately 1 cm of supernatant above
bottom level of the ultracentrifuge tube.
16. Remove bottom of the ultracentrifugation
tube (containing the RNA pellet) over the
surface of the resting supernatant with a hot
scalpel (see Note 18).
17. Carefully remove the last supernatant.
18. Dissolve the pellet in 300 mL TE-SDS buffer
and 1/10 Vol of 8 M LiCl containing 2.2 Vol
ice-cold EtOH (96%).
19. Transfer the dissolved pellet into 1.5-mL
reaction tube.
20. Centrifuge for 15 min with 10,000 rpm at
4 °C.
21. Wash the pellet once in 500 mL ice-cold
EtOH (80%).
22. Centrifuge for 15 min with 10,000 rpm at
4 °C.
23. Discard the supernatant and air-dry for
10 min.
24. Dissolve the pellet in 100 mL of DEPC-H2O.
Methods for Protein Isolation from EPS
Producers
Extraction of Soluble Proteins
The following method is an easy-to-use protocol
for the extraction of cytosolic, soluble proteins
from a fungal exopolysaccharide producer. The
number of tubes can be adapted to the amount
needed. Raising the volume is not advised.
1. Inoculate 100 mL of respective medium with
mycelium of S. rolfsii (one-eight of a freshly
overgrown Petri dish is cut into stripes with a
scalpel) and add a sterile magnetic stirrer bar.
2. Incubate the flasks at 28 °C with 250 rpm on
a magnetic stirrer.
3. Harvest the mycelia after 48 h of cultivation
by filtration through a piece of gauze (70 mM)
(see Note 3).
431
4. Wash mycelia with sterile ddH2O.
5. To remove excess water, squeeze the mycelium in the gauze (see Note 17).
6. Immediately shock-freeze mycelium in liquid nitrogen (mycelia can be stored for several weeks at −80 °C).
7. Transfer 750–1,500 mg of mycelium into
N2-pre-frozen Teflon tubes with two steel
balls.
8. Add one volume of An-Prot-Ex buffer.
9. Disrupt mycelia in a dismembrator (e.g.,
Braun Mikro-Dismembrator 2) for 2 min
(see Note 6).
10. Refreeze in liquid N2.
11. Thaw the mixture on ice and transfer to a
2-mL tube.
12. Remove cell debris by centrifugation for
15 min with 14,000 rpm at 4 °C (see Note 19).
13. Transfer the supernatant into a fresh 1.5-mL
tube and centrifuge for an additional 60 min
with 14,000 rpm at 4 °C to remove the
exopolysaccharide.
14. Transfer the resulting supernatant into a fresh
1.5-mL tube and store on ice or use
immediately.
Extraction of Membrane Proteins
The following method is based on a Web-protocol
(http://www.westernblotting.org/protocol%20
membrane%20extraction.htm), and was optimized to meet the requirements of high-EPS-producing fungi.
1. Inoculate 100 mL of respective medium with
mycelium of S. rolfsii (1/8th of a freshly
overgrown Petri dish is cut into stripes with a
scalpel) and add a magnetic stirrer bar.
2. Incubate the flask at 28 °C with 250 rpm on a
magnetic stirrer.
3. Harvest the mycelia after 48 h of cultivation
by filtration through a piece of gauze (70 mM)
(see Note 3).
4. Wash mycelia with sterile ddH2O.
5. To remove excess water, squeeze the mycelium in the gauze (see Note 17).
6. Immediately shock-freeze mycelium in liquid nitrogen (mycelia can be stored for
several weeks at −80 °C).
432
7. Transfer 750–1,500 mg of mycelium into
Teflon tubes with 2 steel balls pre-frozen in
liquid nitrogen.
8. Disrupt mycelia in a dismembrator (e.g.,
Braun Mikro-Dismembrator 2) for 2 min
(see Note 6).
9. Transfer the powdered mycelium into a tube
(e.g., SS-34) pre-filled with 10 mL membrane-lysis buffer.
10. Vortex for 20 s to homogenize the pellet.
11. Centrifuge with 500×g for 15 min at 4 °C.
12. Transfer supernatant into a fresh SS-34 tube.
13. Add 5 mL lyses buffer to the pellet and resuspend by vortexing for 20 s.
14. Centrifuge with 500×g for 15 min at 4 °C.
15. Combine the supernatants.
16. Centrifuge with 45,000×g for 15 min at
4 °C.
17. Discard the supernatant and wash the pellet
twice in 5 mL lysis buffer by centrifugation
with 45,000×g for 15 min at 4 °C.
18. Dissolve the pellet in 750 mL of membranesuspension buffer and transfer into a 2-mL
reaction tube (see Note 20).
19. The protein solution can be stored at
−80 °C.
Extraction of Soluble and Insoluble
Proteins
The set of proteins isolated depends on the
method used [9, and our own published observations]. We therefore recommend applying different protein extraction methods in order to isolate
as many as possible proteins. In our experience, a
method that allows extracting both proteins of
soluble and insoluble origin in one run is helpful
to isolate proteins that cannot be captured by the
two previous methods (and vice versa). For
example, (soluble or insoluble) proteins with an
alkaline isoelectric point can efficiently be isolated with the following method, which was
adapted from Hurkman and Tanaka [10].
1. Inoculate 100 mL of respective medium
with mycelium of S. rolfsii (1/8th of a
freshly overgrown Petri dish is cut into
stripes with a scalpel) and add a sterile magnetic stirrer bar.
J. Schmid et al.
2. Incubate the flask at 28 °C with 250 rpm on a
magnetic stirrer.
3. Harvest the mycelia after 48 h of cultivation
by filtration through a piece of gauze (70 mM)
(see Note 3).
4. Wash mycelia with sterile ddH2O.
5. To remove excess water, squeeze the mycelium in the gauze (see Note 17).
6. Immediately shock-freeze mycelium in liquid nitrogen (mycelia can be stored for several weeks at −80 °C).
7. Transfer 1.5–2 g of mycelium into a prefrozen Teflon tube with 2 steel balls
(see Note 5).
8. Disrupt mycelia in a dismembrator for 2 min
(see Note 6).
9. Carefully open the Teflon tube and place it
into a ice pan filled to 2 cm with liquid nitrogen, then add 2.5 mL of Tris pH 8.8 buffered
phenol and 2.5 mL of PE-buffer.
10. Continue disruption for 1 min.
11. Transfer the suspension into a 15-mL tube
and homogenize for 30 min at 4 °C.
12. Centrifuge for 10 min with 5,000×g at 4 °C.
13. Transfer the phenol phase (top phase) into a
fresh tube and store on ice.
14. Mix the aqueous phase with 2.5 mL of Tris
pH 8.8 buffered phenol and 2.5 mL of
PE-buffer by vortexing.
15. Centrifuge for 10 min with 5,000 ´ g at 4 °C,
then combine the phenol phase with the phenol phase from the first extraction in a centrifuge tube (e.g., SS-34).
16. Precipitate proteins by adding 5 volumes of
0.1 M ammonium acetate in 100% methanol
(stored at −20 °C).
17. Vortex and incubate at −20 °C for at least 1 h
or overnight.
18. Collect the precipitate by centrifugation for
20 min with 20,000 ´ g at 4 °C.
19. Wash the pellet twice with 0.1 M ammonium
acetate in methanol, then twice with ice-cold
80% acetone (see Note 21).
20. Wash the pellet with cold 70% ethanol.
21. Store the pellet in 70% EtOH at –20 °C, or
dissolve it in 0.5–1 mL of IEF-buffer and
incubate for 1 h at RT for direct use.
39
Nucleic and Protein Extraction Methods for Fungal Exopolysaccharide Producers
Notes
1. Gloves (Nitrile) should be worn throughout
these procedures. At some point, it is advisable to sterilize gloves using a disinfectant.
Particular care should be taken when handling phenol and chloroform (consult safety
data sheets). Phenol is a corrosive agent and
a potential mutagen. All steps involving phenol and chloroform should be performed in a
fume hood.
2. We tested several cultivation media for the
extraction of pure fungal DNA with the different methods. No interference of media
components, extraction buffer, etc., was
observed. When possible and available, a
medium composition that leads to a low level
of EPS should be used.
3. A stable gauze should be used that can be
sterilized in 80% ethanol. Pore size of the
gauze is crucial for removal of the EPS and
retention of the fungal biomass. For different
fungi, different pore sizes might be tested.
Steps including gauze can be substituted by
dilution and centrifugation of the broth.
4. Liquid nitrogen is hazardous; work should
only be performed with extra gloves suitable
for low temperatures and protective glasses.
5. When using the Teflon cups for biomass
homogenization, the sterile Teflon cups (sterilized with 80% of EtOH) should be prechilled for at least 5 min in liquid nitrogen, to
guarantee that the mycelium stays frozen
during homogenization.
6. Using a Braun dismembrator, 2–3 periods of
2 min at maximum shaking frequency (2,000
min-1), followed by a 2-min resting time on
ice, have been found to be most efficient for
disruption of most samples tested. However,
optimization may be needed when testing
other fungal species. Bacteria might be
homogenized by the typical SDS method [2].
7. DNA extraction buffer can be stored for few
weeks at 4 °C; however, using fresh extraction buffer improves yields.
8. DEPC-water is prepared as follows: Add
1 mL of 0.1% Diethylpyrocarbonate (DEPC)
9.
10.
11.
12.
13.
14.
15.
16.
17.
433
to 1,000 mL distilled water. Mix well and let
set at room temperature for 1 h. Autoclave
and let cool to room temperature prior to use.
Work with the solvent isopropanol should be
performed in a fume hood.
Precipitation overnight results in higher
yields.
Drying the pellet for longer than 30 min will
result in only low solubility of the DNA
pellet.
Stirrer speed has an important influence on
mycelial morphology. Stirring at 500 rpm
results in small loosely attached mycelia with
less EPS production than stirring at 250 rpm.
Cutting the top of the pipette tip results in
lower shear forces within the cup, facilitating
more careful removal of the upper phase.
The resting time of 5 min is crucial for DNA
purification and should be kept in detail.
Gently mixing is also essential, as vortexing
will shear genomic DNA.
In case the DNA is still contaminated with
EPS, the following modifications can be
applied. See step 14 of the section: Isolation of
RNA from S. rolfsii using CsCl-Pads, and
insert these additional steps: Step14.1: Divide
the mixture into three equal parts. Step 14.2:
Add 100 mL of 5 M NaCl. Step 14.3: Add
75 mL CTAB/NaCl solution to the extraction
buffer (10% CTAB, 0.7 M NaCl) and mix thoroughly. For preparation of the CTAB/NaCl
solution, first weigh in the NaCl and add CTAB
slowly while heating the solution. Step 14.4:
Incubate the 3 samples at 65 °C for 30 min.
Continue the protocol of Isolation of RNA
from S. rolfsii Using CsCl-Pads with step 16.
Steps including harvesting and washing of
mycelia should be performed as fast as possible to prevent alteration of the transcriptional status of the fungus, which is crucial
(e.g., for a reliable transcriptome analyses).
The exopolysaccharide may swell intensively due to incorporation of water. Thereby,
the amount of mycelium per volume is
reduced significantly. To remove the excess
water after washing the mycelium, squeeze
out the mycelium in the gauze wearing sterile gloves. Keep the time for washing and
434
18.
19.
20.
21.
J. Schmid et al.
squeezing as short as possible and immediately transfer the mycelium into liquid N2.
Cutting the bottom of the tube proved to be
better for pipetting the sample.
After centrifugation the reaction tube contains
~50% clear supernatant and a fluffy pellet
containing cell debris and exopolysaccharide.
At this point, the solution will be turbid,
since the proteins are not soluble in the aqueous environment. For further applications
such as 2D-PAGE use appropriate buffer
systems.
Completely resuspend the pellet each time
by pipetting up and down. Place the resuspended sample at −20 °C for at least 15 min
between each wash.
References
1. Ausubel FH, Brent R, Kingston RE, Moore DD,
Seidman JG, Smith JA et al (1994) Current protocols
in molecular biology. Wiley, Hoboken, NJ
2. Jaufeerally-Fakim Y, Dookun A (2000) Extraction of
high quality DNA from polysaccharides-secreting
xanthomonads. Sci Technol 6:33–40
3. Shankar K, Chavan L, Shinde S, Patil B (2011) An
improved DNA extraction protocol from four in vitro
banana cultivars. Asian J Biotechnol 3:84–90
4. Sakurai K, Mizu M, Shinkai S (2001) Polysaccharidepolynucleotide complexes. 2. Complementary polynucleotide mimic behavior of the natural
polysaccharide schizophyllan in the macromolecular
complex with single-stranded RNA and DNA.
Biomacromolecules 2:641–650
5. De Kruif CG, Tuinier R (2001) Polysaccharide protein interactions. Food Hydrocolloids 15:555–563
6. Schmid J, Müller-Hagen D, Bekel T, Funk L, Stahl U,
Sieber V et al (2010) Transcriptome sequencing and
comparative transcriptome analysis of the scleroglucan producer Sclerotium rolfsii. BMC Genomics
11:329
7. Meyer V, Wedde M, Stahl U (2002) Transcriptional
regulation of the antifungal protein in Aspergillus
giganteus. Mol Genet Genomics 266:747–57
8. Chirgwin JM, Przybyla AE, MacDonald RJ, Rutter
WJ (1979) Isolation of biologically active ribonucleic
acid from sources enriched in ribonuclease.
Biochemistry 18:5294–9
9. Jacobs DI, Olsthoorn MM, Maillet I, Akeroyd M,
Breestraat S, Donkers S et al (2009) Effective lead
selection for improved protein production in
Aspergillus niger based on integrated genomics.
Fungal Genet Biol 46:S141–52
10. Hurkman WJ, Tanaka CK (1986) Solubilization of
plant membrane proteins for analysis by two-dimensional gel electrophoresis. Plant Physiol 81:802–806
Directed Evolution of a Fungal
Xylanase for Improvement
of Thermal and Alkaline Stability
40
Dawn Elizabeth Stephens, Suren Singh,
and Kugen Permaul
Abstract
Pre-treatment of paper pulps with xylanases has been shown to decrease
the amounts of toxic chlorine dioxide used to bleach pulp. Natural xylanases are unable to tolerate the extremes of pH and temperature during the
paper bleaching process and have to be genetically modified to be made
more suitable for such industrial conditions. Such modification can be
achieved using site-directed or random mutagenesis methods. Random
mutagenesis methods are more attractive because detailed information
regarding sequence or structure of the enzyme is not required. This chapter
outlines how the thermal stability and alkaline stability of a glycosyl
hydrolase family 11 cellulase-free xylanase from the fungus Thermomyces
lanuginosus were improved using two random mutagenesis methods,
error-prone PCR and a DNA shuffling method called the staggered extension process.
Keywords
Random mutagenesis • DNA shuffling • Xylanase • Thermal stability •
Alkaline stability • Error-prone PCR • Staggered extension process
Introduction
Evolution is a process occurring over eons of
time, in which the selection for specific traits is
accomplished by applying environmental pres-
D.E. Stephens (*) • S. Singh • K. Permaul
Department of Biotechnology and Food Technology,
ML Sultan Campus, Durban University of Technology,
Durban, Kwa-Zulu-Natal, 4001, South Africa
e-mail: dawnestephens@yahoo.com
sure. In nature, genetic diversity is obtained by
protracted spontaneous mutations that occur
during DNA replication or by recombination
events. Through recursive cycles of mutation,
selection and amplification, new traits accumulate in a population of organisms. Those that
provide an advantage under prevailing environmental conditions are passed from one generation to the next [1]. Man has exploited natural
evolution by using techniques such as crossbreeding in a specific manner to produce plants
and animals with useful characteristics. This
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_40, © Springer Science+Business Media, LLC 2013
435
436
form of sexual recombination is one of the most
powerful evolutionary strategies to generate
new variants. From these crossings, progeny
with improved features are chosen for additional
breeding cycles [2, 3].
Genetic and protein engineering (both rational and random) are modern laboratory techniques for increasing the robustness of proteins
for improved stability to high temperatures,
extremes of pH, oxidizing agents and organic
solvents. Cloning and expression of suitable
genes from a thermophile into a suitable and
faster-growing mesophilic host have allowed
enhanced production of the specific thermostable enzyme required for a particular biotransformation process. Rational engineering is
labour-intensive and requires precise information about enzyme structure. Random mutagenesis or directed evolution offers the fastest and
most effective means of improving biocatalysts,
provided the screening method is sensitive
enough to detect the altered properties being
screened for. During recombination, the
beneficial properties from different variant
genes are combined into a single gene, generating a protein with superior properties to its wildtype parent [4–7].
Xylanases are the predominant enzymes
responsible for the hydrolysis of plant xylans [8].
Their uses are well documented in literature [9–
12], but their predominant application has been
in the biobleaching of paper pulp [13–16]. For
xylanases to be economically feasible for pulp
application, they must be cellulase-free, and
retain stability from 60 to 90 °C at pH 8–10 for
3–5 h [17]. T. lanuginosus DSM 5826 produces
high amounts of a cellulase-free xylanase, exhibits stability over a wide pH and temperature
range, has been cloned into Escherichia coli [18]
and crystallized [19]. It is a potentially good candidate for protein engineering for pulp application. This chapter focuses on the use of random
mutagenesis and DNA shuffling to genetically
alter the xylanase from T. lanuginosus DSM 5826
D.E. Stephens et al.
to render it more temperature and alkaline stable
for future pulp application.
Materials
Cloned Gene(s) of Interest and Strains
Gene(s) of interest should be cloned onto a plasmid vector and transformed into a bacterial host
for easier mutagenesis. The vector and host (bacteria or yeast) are important issues for achieving
maximal expression of cloned genes and valuable
insight for such selection is provided [20]. The
control strain was E. coli containing a pBSK vector without the xylanase gene (xynA). An alternative vector/host expression system for high-level
expression in E. coli such as the Champion™
pET SUMO Protein Expression System is currently available from Invitrogen.
Ampicillin Preparation
1.
2.
3.
4.
5.
6.
Ampicillin-Na salt (Roche Applied Sciences).
Sterile double-distilled water.
Sterile 1.5 mL centrifuge tubes.
Vortex mixer.
10 mL sterile syringes (Braun).
0.22 mm filters (Millipore).
Luria–Bertani Medium Preparation
1.
2.
3.
4.
5.
6.
7.
8.
1.10 g/L bactopeptone.
5 g/L yeast extract.
5 g/L sodium chloride.
15 g/L technical agar (for broth, agar is
omitted).
Distilled water.
Magnetic stirrer.
Sterile petri plates
Liquid-measuring dispenser for broth tubes or
Erlenmeyer flasks.
40
Directed Evolution of a Fungal Xylanase for Improvement of Thermal and Alkaline Stability
Plasmid DNA Isolation
1.
2.
3.
4.
5.
6.
7.
8.
Sterile 1.5 mL centrifuge tubes.
Microcentrifuge.
Vortex mixer.
FastPlasmid Mini kit (Eppendorf).
Isopropanol (95–100%).
Micropipettes.
Sterile tips.
A suitable analytical instrument capable of
accurately quantifying DNA.
Agarose Gel Electrophoresis
1. Agarose (0.8%).
2. Erlenmeyer flask.
3. Sterile 1× TAE buffer (50×, 242 g Tris,
57.1 mL acetic acid, 100 mL of 0.5 M EDTA,
pH 8).
4. Microwave.
5. Gel casting tray with well-makers.
6. Gel loading buffer (0.0375 g bromophenol
blue 4 g sucrose, 1.5 mL 10% SDS, 3 mL
0.5 M EDTA in a total volume of 15 mL).
7. Electrophoresis tank and power supply.
8. Ethidium bromide (0.5 mg/mL).
9. UV transilluminator or gel documentation
system.
437
After PCR, the 786 bp PCR products were
recovered using the GFX PCR DNA and Gel
Band Purification Kit (Amersham).
Restriction Endonuclease Digestion
1. PCR products.
2. pBSK plasmid DNA.
3. EcoRI and XhoI restriction enzymes and buffers (Roche Applied Sciences).
4. Sterile 1.5 mL centrifuge tubes.
5. Sterile distilled water.
6. Microcentrifuge.
7. Vortex mixer.
8. 37 °C heating block.
9. GFX PCR DNA and Gel Band Purification
Kit (Amersham).
Ligation
1. Purified restricted vector and PCR products.
2. Sterile 1.5 mL centrifuge tubes.
3. Rapid DNA Ligation Kit (Roche Applied
Sciences).
4. Microcentrifuge.
5. Vortex mixer.
Preparation of Competent Cells
Random Mutagenesis
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
Sterile 200 mL PCR tubes.
Microcentrifuge.
Vortex mixer.
10 ng plasmid DNA (pX4).
T3 and T7 primers (0.5 mM; Integrated DNA
Technologies).
1U Taq polymerase (Roche Applied
Sciences).
Mg2+ (supplied with Taq), Mn2+, dNTPs
(Roche Applied Sciences).
Diversify random mutagenesis kit (Clontech).
1× PCR buffer (supplied with Taq).
Microcentrifuge.
PCR thermocycler (Genius-Techne). See
Table 40.1 for more details.
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
Sterile 1.5 mL centrifuge tubes.
Microcentrifuge.
Vortex mixer.
A single colony of E. coli XL1 Blue MRF’
(Stratagene).
5 mL sterile SOC medium (20 g/L tryptone,
5 g/L yeast extract, 40 mM glucose, 20 mM
NaCl, 20 mM MgCl2, 20 mM MgSO4, 5 mM
KCl) in a test tube.
37 °C shaking incubator.
29 mL SOC medium in a 50 or 100 mL
Erlenmeyer flask.
Spectrophotometer to measure cell density.
Ice.
35 mL of cold sterilized 100 mM CaCl2 containing 10% glycerol.
438
Table 40.1 Mutagenic PCR conditions used for xynA
Concentration
(mM)
Mg2+
Mn2+
dNTPs
PCR
Program
Condition
A
B
1.5
4.8
–
0.5
0.1
0.2 AG
–
0.8 CT
–
–
94 °C—1 min
35
Matsumura and
Ellington [37];
Chen et al. [38]
C
1.5
–
0.04 AG
0.2 CT
–
DIa
2
0.04
0.02 AG
0.2 CT
–
94 °C—1 min
46 °C—1 min
DII
2
–
0.02 AG
0.2 CT
0.04 dITP
72 °C—2 min
20
30
Xu et al. [39]
Eb
3.5
–
0.04 G
–
–
94 °C—1 min
46 °C—30 s
68 °C—1 min
Fb
3.5
0.64
0.04 G
–
–
72 °C—2 min
25
Diversify kit manual
Gb
3.5
0.64
0.2 G
–
–
Hb (control)
3.5
–
0.2
–
–
42 °C—1 min
No. of cycles
Reference
2 mL unpurified PCR product of DI was used as template for DII
Diversify random mutagenesis kit conditions. Special Diversify dNTP mix with unspecified concentrations was used for conditions E, F and G
a
b
D.E. Stephens et al.
40
Directed Evolution of a Fungal Xylanase for Improvement of Thermal and Alkaline Stability
Preparation of RBB-Xylan-LB Plates
for Transformation and Screening
1.
2.
3.
4.
5.
0.4% RBB-xylan.
10 g/L bactopeptone.
5 g/L yeast extract.
5 g/L sodium chloride.
15 g/L technical agar containing 100 mg/mL
ampicillin.
Transformation
1.
2.
3.
4.
5.
6.
7.
8.
9.
2 mL of the ligated DNA solutions.
150 mL SEM-competent cells.
Ice.
42 °C heating block.
800 mL of fresh SOC medium (no ampicillin)
per transformation reaction.
37 °C shaking incubator.
Hockey stick.
RBB-xylan-LB plates.
37 °C incubator.
Growth of Mutant Library and Enzyme
Extraction
1.
2.
3.
4.
5.
6.
7.
8.
9.
Clones exhibiting xylanase activity.
Sterile toothpicks.
LB-amp plates.
5 mL LB-amp broth.
37 °C shaking incubator.
Sterile 1.5 mL centrifuge tubes.
30% sterile glycerol.
Temperature-controlled centrifuge.
500 mL Bugbuster Protein Extraction Reagent
(Novagen).
10. Shaking incubator at room temperature.
Thermostability Screening Assay
1.
2.
3.
4.
5.
Xylanase supernatant.
80 °C water bath.
1.5 mL centrifuge tubes.
Ice.
Temperature-controlled centrifuge.
439
6. 1% Birchwood xylan substrate (Roth) dissolved in 0.05 M citrate buffer (pH 6.5).
7. 50 °C water bath.
8. DNS (dinitrosalicylic acid) reagent (150 g
potassium sodium tartrate, 8 g NaOH, 5 g
DNS in 500 mL distilled water).
Alkaline Screening Assay
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
0.1% Birchwood xylan (Roth).
1% Agarose.
0.05 M glycine-NaOH buffer (pH 10).
Sterile petri plates.
Ouchterlony well-maker.
60 °C incubator.
1% Congo Red.
1 M NaCl.
60 °C water bath.
1% Birchwood xylan substrate (Roth) dissolved in 0.05 M citrate buffer (pH 6.5).
11. 50 °C water bath.
12. DNS reagent.
Long-Term Thermal and Alkaline
Stabilities
Growth and Extraction
1. 5 mL LB-amp broth per selected mutant
strain.
2. 37 °C shaking incubator.
3. 300 mL LB-amp medium.
4. Spectrophotometer for measuring cell density
5. 1 mM filter-sterilized IPTG (isopropyl-b-dthiogalactoside pyranoside).
6. Centrifuge.
7. 50 mL centrifuge tubes.
8. 150 U DNase I (Roche Applied Sciences).
9. 2 g Lysozyme (Roche Applied Sciences).
10. 120 mL Breaking buffer solution (6.80 g/L
KH2PO4, 0.61 g/L MgCl2·6H2O, 0.77 g/L
dithiothreitol, 0.37 g/L EDTA, 0.10 g/L
phenylmethylsulfonylfluoride).
Stability
1. Xylanase-containing supernatant.
2. 80 °C water bath.
440
3.
4.
5.
6.
7.
8.
9.
10.
D.E. Stephens et al.
1.5 mL centrifuge tubes.
Ice.
Temperature-controlled centrifuge.
1% birchwood xylan substrate (Roth) dissolved in 0.05 M citrate buffer (pH 6.5).
50 °C water bath.
DNS (dinitrosalicylic acid) reagent (150 g
potassium sodium tartrate, 8 g NaOH, 5 g
DNS in 500 mL distilled water).
0.05 M glycine-NaOH buffer (pH 10).
60 °C water bath.
DNA Recombination: Staggered
Extension Process
1.
2.
3.
4.
5.
6.
7.
8.
Overnight cultures of G41 and G53 mutants.
FastPlasmid Mini Kit (Eppendorf).
10 ng plasmid DNA.
1.5 mM MgCl2.
0.5 mM each T3 and T7 primers.
1× PCR buffer and 1 U Taq.
640 mM MnSO4, 0.2 mM dGTP.
Diversify Random Mutagenesis Kit (Clontech).
DNA Sequence Analysis
1.
2.
3.
4.
Automated DNA sequencer.
Chromas Lite software.
ExPASy protein translation tools.
CLUSTALW sequence alignment.
Methods
efficiency of an ep-PCR reaction, and facilitate
easier sequencing. The modified plasmid was
renamed pX4 [21].
Ampicillin Preparation
A 5 mL stock solution of 100 mg/mL of ampicillin was aseptically made using sterile distilled
water and filter sterilized with 0.22 mm filters.
The solution was aliquoted in sterile 1.5 mL centrifuge tubes and stored at −20 °C. For use, the
ampicillin was added to the growth media to a
final concentration of 100 mg/mL. Note that
ampicillin is never autoclaved.
LB Medium Preparation
LB medium supplemented with ampicillin (see
previous) was used for plating of the E. coli harbouring the plasmid with the cloned gene of
interest. This medium was used to maintain all
recombinant strains. LB media components were
combined, dissolved and sterilized. When the
medium cooled, the prepared ampicillin was
added and the medium was mixed gently (to
avoid creation of air bubbles). Agar plates were
then aseptically poured, allowed to set and stored
at 4 °C.
LB medium (without agar) can be made up
in bulk, then dispensed in tubes or
Erlenmeyer flasks (depending on final use) and
sterilized. They can be stored at 4 °C or room
temperature and ampicillin can be added just
prior to use.
Cloning
The xylanase gene used for this research, xynA of
T. lanuginosus, was cloned into a plasmid
Bluescript SK vector (pBSK) using the l-ZapII
cloning system (Strategene) and transformed into
E. coli [18]. This plasmid was called pX3. The
earlier strategy for cloning the xylanase involved
creating a cDNA library, so the xynA gene was
present on a larger insert. The cloned insert was
thus reduced in size to increase the mutational
Plasmid Isolation
E. coli cultures with plasmids were inoculated in
5 mL LB medium containing 100 mg/mL ampicillin and grown for 12–16 h at 37 °C in a shaking
incubator at 150 rpm. Cells were harvested by
centrifugation of broth cultures at 5,000×g for
5 min. For other details refer to the FastPlasmid
Mini Kit (Eppendorf) user manual.
40
Directed Evolution of a Fungal Xylanase for Improvement of Thermal and Alkaline Stability
Measurement of DNA Concentration
This was estimated spectrophotometrically at
260 nm and calculated on the premise that an
absorbance of 1 at OD260 nm corresponds to 50 ng
DNA/mL. Purity of the DNA sample was estimated using the absorbance ratio measured at
both OD260 nm and OD280 nm. A ratio greater than
1.7 is considered ideal purity for procedures like
DNA sequencing. Currently, many instruments
capable of accurately quantifying DNA are
available.
Agarose Gel Electrophoresis
The desired amount of agarose was placed in an
Erlenmeyer flask together with the required
amount of 1× TAE buffer, which was diluted
from the 50× TAE stock. The contents of the flask
were then microwaved for 1 min and poured into
a casting tray with well combs and allowed to set.
Gel loading buffer was added to the DNA samples in a ratio of 1:5, which was then loaded into
the agarose gel wells. Samples were run alongside a DNA molecular weight marker at 90 V for
approximately 1 h. Gels were then stained in
ethidium bromide for 20 min and destained in
distilled water for a further 5–10 min. Care must
be taken to strictly adhere to the recommended
concentration for ethidium bromide staining as
high concentrations can reduce ligation efficiency
of target DNA. Stained gels were then viewed on
a UV transilluminator and the band sizes compared to the DNA molecular weight marker.
Ethidium bromide is a known mutagen and must
be handled with extreme precaution. It is safer to
purchase a ready-made stock to minimize handling [20, 22].
Random Mutagenesis
A broad scope of conditions (see Table 40.1)
ranging from previously published research as
well as the use of a commercial mutagenesis kit,
Diversify Random Mutagenesis Kit (Clontech),
were investigated for mutagenesis of xynA.
441
Commercial mutagenesis kits offer the advantage
of being able to theoretically control the number
of mutations per gene. Plasmid pX4 served as the
template DNA for all mutagenic conditions
tested. The final volume of each PCR reaction
was always 50 mL. Five microlitres of each PCR
product was analysed by agarose gel electrophoresis to determine if the target DNA was successfully amplified. Lighter bands generally imply a
high mutagenic rate in ep-PCR since the Taq
polymerase does not amplify optimally.
Therefore, it cannot form as much product compared to PCR under normal conditions. This represents an early, visible indication of mutation in
the target gene. The remaining PCR products
were recovered after agarose gel elctrophoresis.
The gel was stained for not more than 10 min in
ethidium bromide and then viewed on the UV
transilluminator whilst the PCR-gel bands were
quickly excised. Ethidium bromide and UV light
are mutagens and prolonged contact with the
PCR products can cause irreversible DNA damage and also drastically reduce ligation efficiency.
The GFX PCR DNA and Gel Band Purification
Kit (Amersham) was used according to the manufacturer’s instructions. Many such kits are available for commercial use.
Restriction
Standard protocols were followed for restriction
analysis [20, 22]. Restriction analysis is necessary to create compatible sticky ends to facilitate
ligation of the PCR product into the vector. Note
that both the vector and PCR products must be
restricted separately, and with the same combinations of restriction enzymes. If using more than
one restriction enzyme, it is important to use buffers compatible with both restriction enzymes.
Buffer compatibility charts are usually included
with the restriction enzymes. For digestion, the
following volumes were added to a sterile 1.5 mL
centrifuge tube: 5 mL DNA, 2 mL sterile water,
1 mL buffer H, 1 mL EcoRI and 1 mL XhoI. The
tubes were vortexed and spun down in a minicentrifuge for a few seconds and then incubated in
a 37 °C heating block for 1 h. For notes on
442
restriction digestion, refer to references [20, 22].
Each restriction reaction was purified from solution using the GFX PCR DNA and Gel Band
Purification Kit (Amersham) according to the
manufacturer’s instructions.
Ligation
T4 DNA ligase was used to join the DNA between
the 5¢-phosphate and the 3¢-hydroxyl-group of
adjacent nucleotides of the vector and PCR
inserts. For the creation of recombinant libraries,
a molar vector, insert ratio of 1:3 was used to
guarantee high ligation efficiency. Ligations were
carried out using the Rapid DNA Ligation Kit
(Roche Applied Sciences) according to the manufacturer’s instructions and were directly transformed into competent E. coli cells.
Preparation of Competent Cells
Host cells were made “competent” or capable of
taking up DNA from their surrounding environment, by exposing them to Ca2+, which interacts
with their cell envelopes. E. coli host cells were
made competent prior to transformation [23].
E. coli XL1 Blue was cultured on LB medium
without ampicillin. A single colony was used to
inoculate 5 mL sterile SOC medium (20 g/L tryptone, 5 g/L yeast extract, 40 mM glucose, 20 mM
NaCl, 20 mM MgCl2, 20 mM MgSO4, 5 mM
KCl), incubated at 37 °C and shaken overnight.
One millilitre of this culture was used to inoculate 29 mL of fresh SOC medium and was shaken
at 150 rpm at 37 °C until it reached an OD600 nm of
0.5. At this stage of growth, E. coli cells have
maximum ability to uptake DNA. The culture
was immediately placed on ice and kept cold for
the duration of the procedure. Cold conditions
suspend the continued growth of the cells and
preserve their viability for subsequent transformation. The cells were pelleted at 5,000×g for
10 min and the supernatant was discarded. The
cells were resuspended in 10 mL cold 100 mM
CaCl2, recentrifuged at the same speed and resuspended in 10 mL of cold 100 mM CaCl2. The
D.E. Stephens et al.
entire mixture was incubated on ice for 20 min
and then centrifuged. The competent cells were
subsequently resuspended in 2 mL 100 mM
CaCl2 containing 10% glycerol. One hundred and
fifty microlitres of the prepared competent cells
were dispensed into eppendorfs, stored at 4 °C
overnight, and then maintained at −70 °C, after
snap-freezing in liquid nitrogen. According to the
protocol followed [23], it is postulated that SEMcompetent cells are most efficient when prepared
24 h prior to transformation. The preparations
were therefore incubated at 4 °C overnight to
enhance the effectiveness of the subsequent
transformation procedure.
Preparation of RBB-Xylan-LB Plates
for Transformation and Screening
Mutation sometimes produces clones that contain
the gene of interest; however, it is often so heavily mutated that an active, fully functional protein
is no longer produced [24, 25]. Also, mutations
are more likely to destroy enzyme activity rather
than enhance it. Thus, Remazol Brilliant Blue
dye was linked to birchwood xylan for detection
of xylanase producers during the transformation
process [26, 27]. Only clones capable of producing a fully functional xylanase would be able to
break the linkage between the xylan and the RBB
dye to produce colourless xylooligosaccharides
which was visible as a zone of hydrolysis around
the colony. RBB-xylan was added to LB medium
components, dissolved and sterilized. After cooling to just below 50 °C, filter-sterilized ampicillin was added to the medium and mixed gently
(to avoid creation of air bubbles). Agar plates
were then aseptically poured, allowed to set and
stored at 4 °C.
Transformation
Two microlitres of the ligated DNA solutions were
added to 150 mL of the SEM-competent cells and
incubated on ice for 30 min and thereafter subjected to heat shock for 30 s at 42 °C. An increase
in temperature allows for the creation of transient
40
Directed Evolution of a Fungal Xylanase for Improvement of Thermal and Alkaline Stability
pores that allow for the uptake of DNA attached to
the cell surfaces. Rapid addition of 800 mL of fresh
SOC medium to each of the mixtures followed the
heat shock procedure and they were then shaken
at 37 °C for 1 h. SOC medium is rich in salts and
ions that facilitate the rapid regeneration of the
cell wall. No ampicillin is added to the SOC
medium to prevent additional metabolic stress on
the cells. The cells actively divide and the ligated
products (plasmids) are also amplified. Any uncircularized (or unligated) DNA is degraded by cell
DNases. The transformation mixtures were diluted
1:1 with fresh SOC medium and 100 mL was
plated on RBB-xylan-LB containing ampicillin
and incubated at 37 °C overnight.
Growth of Mutant Library and Enzyme
Extraction
Generally, as the degree of mutagenesis in epPCR reactions increases, the number of transformants obtained also decreases because less DNA
is amplified during the ep-PCR reactions. All
positive transformants, that is, clones exhibiting
xylanase activity, were picked with a sterile
toothpick and initially streaked onto an LB plate
containing 100 mg/mL ampicillin. Subsequently,
a single colony from the overnight plate was
inoculated into 5 mL LB broth containing 100 mg/
mL ampicillin and incubated for 12–16 h at 37 °C
in a shaking incubator at 150 rpm. One hundred
and fifty microlitres of these overnight cultures
were placed in sterile 1.5 mL centrifuge tubes
containing 30% sterile glycerol. These tubes were
stored at −20 °C and served as the master mutant
library stock. Since the E. coli transformant host
is a prokaryote and the xylanase is eukaryotic,
most of the xylanase is located intracellularly and
cannot be properly secreted. It must be lysed to
release the functional xylanase protein.
Mechanical lysis methods such as the use of a
French press or sonicator are unsuitable since
they result in significant heating of the sample
which can denature the xylanase. A gentle chemical lysis agent such as the BugbusterProtein
Extraction Reagent (Novagen) is much more
effective. The use of Bugbuster, however, is
uneconomical for large-scale cell lysis.
443
The rest of the culture was centrifuged at
5,000×g and the media discarded. The pellets
were resuspended in 500 mL cold Bugbuster
reagent. Pellet resuspension must be gentle to
minimize foaming and heat generation. The suspensions were shaken gently at 40 rpm at room
temperature for 20 min to gently lyse E. coli cells
and release the mutant xylanases. The lysates
were then centrifuged at 15,000×g at 4 °C to
remove cell debris. The resulting pellets were
discarded whilst the supernatants were stored at
4 °C for further analysis.
Thermostability Screening Assay
The clear lysate obtained after enzyme extraction
contained the crude xylanase and was used to test
the thermostability of the xylanase variants. The
protocol followed was a combination of the methods used [28, 29]. A temperature of 80 °C was
chosen for screening possible thermostable xylanase variants since XynA is documented to be
stable up to 70 °C [18].
A short-term screen time of 40 min at 80 °C
was used to decrease the number of mutants
screened later. Prior to incubation at 80 °C in a
water bath, 0 min (untreated) samples were
removed from the clear cell lysates and placed on
ice. The crude enzymes were subsequently heated
for 40 min, chilled on ice for 15 min and incubated for 30 min at room temperature to prevent
low-temperature denaturation of the enzymes.
The samples were centrifuged and the supernatants assayed for residual xylanase activity [30].
Activities of the 0 min samples were considered
as 100%, and activities of the 40 min incubation
time were expressed as percentages of the
untreated sample to determine the percentage of
residual activity after heat treatment. The wildtype XynA served as the control.
Alkaline Screening Assay
Alkaline screening was performed in two parts,
viz., a plate assay and a liquid assay that tested
stability at pH 10. Wells were punched (using an
Ouchterlony well-maker) into plates containing
444
0.1% birchwood xylan and 1% agarose made
with 0.05 M glycine–NaOH buffer (pH 10). Ten
microlitres of each crude enzyme was inoculated
into the wells, with one well containing the control XynA on each screening plate. The plates
were incubated at 60 °C for 2 h and then stained
with 1% Congo Red for 25 min. Excess dye was
flushed off with 1 M NaCl for up to 1 h, until
zones of hydrolysis were clearly visible [31, 32].
The supernatants of mutants displaying larger or
more distinct zones than XynA were diluted in
0.05 M glycine-NaOH buffer (pH 10) and incubated for 40 min in a 60 °C water bath and residual activities of the enzymes were determined.
A temperature of 60 °C was used for detection of
alkaline-stable mutants to effectively trim down
the number of mutants screened during longer
incubation periods.
Long-Term Thermal and Alkaline
Stabilities
Growth and Extraction
Mutants displaying more than 60% residual activity after heat and alkaline treatment were selected
for long-term stability testing. These mutants
were inoculated into 5 mL LB-amp broth and
incubated overnight at 37 °C at 150 rpm. One
millilitre of this culture was used to inoculate
300 mL LB-amp medium and shaken at 37 °C
until the OD600 nm of all flasks reached 0.5 absorbance units. The cells were induced for xylanase
production since the pBSK vector has a LacZ
promoter. This means that lactose would induce
the LacZ promoter, and thus leads to expression
of the cloned xynA. IPTG is a lactose analogue,
which constitutively induces the LacZ promoter,
but is not metabolized. IPTG is added to the flasks
and shaken overnight at 37 °C. Thereafter, the
samples were centrifuged at 5,000×g with each
50 mL pellet resuspended in 2 mL of cold lysis
solution. Bugbuster reagent (used previously for
large-scale screening) is unstable for long-term
incubation at high temperatures. Thus, a lysis
solution was designed to eliminate this problem.
First, a breaking buffer solution was made by dissolving 0.37 g/L EDTA in 200 mL distilled water,
D.E. Stephens et al.
followed by the addition of 6.80 g/L KH2PO4,
0.61 g/L MgCl2.6H2O and 0.77 g/L dithiothreitol.
The pH of the solution was brought to 6.8. After
autoclaving and cooling of the breaking buffer
solution, 0.10 g/L phenylmethylsulfonylfluoride
was added. The lysis solution comprised 150 U
DNase I, 2 g lysozyme in 120 mL breaking buffer
solution. The suspensions were left at 4 °C overnight and then centrifuged at 15,000×g. The
supernatant lysates containing the enzymes were
stored at 4 °C until further use.
Stability
For determinations of long-term thermal stability,
samples were incubated at 80 °C and samples
were removed every 15 min for 90 min, incubated
on ice and then assayed for residual xylanase
activity as previously described. For determination of long-term alkaline stability, samples were
diluted in 0.05 M glycine-NaOH buffer (pH 10),
incubated at 60 °C and samples were removed
every 15 min for 90 min, incubated on ice and
then assayed for residual activity. Activities of
each mutant were expressed as percentages of the
0 min sample.
DNA Recombination: Staggered
Extension Process
In this study, highly thermostable xylanases displayed poor catalytic activities, whereas the opposite was found for alkali-stable xylanases. Thus,
the two best thermostable and alkaline-stable xynA
variants were good candidates for DNA recombination for incorporation of both properties to create a single, robust xylanase. The recombination
method used in this study was a modification of
the staggered extension process (StEP) reaction
[33]. Advantages of the StEP method include the
following: (1) it can be performed using a pair of
flanking primers in a single PCR tube; (2) separation of parent templates from the recombined
products is not necessary; and (3) higher recombination frequencies between highly homologous
templates can be achieved. Disadvantages include
that there is a smaller probability of generating
enzymes with unique functions and the sometimes
40
Directed Evolution of a Fungal Xylanase for Improvement of Thermal and Alkaline Stability
non-specific annealing and formation of undesirable recombinants. Other shuffling methods have
been reviewed and there are benefits and disadvantages of each method [34].
The G41 and G53 plasmids were isolated using
the FastPlasmid Mini Kit (Eppendorf). Thermal
stability was considered more important than
alkaline stability; thus, twice the amount of G41
plasmid was added to each reaction as compared
with G53. The final concentration of DNA was
maintained at 10 ng. StEP recombination PCR
reactions contained 10 ng total DNA, 1.5 mM
MgCl2, 0.5 mM each T3 and T7 primers, 1× PCR
buffer and 1 U Taq. Eighty cycles of denaturation
for 30 s at 94 °C followed by annealing for 4 s at
42 °C were carried out on both the reactions. The
recombination products were subsequently cloned
and xylanase activity was tested as previously
described. The study resulted in the creation of
recombinant xylanases that exhibited properties
of both parent, G41 and G53, xylanases [35].
DNA Sequence Analysis
DNA sequencing was carried out to determine
the mutations that were responsible for the
observed changes in stabilities of some xylanase
variants. Many possible automated DNA sequencing methods are currently available. Raw DNA
sequencing data were initially processed using
the Chromas Lite software package (Technelysium,
version 2.0) (www.technelysium.com.au) and
both DNA strands were edited to yield complete
gene sequences. The DNA sequences were then
translated into their protein counterparts using
the Translate tool from the ExPASy Website
(www.expasy.org/tools/dna.htmL) and aligned to
the wild-type parent using the CLUSTALW (version 1.81) alignment program on the GenomeNet
server (www.clustalw.genome.ad.jp). The results
showed that most substitutions occurred in the
b-sheet of xynA. Studies of family 11 xylanases
indicate that this long b-sheet is responsible for
thermostability, because it stabilizes the overall
xylanase structure [36]. Also, noteworthy was the
observed increase in arginine content of the most
stable xylanase variants.
445
Acknowledgements Ms. Siphi Dlungwane is duly
acknowledged for providing technical assistance. The
National Research Foundation of South Africa funded this
research.
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Genome Shuffling Protocol
for the Pentose-Fermenting
Yeast Scheffersomyces stipitis
41
Paramjit K. Bajwa, Nicole K. Harner,
Terri L. Richardson, Sukhdeep Sidhu,
Marc B. Habash, Jack T. Trevors, and Hung Lee
Abstract
This chapter presents the protocol for genome shuffling based on recursive
cross-mating in the pentose-fermenting yeast Scheffersomyces (Pichia)
stipitis. Genome shuffling involves two stages. In the first stage, a pool of
mutants with improved phenotypes is selected. Several rounds of random
mutagenesis can be done using different mutagens, and mutant selection
can be based on different criteria to generate different mutant cell lines. In
the second stage, the genomes of mutants derived from different lines are
mated recursively to allow for genetic recombination, followed by screening after each mating cycle to select for improved phenotypes in the
recombinants. A number of reports have described genome shuffling based
on recursive protoplast fusion in bacteria and yeasts. Recently, we developed mating-based genome shuffling in the pentose-fermenting yeast S.
stipitis. We have used this approach to obtain genetically stable mutants of
S. stipitis with considerably improved tolerance to hardwood spent sulphite liquor (HW SSL), a pulping waste liquor containing a complex mixture of inhibitory substances. This was achieved in the complete absence
of knowledge as to the precise genetic modifications needed to confer HW
SSL tolerance. Here we describe the protocols for recursive UV mutagenesis, cross-mating, sporulation and isolation of recombinants with
improved phenotypic traits.
Keywords
Genome shuffling • Random mutagenesis • Recursive cross-mating •
Pentose fermentation • Scheffersomyces stipitis
P.K. Bajwa • N.K. Harner • T.L. Richardson •
S. Sidhu • M.B. Habash • J.T. Trevors • H. Lee (*)
School of Environmental Sciences, University of Guelph,
50 Stone Road East, Guelph, ON, Canada N1G 2W1
e-mail: hlee@uoguelph.ca
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_41, © Springer Science+Business Media, LLC 2013
447
448
Introduction
Genome shuffling is a microbial strain improvement technology that involves the generation of a
pool of mutants with improved phenotypes, followed by iterative recombination between their
genomes. This approach offers a number of advantages compared to the classical and molecular
methods of microbial strain improvement.
Classical strain improvement using mutagenesis
and selection is time-consuming, laborious and
based on a single starting strain. Molecular
approaches of strain improvement are only applicable to genes that can be isolated, along with
some knowledge of what genetic modifications
are needed to produce the desired phenotypic
effect. To improve a cell’s complex phenotype,
such as stress tolerance, likely requires modification
in a number of known and unknown genes; hence,
the molecular methods are inadequate. Genome
shuffling is particularly suitable for the engineering of complex multi-genic phenotypic traits that
are difficult to modify by either the classical or
molecular strain improvement approaches, as it
does not require a priori knowledge of the set of
genes to be modified to confer beneficial changes.
Genome shuffling was originally developed
for bacteria and then extended to yeasts. In bacteria, genome shuffling has been used successfully
to increase the tylosin titer in Streptomyces fradiae [1], lactic acid tolerance by Lactobacillus
[2], and pentachlorophenol degradation and tolerance by Sphingobium chlorophenolicum [3]. In
yeasts, particularly Saccharomyces cerevisiae,
genome shuffling has been used to improve thermotolerance, ethanol productivity, ethanol tolerance and acetic acid tolerance [4–7].
In genome shuffling, genome recombination is
carried out using either recursive protoplast fusion
or cross-mating. Most of the reports on genome
shuffling are based on protoplast fusion [1–5].
We have developed a genome shuffling protocol
based on recursive mating in the pentose-fermenting yeast Scheffersomyces stipitis and isolated
mutant strains with improved tolerance to hardwood spent sulphite liquor (HW SSL) [8]. This
approach led to a rapid improvement in tolerance
P.K. Bajwa et al.
to HW SSL in the selected strains, which retained
their growth and fermenting ability. In our
approach, the haploid cells from a pool of S. stipitis mutants are mated to form diploid zygotes. In
malt extract agar, the diploid zygotes undergo
meiosis to form two-spored asci [8, 9]. The ascus
wall dissolves on its own to release the hat-shaped
ascospores. When transferred to a rich medium,
the ascospores germinate to produce vegetative
haploid cells. The recombinants with improved
phenotypes are identified and selected using suitable selection regimes (Fig. 41.1). The process of
mating and sporulation is repeated several times
to allow for recombination between multiple parents and the pooling of beneficial alleles from
various genomes.
Materials
Random Mutagenesis
1. A physical (ultraviolet light) or a chemical
(e.g., N-methyl-N¢-nitro-N-nitrosoguanidine
or ethylmethanesulphonate) mutagen.
2. S. stipitis culture grown for 48 h in minimal
medium broth (0.67% [w/v] yeast nitrogen
base (YNB) without amino acids and 2%
[w/v] xylose).
3. Sterile Petri dishes, 15-mL centrifuge tubes
and a tube rotator.
4. Selective agar (1.5% [w/v] medium).
5. Square plastic Petri dishes (120 × 120 × 17 mm).
Yeast Mating and Genome Shuffling
1. Two auxotrophic yeast strains with different
nutritional requirements.
2. YEPD broth (1% [w/v] yeast extract, 2% [w/v]
peptone, 2% [w/v] glucose).
3. Minimal medium plates (0.67% [w/v] YNB
without amino acids, 2% [w/v] xylose, 1.5%
[w/v] agar).
4. Malt extract (ME) agar plates (3% [w/v] ME,
1.5% [w/v] agar).
5. Sterile Petri dishes, 15-mL centrifuge tubes
and a tube rotator.
41
Genome Shuf fling Protocol for the Pentose-Fermenting Yeast Scheffersomyces stipitis
Fig. 41.1 Schematic of
genome shuffling in yeasts
449
450
Methods
Random Mutagenesis Procedure
Random mutagenesis can be carried out using a
physical or chemical mutagen. Freshly grown
yeast cells are exposed to the mutagen for a predetermined time. The cells are then allowed to
recover overnight in liquid or solid medium followed by screening and selection of the mutant
colonies in a selective medium. In the following,
we describe random mutagenesis using UV light.
1. Transfer a loopful of S. stipitis cells from an
isolated colony on YEPD agar plate to 20 mL
of broth containing 0.67% [w/v] YNB without
amino acids and 2% [w/v] xylose in a 125-mL
Erlenmeyer flask at 28 ± 1 °C with shaking at
180 rpm for 48 h.
2. Aseptically transfer 1 mL of the 48-h grown
yeast culture (OD600 between 8 and 10) to an
empty sterile Petri dish.
3. Place the Petri dish under the UV light source
at a distance of about 40 cm.
Fig. 41.2 Test tube rotator
P.K. Bajwa et al.
4. Remove the Petri dish cover and turn on the
UV light. Prepare a yeast survival curve based
on length of UV exposure. For S. stipitis wildtype (WT) cells, the length of UV exposure
tested ranged from 10 to 60 s and 50% survival was achieved at 20 s.
5. Transfer the UV-irradiated culture to a 15-mL
sterile centrifuge tube covered with aluminium
foil and incubate in a tube rotator (Fig. 41.2)
for 24 h at 23 ± 1 °C. The tubes are rotated
about their vertical axis at about 60° from horizontal at 90 cycles per min as previously
described [10, 11].
6. Spread 100 mL of the UV-exposed culture on
several selective medium plates and incubate
at 28 ± 1 °C for 4–5 days.
7. Isolate the mutant colonies appearing on the
selective medium plates and maintain them on
YEPD agar plates.
8. Individual mutants are further tested in
liquid broth. Mutants that retain good fermenting ability in liquid broth were selected for
further rounds of mutagenesis and genome
shuffling.
41
Genome Shuf fling Protocol for the Pentose-Fermenting Yeast Scheffersomyces stipitis
451
Fig. 41.3 Schematic of UV mutagenesis followed by screening on gradient plates to select for HW SSL tolerant
mutants
In our earlier study [12], we used three consecutive rounds of UV mutagenesis followed by
screening to obtain mutants of S. stipitis NRRL
Y-7124 with improved tolerance to HW SSL. A
schematic of the random mutagenesis procedure
is presented in Fig. 41.3. HW SSL is a waste
liquor from the Tembec pulp and paper mill in
Témiscaming, Québec, Canada. It contained, in
[w/v]: 0.076% arabinose, 0.25% galactose, 0.33%
glucose, 0.55% mannose, 2.2% xylose, 1% acetic
acid, 0.18% furfural and 0.11% hydroxymethyl
furfural. Prior to use, the pH of HW SSL was
increased from 2.5 to 5.5 with 10 M NaOH. The
liquor was then boiled for 5 min in a microwave
oven, followed by gradual cooling to room
temperature.
For UV mutagenesis, the UV exposure time
was optimized to achieve a survival rate of about
50%. We hypothesized that changes in many
genes are needed to collectively confer tolerance
to HW SSL in yeasts. Thus, we used a low UV
dosage resulting in a high survival rate in order to
maximize those surviving populations carrying a
small number of mutations for screening. Higher
UV doses resulting in a lower survival rate may
result in many members of the surviving popula-
tions carrying multiple mutations, some of which
may not be beneficial. The presence of
nonbeneficial mutations will likely mask the
effect of the beneficial mutations, resulting in
these mutants not being selected in the screen.
An increasing concentration of HW SSL was
used as the selective agent for initial mutant
screening on agar plates after UV mutagenesis.
This was done by spreading 100 mL of the
UV-exposed culture on HW SSL (pH 5.5) gradient plates. The plates were incubated at 28 °C for
5–10 days. Colonies growing at a higher concentration of HW SSL as compared to the WT strain
were isolated and maintained on YEPD plates.
HW SSL gradient plates were prepared by successively pouring two layers of agar into a square
plastic Petri dish (120 × 120 × 17 mm). The bottom
layer consisted of 25 mL of HW SSL agar medium
(Fig. 41.4a). The plate was allowed to solidify at a
slightly inclined position. The Petri dish was then
placed in a horizontal position and a second layer
of plain agar was poured over the first layer
(Fig. 41.4b). The Petri dish was incubated at 30 °C
for 2 days before use. This allowed diffusion of
the HW SSL components through the agar layer,
thereby establishing a concentration gradient.
452
P.K. Bajwa et al.
The first layer of agar with HW
SSL is poured into the Petri dish
and allowed to solidify while the
plate is held in an inclined
position
The second layer of plain agar
is poured into the Petri dish
and allowed to solidify while
the plate is held in a leveled
position
Fig. 41.4 Preparation of HW SSL gradient agar plate
Because performance of the mutants on the
plate screen may not translate to the broth, the
mutants were further tested for growth in liquid
HW SSL. Also, since our end goal is fermentation of the sugars in the waste liquor, the fermenting ability of the mutants needs to be verified by
screening in broth. Therefore, the mutants with
improved HW SSL tolerance were assessed for
the ability to ferment xylose, glucose, mannose,
galactose and arabinose in defined media as well
as HW SSL. Six improved mutants obtained after
several rounds of UV mutagenesis and screening
were used as the starting strains for genome
shuffling.
Optimization of Yeast Mating
and Genome Shuffling
1. Grow each auxotrophic yeast strain in 5 mL of
YEPD broth in a 15-mL centrifuge tube at
28 °C with shaking at 180 rpm for 24 h.
Auxotrophic yeast strains can be obtained by
random mutagenesis [13].
2. Mix 2 × 108 cells of each auxotrophic strain
together and spread the mixed cell suspension
on ME plates. Also, spread 2 × 108 cells of
each auxotroph separately on ME plates as
controls.
3. Incubate the ME plates at 28 °C for 7–10 days
to allow mating and sporulation to occur.
Examine the cells periodically under the
microscope to follow spore formation. S. stipitis forms two hat-shaped spores per ascus.
4. After sporulation, scoop out all the cells from
each plate. Suspend them in sterile water, centrifuge and wash the cells several times with
sterile water. The cells are then suspended in
5 mL of sterile water.
5. Spread the cells on minimal media plates. The
auxotrophic strains cannot grow on these
plates. Only the recombinant cells resulting
from mating between the two auxotrophic
strains can grow on minimal media plates.
6. Incubate the plates at 28 °C for 4–5 days.
Enumerate the colonies appearing on minimal
media plates and determine the mating
frequency.
7. To check for the stability of recombinants,
repeatedly streak the colonies derived from
the mating on minimal media plates.
In our lab, the mating and sporulation protocol
was optimized using two auxotrophic strains of
S. stipitis (FPL Y14—ura3, ade2, met1 and FPL
Y18—ura3, leu2 delta). Each parent carried a
common auxotrophic requirement for uracil in
addition to the other different auxotrophic markers. If mating and successful recombination have
occurred, then the recombinants would require
only uracil. Therefore, the sporulated cells were
spread on uracil-containing minimal media plates
(0.67% w/v YNB w/o amino acids, 2% w/v xylose
and 0.005% w/v uracil). Only the recombinant
cells could grow on these plates (Fig. 41.5). The
mating efficiency of the haploid S. stipitis auxotrophs was estimated to be 0.05% after one
round of mating and sporulation. The estimate
was based on the number of recombinants
obtained on uracil-containing plates after mating
the two auxotrophic S. stipitis strains. The optimized protocol was used for genome shuffling of
UV-induced mutants of S. stipitis.
Six HW SSL-tolerant mutants obtained after
the third round of UV mutagenesis served as
the parental strains for genome shuffling. The
41
Genome Shuf fling Protocol for the Pentose-Fermenting Yeast Scheffersomyces stipitis
453
Fig. 41.5 Growth of the sporulated cell suspensions
obtained from individual auxotrophs and mated populations on minimal media plates + uracil. Sectors 1 and 2
represent the growth of individual auxotrophs FPL Y14—
ura3, ade2, met1 and P. stipitis FPL Y1—ura3, leu2 delta,
respectively. Sector 3 represents the growth of sporulated
cells obtained after mating of the two auxotrophs
Fig. 41.6 Growth of S. stipitis WT (left), UV-induced
mutant (PS302, middle) and genome shuffled strain
(GS401, right) on HW SSL (pH 5.5) gradient plate
selected mutants were individually grown overnight in YEPD broth followed by mixing of
1 × 108 cells of each mutant on ME plates. The
plates were incubated for 7 to 10 days to allow
mating and sporulation to occur in the same way
as described above for the auxotrophic strains.
All the cells were scooped out from the ME plates
and suspended in 5 mL of sterile water. After
centrifugation and several washings with sterile
water, the spore suspension was transferred to
20 mL of fresh YEPD broth in 125-mL Erlenmeyer
flask and incubated at 28 °C with shaking
(180 rpm). After 48 h of growth, 100 ml of cell
suspension was spread on several HW SSL (pH
5.5) gradient plates. Colonies appearing at a
higher concentration of HW SSL as compared to
the best UV-induced mutant were isolated. Three
consecutive rounds of genome shuffling involving mating, sporulation and selection on HW SSL
plates were done. Improved mutants obtained
from one round served as the starting strains for
the next round of genome shuffling. Since we
started with six UV-induced mutants, three rounds
of genome shuffling were deemed sufficient to
recombine all the beneficial mutations together.
A greater number of rounds would be desirable if
we had started with a larger mutant pool.
Figure 41.6 illustrates the growth of S. stipitis
WT, UV-induced mutant and a genome shuffled
mutant on the HW SSL gradient plate. The
genome shuffled strain (GS401) was clearly more
tolerant than the UV-induced mutant (PS302),
which in turn was more tolerant than the WT, on
HW SSL gradient plate. The isolated mutants
were further tested for growth in liquid HW SSL
to confirm their HW SSL tolerant character. The
WT was unable to grow in HW SSL unless diluted
to 65% (v/v) or lower. The UV-induced mutants
grew in 75% (v/v) HW SSL. Two mutants
obtained after three rounds of genome shuffling
grew in 85% (v/v) HW SSL and one of these
could survive in 90% (v/v) HW SSL, although no
increase in cell number was seen [8]. The genome
shuffled mutants consumed 4% [w/v] xylose or
glucose in defined media more efficiently and
produced more ethanol as compared to the S.
stipitis WT strain. These mutants also utilized
mannose and galactose and produced the same
amount of ethanol as the WT.
454
Summary
The results from our research demonstrated the
utility of genome shuffling via cross-mating as a
means for industrial strain improvement in S.
stipitis. The protocol is easy, inexpensive and
convenient. The key requirement is the availability of a screening method that is specific and sensitive. The improved haploid recombinant(s) are
genetically stable and amenable to be further
improved (i.e., they are not dead-end strains). For
example, the selected genome shuffled strains in
our study can be subjected to further genome
shuffling again to select for mutants with
improved tolerance to other stresses such as acetic acid and other pretreatment-derived inhibitors. Beneficial mutations in different mutant
lines can be easily recombined into one strain.
The same method can be applied to other yeasts,
including native pentose-fermenting yeasts such
as Pachysolen tannophilus, for which a genetic
mating system has been described [14].
Acknowledgments We thank Juraj Strmen and Frank
Giust of Tembec Inc. (Témiscaming, Québec, Canada) for
providing the HW SSL, and Tom Jeffries (USDA,
Madison, Wisconsin, USA) for providing the auxotrophic
S. stipitis strains.
References
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to improve thermotolerance, ethanol tolerance and
ethanol productivity of Saccharomyces cerevisiae.
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6. Hou L (2010) Improved production of ethanol by
novel genome shuffling in Saccharomyces cerevisiae.
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Li P, Jiang X-H (2010) Drug resistance marker-aided
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38:415–422
8. Bajwa PK, Pinel D, Martin VJJ, Trevors JT, Lee H
(2010) Strain improvement of the pentose-fermenting
yeast Pichia stipitis by genome shuffling. J Microbiol
Methods 81:179–186
9. Melake T, Passoth VV, Klinner U (1996) Characterization of the genetic system of the xylose fermenting
yeast Pichia stipitis. Curr Microbiol 33:237–242
10. Schneider H, Wang PY, Chan YK, Maleszka R (1981)
Conversion of D-xylose into ethanol by the yeast
Pachysolen tannophilus. Biotechnol Lett 2:89–92
11. Barbosa Mde F, Lee H, Collins-Thompson DL (1990)
Additive effects of alcohols, their acidic by-products,
and temperature on the yeast Pachysolen tannophilus.
Appl Environ Microbiol 56:545–550
12. Bajwa PK, Shireen T, D’Aoust F, Pinel D, Martin VJJ,
Trevors JT et al (2009) Mutants of the pentose-fermenting yeast Pichia stipitis with improved tolerance
to inhibitors in hardwood spent sulphite liquor.
Biotechnol Bioeng 104:892–900
13. Hashimoto S, Ogura M, Aritomi K, Hoshida H,
Nishizawa Y, Akada R (2005) Isolation of auxotrophic
mutants of diploid industrial yeast strains after UV
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J Gen Microbiol 129:1489–2494
Detection and Identification
of Fungal Microbial Volatile Organic
Compounds by HS-SPME-GC–MS
42
Bernhard Kluger, Susanne Zeilinger,
Gerlinde Wiesenberger, Denise Schöfbeck,
and Rainer Schuhmacher
Abstract
A method based on solid phase microextraction coupled to gas chromatography–mass spectrometry (GC–MS) for the detection and identification of
microbial volatile organic compounds (MVOCs) in the headspace of
filamentous fungi is presented. MVOCs are identified by comparison of
mass spectra and linear temperature programmed retention indices
(LTPRIs) with database entries and LTPRIs published in literature. The
presented method enables the monitoring of the formation of volatile
metabolites for defined time intervals during cultivation of the investigated fungus. The experimental procedure is exemplified with Fusarium
graminearum and Trichoderma atroviride but can also be used to detect,
identify and profile MVOCs produced by other filamentous fungi.
Keywords
Solid phase microextraction • Gas chromatography–mass spectrometry •
Microbial volatile organic compounds • Linear temperature programmed
retention index
Introduction
B. Kluger • D. Schöfbeck • R. Schuhmacher (*)
Department for Agrobiotechnology (IFA-Tulln),
University of Natural Resources and Life Sciences
Vienna, Konrad Lorenz Str. 20, Tulln 3430, Austria
e-mail: rainer.schuhmacher@boku.ac.at
S. Zeilinger
Research Area Gene Technology and Applied
Biochemistry, Institute for Chemical Engineering,
Vienna University of Technology, Getreidemarkt 9,
Vienna 1060, Austria
G. Wiesenberger
Institute of Applied Genetics and Cell Biology,
University of Natural Resources
and Life Sciences Vienna, Konrad Lorenz Str. 24,
Tulln 3430, Austria
Filamentous fungi produce and release microbial
volatile organic compounds (MVOCs), which are
frequently involved in self-signalling as well as
the interaction with other organisms, such as
plants and microbes. Because of their volatile
nature, these compounds can easily be transported
through air and therefore can act over relatively
long distances [1]. Complex mixtures of MVOCs
are being produced, individual metabolites of
which belong to different structure classes, such
as mono- and sesquiterpenes, alcohols, ketones,
lactones and the so-called C8-compounds [2–4].
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_42, © Springer Science+Business Media, LLC 2013
455
456
Nowadays, volatile components are usually
detected and identified by gas chromatography–
mass spectrometry (GC–MS) [5]. Gas chromatography (GC) is applied for the separation of
complex mixtures of (at least partly) evaporable
substances on capillary columns available with
stationary phases of different polarity. GC can
easily be coupled to mass spectrometry, which
offers a powerful tool for sensitive and selective
detection as well as the simultaneous identification
and structure characterisation of natural products
such as MVOCs.
Several sample preparation techniques have
been used for the extraction of MVOCs from fungal cultures. For solid [6] or liquid growth media
[7], solid phase extraction using C18 or silica gel
columns or steam distillation extraction [8] have
been described. While these methods require the
destruction of the fungal culture, headspace (HS)
techniques such as online gas enrichment on
adsorption tubes [9], closed-loop stripping analysis [10], dynamic headspace (purge and trap) [11]
or solid phase microextraction (SPME) [12–14]
can be used to sample volatile metabolites during
culture growth. Since these noninvasive HS methods allow the extraction, enrichment and detection of volatile metabolites from the living fungal
culture, they provide a more representative picture of the MVOCs actually produced [15]. In
addition, HS extraction helps to reduce the timeconsuming manual preparation of samples. This
holds particularly true for SPME because this
technique allows quick sample extraction and can
be fully automated and coupled online to GC–MS
analysis.
The HS-SPME technique [16] has already
been demonstrated to be very useful for the
extraction of volatile compounds from a wide
variety of samples, including MVOCs from plants
[15] and different fungal genera (e.g. Aspergillus,
Penicillium, Mucor, Fusarium, Trichoderma) [6,
12, 17].
Automated detection, identification and structure characterisation of MVOCs from GC–MS
chromatograms require efficient software tools
for comprehensive data evaluation, such as chromatographic peak detection, deconvolution (i.e.,
“purification”) of mass spectra and calculation of
B. Kluger et al.
retention indices. Suitable software programs are
available and can freely be downloaded from the
internet (e.g. AMDIS, MetaboliteDetector) [18,
19]. Assignment of chromatographic peaks and
putative identification of metabolites require the
comparison of measured mass spectra as well as
experimentally derived retention indices (i.e., linear temperature programmed retention index,
LTPRI) with reference spectra and values. For
both parameters, (pre-)defined criteria have to be
fulfilled for positive identification. In this protocol we present a comprehensive procedure that is
based on a method by Stoppacher et al. [20] and
has been further developed for the cultivation,
HS-SPME extraction, GC–MS analysis, automated detection and identification of MVOCs
produced by living cultures of filamentous fungi.
The procedure allows high flexibility, as growth
conditions can be controlled independently from
the sampling and extraction steps.
Materials
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
Potato dextrose agar (PDA).
Distilled water.
Autoclave.
50 °C water bath.
85-mm plates (Petri dishes).
20-mL headspace vials.
Cotton plugs and rubber straps.
Fungal culture, e.g. Trichoderma atroviride
(ATCC 74058), Fusarium graminearum
(PH-1, NRRL 31084).
Thermostat for incubator at 28 °C.
Cork borer for cutting 5-mm discs from agar
plate
Sterile syringe needles.
Fusarium minimal medium (FMM) plates
(1 g KH2PO4, 0.5 g MgSO4*7H2O, 0.5 g KCl,
2 g NaNO3, 30 g sucrose, 20 g agar, 10 mg
citric acid, 10 mg ZnSO4·6H2O, 2 mg
Fe(NH4)2(SO4)2·6H2O, 0.5 mg CuSO4*5
H2O, 0.1 mg MnSO4, 0.1 mg H3BO4, 0.1 mg
Na2MoO4·2H2O in 1 L deionised water).
Growth chamber 20 °C.
Mung beans substrate (MBS) medium (put
10 g of mung beans to 500 mL boiling water,
42
Detection and Identification of Fungal Microbial Volatile Organic Compounds…
cook for 20 min, filtrate the extract through a
paper filter, fill up to 1 L and autoclave for
20 min at 121 °C).
15. Sterile scalpels.
16. Orbital shaker.
17. Baffled flasks (250 mL).
18. Glass wool filter (remove the plunger of a
10-mL syringe and stuff it with glasswool to
the 5-mL mark).
19. 50-mL falcon tubes.
20. Centrifuge with swing-out rotor for 50-mL
falcon tubes (optional).
21. Light microscope and haemocytometer.
22. Laminar flow work bench; e.g. Heraeus
Instruments LaminAir® HB2472 (Heraeus,
Hanau, Germany).
23. Synthetic air (Messer, Gumpoldskirchen,
Austria).
24. Sterile filter, e.g. 0.2 mm PTFE Midisart BV
membrane filter (Sartorius, Göttingen,
Germany).
25. Screw caps containing 1.3-mm gas-tight silicone/teflon septa (Gerstel, Mülheim, Germany).
26. Climate chamber: 28 °C, 70 % humidity.
27. SPME headspace autosampler, e.g. Gerstel
MPS 2XL equipped with SPME fibre holder
(Gerstel, Mülheim, Germany; see Note 1).
28. SPME fibre, e.g. with 65 mm PDMS/DVB coating (Supelco, Bellefonte, USA; see Note 2).
29. Headspace inlet glass liner, 1.5 mm i.d. (e.g.
Supelco, Bellefonte, USA).
30. Gas chromatograph, e.g. Agilent 6890 N
(Agilent, Waldbronn, Germany).
31. Helium (5.0) (Messer, Gumpoldskirchen,
Austria).
32. GC capillary columns (see Note 3).
• Column 1: HP-5MS 30 m × 0.25 mm ×0.25 mm
(e.g. Agilent, Waldbronn, Germany).
• Column 2: DB-Wax 30 m × 0.25 mm × 0.25 mm
(e.g. Agilent, Waldbronn, Germany).
33. Mass Selective Detector, e.g. Agilent Mass
Selective
Detector
5975B
(Agilent,
Waldbronn, Germany).
34. Alkane standards (C8–C20), (C21–C40) (SigmaAldrich, Vienna, Austria) used for LTPRI
calibration.
457
35. Pentane 99 % (Sigma-Aldrich, Vienna,
Austria), hexane 99.5 % (Merck, Darmstadt,
Germany), heptane 99.5 % (J.T. Baker,
Deventer, Netherlands), octane 99 % (SigmaAldrich, Vienna, Austria), nonane 99 %
(Sigma-Aldrich, Vienna, Austria), decane
p.a. (Promochem, Wesen, Germany) also
used for LTPRI calibration
36. Software AMDIS—automated mass spectral
deconvolution and identification system
(version 2.69) (see Note 4)
Method
The method presented below describes a detailed
procedure for cultivation, sample preparation,
HS-SPME-GC–MS analysis, identification and
profiling of MVOCs produced by filamentous
fungi. The protocol will be exemplified for the
fungi Trichoderma atroviride and Fusarium
graminearum. Depending on the fungus to be
cultivated (using agar discs or alternatively spore
suspension) and studied, different parts of the
method may need modifications (e.g. cultivation
conditions, SPME fibre coating or extraction
temperature).
Preparation of PDA Plates
and Slants in Headspace Vials
In addition to biological samples, a HS vial containing only PDA medium (medium blank)
should be prepared and measured to exclude false
identification of compounds released from the
culture medium (see Note 5).
1. For preparing agar slants in headspace (HS)vials, sterilise the vials sealed with cotton
plugs and rubber straps.
2. Prepare 1 L of medium by suspending 39 g of
PDA in 1 L of distilled water.
3. Sterilise at 120 °C for 15 min and afterwards
cool the medium down to 50 °C in a waterbath.
4. For PDA plates, pour 25-mL aliquots of the
medium into sterile 85-mm Petri dishes.
458
5. Transfer 5-mL aliquots of the autoclaved PDA
medium equilibrated to 50 °C to the sterile HS
vials.
6. Place HS vials in slanted position until the
medium has solidified (see Note 6).
Cultivation of Fungi Using Agar Discs
(e.g. Trichoderma atroviride)
1. Prepare a fungal pre-culture (e.g. Trichoderma
atroviride) by cultivating the fungus on PDA
plates for 7 days in a 28 °C incubator in the
dark.
2. For cultivating the fungus in HS vials containing agar slants, cut a 5-mm disc from the preculture plate containing sporulated fungal
mycelium using a cork borer.
3. Transfer the disc from the plate to the HS vial
by placing it at a central position of the surface
of the PDA slant using a sterile syringe
needle.
4. Re-seal the HS vial with the sterile cotton plug
and the rubber strap.
Cultivation of Fungi Using Spores (e.g.
Fusarium graminearum)
As an alternative to cultivation using agar discs,
a spore suspension at a defined concentration can
be pipetted on top of agar surface and used for the
cultivation in HS vials.
1. Grow fungi (e.g. Fusarium graminearum) on
an FMM plate at 22 °C in the dark until the
mycelium reaches the edge of the Petri dish.
2. For sporulation of the required strain, scrape
five to ten small pieces of the mycelium (total
area about 2 cm2) with a sterile scalpel from
the surface of the plate (see Note 7) and suspend them in 50 mL of MBS medium in a 250mL baffled flask. Shake the flask on an orbital
shaker at 120 rpm at 20 °C until spores are
formed (2–4 days; see Note 8).
3. Separate the spores from the mycelium by
filtration of the culture through a column containing tightly packed glass wool, collect the
flow though in a 50-mL Falcon tube. If
B. Kluger et al.
desired, the spores can be concentrated by
gravity (overnight at 4 °C) or centrifugation
(5 min at 4,000 rpm) and removal of the
supernatant. Resuspend the spores in a small
amount of the supernatant or in sterile water
and store the suspension at 4 °C until further
use (see Note 9).
4. Determine the spore count using a
haemocytometer (e.g. Fuchs-Rosenthal).
5. Inoculate a HS vial with 8 × 103 spores by
pipetting the required volume to the centre of
the PDA surface.
6. Re-seal the HS vial with the sterile cotton plug
and the rubber strap.
Sample Preparation of Fungal Cultures
for Headspace Extraction
This method enables one to study the synthesis
rate of volatiles in defined time periods at different time points during fungal cultivation. This
step of the method is applied to purge culture
vials with air and thereby remove all volatiles
from the HS above the cultures (see Note 10).
1. Carry out the procedure at a defined time point
(e.g. 6 h) prior to GC–MS measurement in a
laminar flow work bench.
2. Remove the cotton plug from the HS vial.
3. Purge the headspace above culture with a gentle stream of air through a sterile filter for 30 s.
Caution: Spores might be expelled from the
HS vial to ambient air.
4. Tightly seal the HS vials with screw caps containing gas-tight silicone/teflon septa.
5. Incubate the cultures for a defined time period
at controlled conditions in the tray of the
SPME headspace autosampler (e.g. at 28 °C)
until measurement (see Note 11) (Fig. 42.1).
HS-SPME-GC–MS Analysis
of Filamentous Fungi
An SPME headspace autosampler is used to
extract volatiles automatically from the headspace of fungal cultures and transfer the extracted
MVOCs into the injector of the GC–MS system.
42
Detection and Identification of Fungal Microbial Volatile Organic Compounds…
459
Fig. 42.1 HS vials with
agar slants and culture agar
discs before purging with
synthetic air, sealed with
cotton plug and rubber
strap, (left) and after
purging, sealed with screw
cap (right)
Application of an autosampler increases precision and sample throughput.
1. Equilibrate the fungal cultures and blank sample at the extraction temperature of 30 °C for
5 min (see Note 12).
2. Insert the SPME fibre (e.g. 65 mm DVB/
PDMS coating) into the sample vial and
expose fibre coating to the HS above the cultivated fungus or blank sample (see Note 2).
3. Extract for 45 min at 30 °C without agitation
(fibre penetration depth 21 mm).
4. Transfer the fibre to GC–MS injection port
and desorb volatiles bound to the fibre for
2 min in a split/splitless injector (splitless
mode, 250 °C, fibre penetration depth of
57 mm, headspace inlet glass liner, 1.5 mm
i.d.).
5. After sample injection/desorption, clean the
fibre for 3 min in a needle heater station (gentle nitrogen stream, 250 °C, fibre penetration
depth of 57 mm).
6. Use helium (5.0) as carrier gas at a constant
flow rate of 1 mL/min.
7. Analyse the compounds desorbed from the
fibre on two columns for identification (see
Note 3):
• HP-5MS (30 m × 0.25 mm × 0.25 mm).
• DB-Wax (30 m × 0.25 mm × 0.25 mm).
8. Temperature oven program: 40 °C (hold
2 min), 10 °C/min to 200 °C, 25 °C/min to
260 °C (hold 5 min) (see Note 13).
9. Mass selective detector settings: electron ionisation (EI) at 70 eV, source 230 °C, quadrupole 150 °C, solvent delay of 4 min, full scan
(m/z 45–400) (Fig. 42.2).
GC–MS Measurements of Alkane
Standards for Calibration of LTPRI
Values
This step of the protocol describes the experimental details for the HS-SPME-GC–MS measurements of alkane standards (C5–C40). Alkane
standards have to be measured with the same
GC–MS method as the fungal samples. Their
retention times are required to determine LTPRI
values of fungal metabolites. LTPRI values were
calculated according to the formula suggested by
Van den Dool and Kratz in 1963 [21] (Fig. 42.3).
Compared to the procedure described in the section “HS-SPME-GC–MS Analysis of Filamentous
Fungi”, different extraction parameters are necessary for alkane standards (see Note 14). The
alkane standard mixture C8–C20 can be measured
with the protocol described in the section
“HS-SPME-GC–MS Analysis of Filamentous
Fungi”.
1. Alkane standard mixtures C8–C20 and C21–C40
are commercially available as solutions in
hexane with a concentration of 40 mg/L for
each alkane.
460
B. Kluger et al.
Fig. 42.2 Total ion chromatogram (displayed from 8 to 24 min) resulting from MVOCs of Trichoderma atroviride
after 48 h of cultivation using the HS-SPME-GC–MS method
Fig. 42.3 Overlay of two chromatograms (sample,
alkane standard) and formula for the calculation of the linear temperature programmed retention index (LTPRI).
The LTPRI represents the relative retention time of an
analyte, which is calculated based on the alkane standards,
one eluting before and the other eluting after the analyte
2. Prepare C5–C10 alkane standard mixture separately by using pure compounds (pentane:hex
ane:heptane:octane:nonane:decane
17:6:3:2:1:0.5 [v/v]).
3. Transfer each of the three alkane standard
mixtures to a separate empty clean HS vial
C5–C10: 1 mL, C8–C20: 10 mL, C21–C40: 40 mL
and immediately seal HS vials with screw caps
containing gas-tight silicone/teflon septa.
4. Equilibrate the HS vials including the alkane
standard mixture without agitation prior to
SPME extraction (C5–C10: 0.01 min, 90 °C;
C8–C20: 5 min, 30 °C; C21–C40: 30 min,
120 °C).
42
Detection and Identification of Fungal Microbial Volatile Organic Compounds…
5. Extract without agitation using a fibre penetration depth of 21 mm (C5–C10: 0.01 min, 90 °C;
C8–C20: 45 min, 30 °C; C21–C40: 60 min,
120 °C).
6. Follow the steps in the section HS-SPMEGC–MS Analysis of Filamentous Fungi, starting from step 4.
Data Evaluation
Putative identification of volatile metabolites is
based on two parameters: (1) LTPRI values and
(2) mass spectra. Consideration of both parameters requires the comparison with reference values. These reference LTPRI values or mass
spectra can either be taken from databases, the
scientific literature or from own measurements of
authentic standard substances (see Note 15).
Independent on where the reference LTPRI values and mass spectra originate from, a database
containing these reference values has to be created
before measured chromatographic peaks can be
assigned to certain metabolites. Moreover, specific
criteria have to be defined, which must be fulfilled
for putative identification of MVOCs, which can
be used for the profiling of filamentous fungi.
Establishment of a Reference Database
A reference database has to be established that
serves as a positive list for metabolite
identification. Therefore, mass spectra, LTPRI
values and a unique identifier (e.g. CAS number)
have to be selected for each compound contained
in the database. Comprehensive databases, such
as the Wiley Registry 9th/NIST 2008 mass spectral library, may serve for initial identification of
metabolites based on spectral similarity, because
only metabolites that are contained in the reference database can be identified by this approach.
For the determination of literature LTPRI values,
different sources, such as the NIST Chemistry
Webbook or the Adams library, are available (see
Note 15). Exact description of the data format
required for the reference database used by
AMDIS is beyond the scope of this protocol.
461
A detailed description of the AMDIS software is
given in the user manual, which can be accessed
via the Internet.
1. Use reference mass spectra obtained from
comprehensive databases (e.g. Wiley Registry
9th/NIST 2008, standard compounds).
2. Select appropriate reference spectra to be used
with the reference database.
3. Convert selected mass spectra to a format that
can be used by the program AMDIS to build a
reference database.
4. Add LTPRI values obtained from the literature
(e.g. NIST Chemistry Webbook) to reference
database.
Calibration of LTPRI Values for Alkane
Standards
In order to enable calibration of LTPRI values
corresponding to chromatographic peaks of
MVOCs (which follows in the steps of the section Automated Peak Detection, Deconvolution
of Mass Spectra and Calculation of LTPRI
Values), alkane standard chromatograms have to
be evaluated by AMDIS.
1. Open AMDIS program and load an alkane
standard data file.
2. Load settings for LTPRI calculation.
3. Use appropriate deconvolution settings (see
Note 16).
4. Use an alkane standard database containing
the spectra of C5–C40.
5. Analyse every alkane standard data file
(C5–C10, C8–C20 and C21–C40). Alkane standard peaks are detected automatically, mass
spectra are deconvoluted and retention times
of alkanes are used for the calibration of
LTPRI values according to the formula [21].
Automated Peak Detection,
Deconvolution of Mass Spectra
and Calculation of LTPRI Values
Metabolic profiling produces great quantities of
data, and manual data evaluation is difficult to
standardise and can be very labor-intensive and
462
time-consuming [22]. Therefore, the application
of tailor-made software programs (e.g. AMDIS,
MetaboliteDetector) is highly recommended (see
Note 4). The following protocol uses the program
AMDIS for LTPRI determination, component
detection and assignment of metabolites. The
program is freely available and accepts a wide
variety of manufacturers’ raw data formats [23].
1. Load sample data file with AMDIS (blank or
fungal culture chromatogram; see Note 5).
2. Load LTPRI calibration for the LTPRI calculation of detected peaks.
3. Use appropriate deconvolution settings (see
Note 16).
4. Exclude m/z values corresponding to siloxanes that are released by the column or the
fibre such as m/z 207 (see Note 17).
5. Analyse the opened data file for peak detection and deconvolution of mass spectra. LTPRI
of every detected peak is calculated based on
the alkane standard calibration made before.
Selection of Identification Criteria
and Assignment of Metabolites
Several criteria have to be fulfilled to assure the
identification of unknown MVOCs. Therefore,
obtained gas chromatograms and mass spectra
have to be compared to the reference database
established in Establishment of a Reference
Database. Moreover, when analysing fungal samples with the aim to identify MVOCs of this particular fungus for the first time, measurements
should be carried out on two columns with stationary phases of different polarity. The following protocol uses the comparison of mass spectra
and LTPRI values for the identification of MVOCs
using the deconvolution program AMDIS.
1. Compare the mass spectra of detected peaks to
the reference spectra contained in the reference database (e.g. Wiley Registry 9th/NIST
2008 mass spectral library) established in
Establishment of a Reference Database.
2. Open the menu “Analyze—Search NIST
library” in AMDIS.
B. Kluger et al.
3. Use following parameters for analysis:
• Hits reported per search: “Min. match factor ³90” (see Note 18).
• Select from “all components”.
• Number of components searched: “All
above threshold 0.0 % of total signal”.
• Libraries: Select the reference library to
compare a component spectrum.
4. Put all components that fulfil the criteria mentioned in step 3 on a positive list of tentatively
identified compounds.
5. Compare the experimentally defined LTPRI
values of tentatively identified compounds
with match factors ³90 with published LTPRIs
from literature with AMDIS (see Note 18).
6. Repeat the procedure for samples measured
on both, an HP-5MS and DB-Wax column.
For identification the compound has to fulfil
the criteria on both columns.
7. Confirm the mass spectrum and LTPRI of a
putatively identified substance with obtained
data of an appropriate pure standard compound (see Note 19).
8. Store mass spectra and LTPRI values of all
MVOCs identified according to the chosen
criteria in a separate database (see Note 20).
Notes
1. Manual extraction and injection of extracted
metabolites are also possible. For this purpose, a manual fibre holder is needed. It has
to be noted that the use of a manual fibre
holder is not only more labor-intensive, but
also leads to less reproducible measurements
results.
2. Choice of fibre coating material and fibre
thickness depends on the MVOCs produced
by the fungus and has to be optimised according to the analytical purpose. The selection
of adequate fibre coating material for the
extraction of metabolite classes is mainly
associated with the polarity of the metabolites to be extracted and is crucial to achieve
efficient extraction of MVOCs. Different
42
Detection and Identification of Fungal Microbial Volatile Organic Compounds…
fibre coatings, such as polyacrylate (PA),
polydimethylsiloxane (PDMS), PDMS/divinylbenzene (DVB), carboxene (CAR)/PDMS
and DVB/CAR/PDMS, are available.
3. Reliable identification of metabolites generally requires the application of two stationary phases of different polarity. In general,
retention behaviour of a selected analyte to
be identified is investigated, on both a nonpolar stationary phase (e.g. 100 % PDMS or
95 % dimethylpolysiloxane 5 % diphenylpolysiloxane), and a more polar stationary
phase (e.g. polyethylene glycol) with different chromatographic selectivity. For the columns mentioned in this protocol, LTPRI
values for various compounds are available
(e.g. NIST Chemistry Webbook).
4. The software program AMDIS (http://chemdata.nist.gov/mass-spc/amdis/downloads/)
[18] has been used for automated data
processing in this protocol. The software
program MetaboliteDetector (http://metabolitedetector.tu-bs.de/download.htmL) [19]
can also be used alternatively. Both programs
are freely available. They can be used for
automated LTPRI calibration and evaluation
of retention indices, automated deconvolution of raw mass spectra, peak detection and
calculation of mass spectral match factors.
Furthermore, both programs offer the option
of automated compound identification and
assignment of metabolites from a reference
list, if predefined identification criteria are
met. In contrast to the AMDIS program,
MetaboliteDetector additionally offers
batchwise processing and quantification of
identified as well as “unknown” metabolites.
5. Always include a blank sample that contains
only nutrition medium as a control to prevent
false positives originating from culture
medium, glassware, silicon septa, lab environment, for example. This sample should be
treated the same as the fungal culture sample
with regard to sample preparation, HS-SPMEGC–MS analysis and data evaluation.
Identified compounds in the blank sample
are not further considered for the database
including identified MVOCs.
463
6. Alternatively, only 2-mL aliquots of the autoclaved PDA medium can be transferred to
sterile HS vials and the vials placed in a horizontal position for medium solidification to
prevent a nutrition gradient.
7. For a good yield of spores it is important to
shred the mycelium into small pieces using a
scalpel before inoculating the MBS.
8. The sporulation should be carried out for at
least 2 days, but it should be stopped when
the first germinating conida are visible.
9. It is preferable to use freshly made spore suspensions because the age of the spores can
have a significant influence on germination
onset and growth and, consequently, the
HS-profile.
10. The purging step is recommended for two
reasons. First, removal of all metabolites
accumulated above the culture enables the
monitoring of MVOCs synthesis starting
from that particular time point. Second,
replacement of cotton plug by tightly sealing
screw caps usually leads to uncontrollable
loss of metabolites from the culture vial and
reduces the precision of the method.
11. Incubation temperature has to be selected as
required for the fungal culture. Standardisation
of incubation temperature is recommended
in order to reduce biological variability.
12. Extraction temperature was set to 30 °C in
order to minimise affection of the biological
culture while simultaneously achieving standardised conditions by heating to a temperature above room temperature. If required,
extraction can also be carried out without
temperature control or under cooled conditions. However, it has to be considered that
extraction yield will decrease at lower
temperature.
13. Conditions have been chosen with the aim to
achieve appropriate chromatographic separation of MVOCs. Improved chromatographic separation especially for terpenoids
can be obtained by lowering the rate of temperature increase/time interval.
14. For calculation of correct LTPRI values, narrow and symmetric chromatographic peaks
of Gaussian shape are required for each
464
alkane standard. Therefore, different SPME
extraction methods have to be used for alkane
standard mixtures C5–C10 and C21–C40.
Alkane standard mixtures injected at too high
concentrations or after extraction with an
inadequate SPME program will result in poor
peak shapes and consequently lead to inaccurate LTPRI values. Hence, it is highly recommended to optimise the analysis of
n-alkanes before the measurement of fungal
cultures. It might happen that not all n-alkanes
elute from the column until the end of a
single chromatographic run. In particular,
high-boiling alkanes might need higher temperatures in order to elute during the chromatographic run. Therefore, it is recommended
to bake out the column at a temperature close
to maximal tolerable temperature after measuring alkane standard mixtures.
15. Usually more than one LTPRI value can be
found for a particular stationary GC-phase in
the literature. However, the choice of the reference LTPRI should not be made arbitrarily.
NIST chemistry Webbook (http://webbook.
nist.gov/chemistry/) [24], for example, contains a collection of various LTPRI values
published in the literature for every compound. LTPRI values from the literature
should be as consistent as possible with measured LTPRI values regarding column coating, column dimensions and the gas
chromatographic method to assure best compatibility. We recommend the use of the
median of published LTPRI values.
16. Deconvolution settings have to be adjusted
according to the peak intensity, width and
shape in chromatograms. The following settings shall serve as an example and have
been found suitable in our laboratory for the
evaluation of raw chromatograms obtained
with instrumentation used in the present
study. These settings might serve as orientation and will have to be modified according
to the samples to be evaluated:
• Component width: “32”
• Adjacent peak subtraction: “One”
• Resolution: “Medium”
B. Kluger et al.
17.
18.
19.
20.
• Sensitivity: “Medium”
• Shape requirements: “Medium”
• User manual and detailed program description can be downloaded from the NIST
Webpage.
The following m/z values are well known to
be caused by decomposition of polysiloxane
material used as GC stationary phase or fibre
coating and therefore should be excluded
from further data evaluation: m/z 77, 207,
267, 281, 341, 355. These signals can also be
recognised from typical isotopic distribution
of silicium.
Identification criteria for compounds may
vary according to the analytical study. For
metabolic profiling of MVOCs the application of strict criteria is advised due to the fact
that only few standards are commercially
available for tentatively identified compounds. Strict criteria help to exclude false
positives. Therefore, it is recommended to
confirm the results for identification of a
compound on two columns of different stationary phase. We set the tolerance for the
maximum relative deviation to ±2 % of literature values and a minimum match factor
for the similarity of mass spectra of ³90 %.
Additionally, the compound should be
confirmed by a standard, if available.
Many MVOCs identified are not available as
standard compounds; therefore, it is not possible to confirm every MVOC. Dilute available standard compounds in a concentration
resulting in narrow and symmetric peak
shapes. No general concentration for standards can be recommended due to the fact
that every substance has a different binding
behaviour on particular fibre coatings.
Standards can be diluted in n-hexane or
ACN:H2O (1:1) depending on solubility.
This database can be used for the further
metabolic profiling of, for example, this particular fungal strain, species or genus.
Acknowledgement The financial support by the
Austrian Science Fund (FWF projects F3702 and F3706)
is gratefully acknowledged.
42
Detection and Identification of Fungal Microbial Volatile Organic Compounds…
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Transformation Methods
for Slow-Growing Fungi
43
Suman Mukherjee and Rebecca Creamer
Abstract
It is challenging to prepare protoplasts and establish a transformation system in any toxin that produces slow-growing fungus, like Undifilum oxytropis. This chapter describes a protocol to generate a nontransient
transformation system with the help of introducing a foreign heterologous
gene to a toxin producing slow-growing fungi. This system will be useful
to study genes in metabolically important pathways.
Keywords
Slow-growing fungi • Protoplast • Transformation • Undifilum oxytropis •
Green fluorescent protein
Introduction
The preparation and regeneration of protoplasts
and the subsequent development of transformation systems are well established for several
fungi. Several groups, such as Fierro et al. and
Akamatsu et al., established transformation systems for ascomycete fungi, whereas others, such
S. Mukherjee (*)
National Institutes of Health, Laboratory of Biochemistry
and Genetics, NIDDK, 8 Center Drive, Room 326,
Bethesda, MD 20892, USA
e-mail: suman.mukherjee@nih.gov; creamer@nmsu.edu
R. Creamer
Department of Entomology, Plant Pathology,
and Weed Science, New Mexico State University,
Box 30003, MSC 3BE, 945 College Ave, Las Cruces,
NM 88003, USA
as Panaccione et al., developed transformation
systems for the endophyte Neotyphodium [1–3].
All of these systems were developed for fungi
that rapidly grow and reproduce.
Toxic locoweeds are found to be associated
with a slow-growing endophytic fungus, Undifilum
oxytropis. U. oxytropis is an Embellisia-like fungus that does not produce external mycelia on
host [4]. When cultured in vitro, U. oxytropis
grows up to 0.03–0.34 mm/day on potato dextrose
agar (PDA) plates at room temperature [5]. U.
oxytropis produces a polyhydroxy alkaloid,
swainsonine (1, 2, 8-trihydroxyindolizidine), naturally and in culture [5, 6], which has been correlated with toxicity of locoweeds [5, 7]. This
protocol details the generation of protoplasts and
establishment of a stable transformation system
from slow-growing toxin-producing fungi [8].
The current protocol was established for
U. oxytropis but can be extended to several
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_43, © Springer Science+Business Media, LLC 2013
467
468
slow-growing toxin-producing fungi, including
endophytes. Protoplasts of U. oxytropis were prepared and transformed with a fungal-specific
vector that expresses green fluorescent protein
(GFP). Regenerated fungal mycelia were screened
for GFP expression. In this chapter we describe
protoplast preparation, regeneration with selection, nontransient GFP expression, and a stable
transformation system using a slow-growing fungus. This technology can provide tools to understand metabolically important pathways of
toxin-producing fungi.
S. Mukherjee and R. Creamer
31.
32.
33.
34.
35.
36.
37.
38.
39.
Phenylmethylsulphonyl floride
EDTA
Triton-X 100
Glycerol
SDS-polyacrylamide gel
Immobilon polyvinylidene fluoride (PVDF)
Tween-20
Tris buffered saline
Rocking platform
Methods
Fungal Strains and Culture Conditions
Materials
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
25.
26.
27.
28.
Potato dextrose agar plates (PDA)
Water agar media
Fungal-specific vector
Potato dextrose broth
Antibiotic hygromycin B (Hyg)
Platform shaker at 200 rpm
Miracloth
Buchner funnel
b-glucuronidase
Lysing enzyme
b-d-Glucanase G
Bovine serum albumin
MgSO4
NaH2PO4
Sorbitol
Tris–HCl
Refrigerated centrifuge
CaCl2
Hemocytometer
4’, 6-Diamidino-2-phenylindole
Deionized water
Fluorescence microscope
TE buffer
Sucrose
Yeast extract
Casein hydrolysate
Bacto Agar
Poly ethylene glycol, 4,000 molecular
weight
29. NaCl
30. NaF
1. Fungal isolates can be isolated from plants or
harvested from colonies grown on potato dextrose agar plates (PDA).
2. If collecting from plant tissues, surface sterilize for 30 s in 70 % ethanol, followed by 3 min
in 20 % bleach, and then 30 s in sterile water.
3. Dry the plant tissue and place on water agar
media; incubate media plates at room temperature (25 °C) for 2 weeks.
4. Transfer fungal hyphae from water agar media
to PDA plates and grow at room temperature
for at least 14 days [9].
5. Preserve fungal isolates and store at both 4 °C
and −80 °C. Fungal plates can also be maintained in a sealed plastic box at room temperature for further experiments (Fig. 43.1).
Hygromycin B Selection
1. pPd-EGFP vector was used for U. oxytropis
transformation; an ascomycete-specific vector
can be used for transformation.
2. pPd-EGFP is a fungal-specific expression vector that encodes the antibiotic hygromycin B
(Hyg) resistance gene driven by the Aspergillus
nidulans trpC-promoter [10]. The vector also
expresses GFP driven by the Cryphonectria
parasitica glyceraldehydes-3-phosphate dehydrogenase (gpd) promoter.
3. Generate working stock of Hyg at a concentration of 1 mg/mL (dissolve in sterile water).
43
Transformation Methods for Slow-Growing Fungi
469
Fig. 43.1 Unidifilum
oxytropis (25-1 isolate)
grown on PDA plates
for 4 weeks
4. Add Hyg to a final concentration of 20 mg/mL
for a 20 mL of PDA plate. For U. oxytropis
20 mg/mL of Hyg is the lethal dose, so any
growth of mycelia on plate represents growth
of positive transformants.
5. Lethal dose differs among fungal species, so a
kill curve must be determined with the required
antibiotic using serial dilutions of the antibiotic on PDA plates.
6. Measure fungal growth on the PDA plate in
mm over a 14-day span or more depending on
the growth rate of the respective fungus. Use
the data to generate a kill curve for hygromycine sensitivity of the fungi.
Preparation of Protoplasts,
Transformation, and Regeneration
This method was used to prepare fungal
protoplasts.
1. Add 1 g of fungal mycelia into potato dextrose broth in a 250-mL conical flask.
2. Incubate with shaking for 2 weeks on a
platform shaker at 200 rpm at room
temperature.
3. Filter the fungi and media to collect the fungal mass using miracloth in a Buchner
funnel.
4. Re-suspend mycelial mass in 100 mL of
digestion buffer.
5. Digestion buffer composition: 1 mL of
b-glucuronidase, 75 mg lysing enzyme,
800 mg b-d-Glucanase G, 600 mg bovine
serum albumin dissolved in 100 mL osmotic
medium (1.2 M MgSO4 with NaH2PO4,
pH 5.8).
6. Incubate at room temperature on an orbital
shaker for 3 h.
7. The 3 h of incubation time after addition of
digestion buffer provides optimum protoplasts yield for U. oxytropis. Optimum incubation time depends on fungal species and
isolates.
8. After incubation, collect 8 mL of digested
mycelial mass in 30-mL glass centrifuge
tubes.
9. Combine mycelial mass with 10 mL of
trapping buffer (0.4 M Sorbitol in 100 mM
Tris–HCl, pH 7.0).
10. Centrifuge at 6,000 rpm at 4 °C for 15 min.
11. Collect protoplasts at the interface of the two
layers.
12. Add 2 volumes of 1 M sorbitol to the collected protoplasts.
13. Centrifuge the protoplasts at 6,000 rpm at
4 °C for 5 min.
14. Collect pellets after decanting the supernatant.
470
15. Suspend the collected protoplasts (pellets)
in 100 mL of STC buffer (STC buffer: 1 M
sorbitol in 100 mM Tris–HCl, pH 8.0,
100 mM CaCl2).
16. Count aliquots of protoplasts using a hemocytometer with a light microscope.
17. Keep suspended protoplasts on ice for transformation experiments (109 protoplasts per
mL can be obtained using this method).
DAPI Staining
To check for quality/viability of protoplasts,
DAPI (4¢, 6-Diamidino-2- phenylindole) stain
can be used.
1. DAPI stock solution is 5 mg/mL (10 mg in
2 mL of deionized water).
2. Add DAPI stock solution into the protoplasts
to make a 100 ng/mL final concentration.
3. Incubate 10 mL of the protoplast /DAPI solution in a sterile microcentrifuge tube for 30 min
at room temperature and observe in fluorescence
microscope with excitation at 360 nm [8].
4. DAPI stock solution should be stored in a
brown bottle, protected from light at −20 °C;
working solution can be stored at 4 °C.
Transformation Procedure
1. Dilute 5 mg of vector in 10 mL of TE (0.5×,
pH 8) buffer.
2. Add the vector solution (10 mL) to purified
protoplasts.
3. Incubate on ice for 1 h.
4. GFP expression can be observed at this point
using a fluorescence microscope [8].
5. Add 1 mL of PTC buffer (PTC buffer: 40 %
poly ethylene glycol, 4000 molecular weight,
100 mM Tris–HCl, pH 8.0 and 100 mM
CaCl2) to each tube of protoplast.
6. Incubate 25 min at room temperature.
7. Plate the transformation mixture as 2, 20,
and 200 mL droplets onto Petri dishes.
8. Add 12.5 mL regeneration medium (1 M of
sucrose, 0.001 w/v of yeast extract, 0.001 w/v
of casein hydrolysate and 0.016 % of Bacto
Agar) to each plate.
S. Mukherjee and R. Creamer
9. Add further 12.5 mL regeneration media
containing 40 mg/mL Hyg overlaid on each
plate (double the amount of Hyg to be added
to avoid dilution of the antibiotic).
10. Incubate plates for 4 days at room temperature (see Note 1).
Sporulation and Hyphal Tipping
1. Grow transformant on Hyg-containing water
agar plates (complete separation of clonal
hyphae can be observed in water agar plate).
2. Collect single hyphal tips under microscope to
start a mononucleate culture on PDA plates
again.
3. Observe GFP expression after 2 weeks of
growth on PDA plates (see Note 2).
Microscopy
1. Take fungal mycelia on a slide, add 10 ml of
water, and observe under fluorescence
microscope.
2. Use fluorescence microscope to capture
fluorescence images at a combined objective
and eyepiece magnification of 40× and 10×,
respectively. Mycelium demonstrated strong
expression of the GFP gene, as observed by
emission at 488 nm using an Axiovert 200 M
microscope (any other fluorescence microscope can be used).
3. GFP expression of emerged mycelia was
observed microscopically after 3 weeks of
growth (Fig. 43.2). Expression was retained
after several weeks and passages of mycelial
growth on PDA plates as observed (Fig. 43.3)
(see Note 3).
Detection of Transformed Gene
or Marker by Immunoblot
1. Harvest fungal tissues from transformed and
untransformed U. oxytropis for western blot
experiment.
2. Homogenize filtered fungal tissue with liquid nitrogen in extraction buffer (Extraction
43
Transformation Methods for Slow-Growing Fungi
471
Fig. 43.2 (a) Mycelial mass of U. oxytropis 3 weeks
post-transformation on hygromycin selection PDA plate.
(b) Fluorescence image of mycelial growth of U. oxytro-
pis 3 weeks post-transformation on hygromycin selection
PDA plate expressing GFP. (c) Overlay of (a) and (b)
buffer: 50 mM Tris–HCl, pH 7.5, 100 mM
NaCl, 50 mM NaF, 2 mM phenylmethylsulphonyl floride, 5 mM EDTA, 1 % Triton-X
100, 10 % glycerol).
Centrifuge at 4 °C for 10 min at 13,000 rpm.
Quantify protein concentration of lysates
using a protein assay kit.
Load total protein (20 mg) on a 12 % SDSpolyacrylamide gel.
Transfer the gel to Immobilon polyvinylidene
fluoride (PVDF) transfer membrane.
Block the membrane in 3 % (w/v) bovine
serum albumin and dissolve in 1× Tris buffer
saline with Tween-20 (TBST) for 1 h at room
temperature (on rocking platform).
Dilute GFP or selective marker antibody as
recommended by manufacturer.
Add primary antibody solution to the membrane and incubate for 1 h at room temperature (on rocking platform).
10. Wash the membrane by rocking in 10 mL of
1× TBST three times, 10 min each.
11. Add secondary antibody solution to the
membrane and incubate for 1 h at room temperature (on rocking platform).
12. Wash the membrane by rocking in 10 mL of
1× TBST three times, 10 min each.
13. Visualize blots using the detection kit
(Fig. 43.4) (see Note 4).
3.
4.
5.
6.
7.
8.
9.
Notes
1. Although serial dilutions were performed on
PDA plates, observation of individual colonies is difficult after 7 days of regenerated
growth because the transformants can merge
with each other. A count of individual colonies
should be done at an early stage to measure
transformation efficiency. Fourteen days after
472
Fig. 43.3 (a) Mycelial mass of U. oxytropis 9 weeks
post-transformation on hygromycin selection PDA plate.
(b) Fluorescence image of mycelial growth of U. oxytro-
Fig. 43.4 Immunoblot of hyphal samples of U. oxytropis
using anti-GFP antibody. 27 kDa band indicates GFP
expression. Lane 1, non-transformed hyphae; lane 2,
hyphae 3 weeks post-transformation; lane 3, hyphae 9
weeks post-transformation
transformation, complete mycelia formation
was observed on Hyg-containing PDA plates.
No hyphae were observed in untransformed
protoplasts plated on Hyg-containing PDA
plates [8].
S. Mukherjee and R. Creamer
pis 9 weeks post-transformation on hygromycin selection
PDA plate expressing GFP. (c) Overlay of (a) and (b)
Slow-growing fungi such as U. oxytropis
may take 7 days to grow mycelia after regeneration. In other ascomycetes, mycelial growth
can be observed within 24 or 48 h.
Perform a control transformation mixture
without vector (containing antibiotic marker)
following similar protocol as above. Plate on
Hyg-containing PDA plate as control.
2. Sporulation is a rare event for U. oxytropis,
and conidial germination rates are poor.
Hyphal tipping was used to start new cultures.
For other fungi, single spores can be used to
start new cultures on PDA plates.
3. GFP expression was observed for several
weeks and months in transformed fungal
mycelia, which indicates the establishment of
a stable transformation system. We used GFP
43
Transformation Methods for Slow-Growing Fungi
marker for fluorescence microscope study, but
any marker can be used.
4. We used GFP as a marker to investigate transformation, but any marker can be detected
with this method if suitable antibody is present.
Blocking buffer can be made also with PBST
with 2 % dry non-fat milk. Primary and secondary antibody solution was generated by
diluting designated antibodies in blocking
solution.
Acknowledgements The authors would like to thank Dr.
Richard Richins, Dr. Soum Sanogo, Dr. Swati Mukherjee,
and Deana Beacom of New Mexico State University for
technical assistance and constructive discussion.
References
1. Akamatsu H, Itoh Y, Otani H, Kohmoto K (1997)
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2. Fierro F, Gutierrez S, Diez B, Martin JF (1993)
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notatum and chromosome I (10.4 Mb) in Penicillium
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J, Hambleton S (2009) Undifilum, a new genus for
endophytic Embellisia oxytropis and parasitic
Helminthosporium bornmuelleri on legumes. Botany
87:178–194
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Harris CM, Schneider MJ, Ungemach FS, Hill JE,
Harris TM (1988) Biosynthesis of the toxic indolizidine alkaloids slaframine and swainsonine in
Rhizoctonia leguminicola: metabolism of 1-hydroxyindolizidines. J Am Chem Soc 110:940–949
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Thacker E (2002) Predicting toxicity of tall larkspur
(Delphinium barbeyi): measurement of the variation
in alkaloid concentration among plants and among
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Development of a transformation system in the
swainsonine producing, slow growing endophytic
fungus, Undifilum oxytropis. J Microbiol Methods 81:
160–165
Ralphs MH, Creamer R, Baucom D, Gardner DR,
Welsh SL, Graham JD et al (2008) Relationship
between the endophyte Embellisia spp. and the toxic
alkaloid swainsonine in major locoweed species
(Astragalus and Oxytropis). J Chem Ecol 34:32–38
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expression vectors. J Virol 74:7568–7577
Enzymatic Saccharification
of Lignocellulosic Biomass
44
Manimaran Ayyachamy, Vijai Kumar Gupta,
Finola E. Cliffe, and Maria G. Tuohy
Abstract
The conversion of polymers present in the lignocellulosic biomass into
fermentable sugars can be achieved through physical/chemical and enzymatic pretreatments. The microbial conversion of biomass to bioenergy
will be cost-effective only if all of the components in the biomass are converted into value-added products. The combination of appropriate chemical and enzymatic conversion methods is very important to develop an
effective biomass to biofuels and biorefineries conversion technology.
Keywords
Lignocellulosic biomass • Pretreatment • Saccharification • Enzyme
cocktails • Biofuels • Biorefineries
Introduction
M. Ayyachamy (*) • F.E. Cliffe • M.G. Tuohy
Molecular Glycobiotechnology Research Group,
Department of Biochemistry, School of Natural Sciences,
National University of Ireland,
University Road, Galway, Ireland
e-mail: manimaran.ayyachamy@nuigalway.ie
V.K. Gupta
Molecular Glycobiotechnology Research Group,
Department of Biochemistry, School of Natural Sciences,
National University of Ireland, University Road,
Galway, Ireland
Assistant Professor of Biotechnology, Department
of Science, Faculty of Arts, Science & Commerce,
MITS University, Rajasthan, India
Rapidly depleting stocks and increasing demand
for petroleum-derived energy sources, in combination with exponentially increasing global
warming, have forced the public and private sectors globally to look for alternative technologies
to meet future energy needs. Biofuels are promising as future candidate fuels as they are largely
produced from renewable plant resources or
wastes. Biofuels have great potential not only as
environmentally clean, renewable energy sources,
but also they provide a mechanism to reduce reliance on imported energy sources [1, 2].
Plant biomass (or lignocellulosic biomass) is
approximately composed of cellulose (20–50%)
and hemicellulose (15–35%) and strongly
intermeshed with the aromatic copolymer lignin
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_44, © Springer Science+Business Media, LLC 2013
475
476
[3–8]. The conversion of polysaccharides present
in the biomass into fermentable sugars can be
achieved through physical/chemical and enzymatic methods [6, 9]. The microbial conversion
of biomass to energy will be cost-effective only
if all of the components in the biomass are
converted into value-added products, which is
the basis of the lignocellulosic biorefinery concept [10–13].
A primary target of a biomass to biorefinery
strategy is to maximize the conversion of the
significant reservoir of complex carbohydrates to
feedstocks that are suitable for the production of
biofuels and other commodity products through
fermentation or thermochemical processing [12,
14]. To maximize recovery of fermentable sugars, lignocellulosic biomass is generally pretreated with chemicals and/or microbial enzymes
to reduce the size of the feedstock and also to
open up the plant structure [15, 16]. Cellulose
and hemicellulose, which are the principal sugar
components, are bound with lignin in a complex,
highly ramified and cross-linked matrix that is
designed to resist chemical and microbial attack.
This structure limits bioconversion, either by
microorganisms or the enzymes they produce;
therefore, a pretreatment is generally required
before lignocellulosic biomass can be subjected
to microbial or enzymatic conversion.
Pretreatment technologies for lignocellulosic
biomass can be assigned into four broad categories: physical, chemical, physicochemical, and
biological methods. An ideal pretreatment should
remove lignin and thus reduce the crystallinity of
cellulose [17], increase the porosity and accessibility of the cellulose (and hemicellulose) to
enzymatic hydrolysis, generate low levels of
compounds inhibitory to both enzymes and fermentation microorganisms, be of low cost and
have low energy requirements. Overall, the pretreatment should result in a reduction in the
recalcitrance of lignocellulose and increase
accessibility to enzymes. Pretreatments vary
from hot-water extraction, steam pretreatments
(often with an oxidant or other chemical), to
weak and strong acid and alkali pretreatments
[18, 19]. However, pretreatments may affect the
composition and interactions between biomole-
M. Ayyachamy et al.
cules in the substrate in a way that is not necessarily advantageous to downstream bioconversion
[20, 21].
The ideal pretreatment should minimize the
need for feedstock particle size reduction, limit
the formation of sugar degradation compounds,
minimizes energy demands, and reduces the
overall costs. During the chemical pretreatment
process, sugar-derived compounds such as
furfural and 5-hydroxymethyl furfural are
formed as co-products [22]. These compounds
are very toxic to microorganisms and also inhibit
subsequent enzymatic saccharification process.
Therefore, these compounds must be removed or
neutralized prior to the microbial fermentation
and enzymatic hydrolysis. Biological pretreatments are based mainly on the use of white rot
fungi to delignify biomass in a low-cost treatment approach (some loss of hemicellulose and
minor amounts of cellulose occurs as a result of
this type of microbial pretreatment). Microbial
delignification, although gentle and effective, can
remove upto 32% of the lignin from biomass
materials, such as corn stover [23]. However, this
pretreatment method generally does not give high
sugar yields during subsequent hydrolysis.
Furthermore, microbial pretreatment times are
lengthy, typically requiring 18–35 days.
Enzymatic saccharification of lignocellulosic
biomass is a very complex process and the hydrolysis of all of the polysaccharides requires a repertoire of several hydrolytic enzymes. In the
biosphere, a variety of pathogenic and saprophytic microorganisms play an unparalleled role
in bringing about the depolymerization and
decomposition of plants and plant-derived residues and wastes. Fungi are key microbial players
in the biological conversion of plants and plantderived wastes. These eukaryotic microorganisms secrete a vast array of carbohydrate
hydrolases (collectively termed glycosyl hydrolases) as well as a range of peptidases and ligninmodifying enzymes that reduce plant biomass to
its simple building blocks. For decades, scientists
have worked to identify the types and modes of
action of cellulases and hemicellulases produced
by fungi. Species of ascomycete fungi, namely
Trichoderma reesei and Aspergillus niger, have
44
Enzymatic Saccharification of Lignocellulosic Biomass
been exploited as cell factories for production of
commercial cellulase and hemicellulase enzyme
products. More recently, additional fungal species have been investigated as sources of cellulases and hemicellulases with enhanced catalytic
efficiency and thermostability [24, 25].
Thus, the combination of chemical and enzymatic methods is required for an efficient conversion of biomass into fermentable sugars, and
selection of appropriate combinations of chemical and enzymatic conversion methods is very
important to develop an effective biomass to biofuels and biorefinery conversion technology.
Materials
1. Feedstocks: lignocellulosic biomass collected from local sources (see Note 1).
2. Enzymes: cellulolytic and hemicellulolytic
enzymes were obtained from Novozymes
(Denmark) and Genencor (USA).
3. Novozymes: Celluclast, Cellic C Tech 2,
Cellic H Tech 2.
4. Genencor: Accellerase 1500, Spezyme CP,
Accellerase CY, Accellerase XC, Accellerase
XY, Accellerase BG.
5. Buffers (100 mM): pH (5–6)—Sodium acetate buffer; pH (7–8)—Sodium phosphate
buffer (pH 7-8); pH 9—Glycine-NaOH buffer (see Note 2).
6. Substrates: wheat arabino xylan (Megazyme
International Ireland Ltd., Bray, Co.
Wicklow), oat spelts xylan (SigmaAldrich, Dublin, Ireland), birch wood xylan
(Sigma-Aldrich), b-glucan (from barley;
Megazyme Intl. Ireland Ltd.), and carboxy
methyl cellulose and 4-nitrophenyl a- or
b-glycosides (Sigma-Aldrich) (see Note 3).
7. Dionex high-performance anion exchange
chromatography (HPAEC) ICS-3000 ion
chromatography system equipped with an
autosampler, a UV detector, and ED40
pulsed amperometry detector, fitted with a
gold electrode (Dionex Corporation,
Sunnyvale, CA, USA). SD-10 and PA-100
analytical and guard columns (Dionex
477
8.
9.
10.
11.
12.
13.
14.
15.
16.
Corporation, Sunnyvale, CA, USA)
Chromeleon™ Version 6.70 software for
data collection, processing, and analysis
(Dionex Corp., Sunnyvale, CA, USA) (see
Note 4).
Chemicals: Unless otherwise stated, all
chemicals and reagents were purchased from
Sigma-Aldrich (Dublin, Ireland).
Fine-grade muslin was purchased from a
local supplier.
Grant water baths (Mason Technology,
Ireland).
Thermostatically controlled New Brunswick
Scientific Innova 44 platform, shaking incubator (New Brunswick Scientific, USA).
High-temperature oven (Fisher Scientific,
Ireland).
BioTek Powerwave XS2 microplate reader
with incubation facilities to 50 °C (BioTek,
USA).
PDFE membrane (0.2 mm) syringe-less filters
(Millipore, USA).
Laboratory plastics, i.e., disposable micropipette tips, microcentrifuge tubes, and flatbottomed, high-optical-quality, low-proteinabsorbing microtitre plates (Sarstedt,
Sinnottstown, Co. Wexford, Ireland).
Safety apparel (Caulfields Ltd., Galway,
Ireland).
Methods
Feedstock Collection
and Characterization
1. Lignocellulosic biomass is collected from
local areas and is washed thoroughly with tap
water to remove contaminants.
2. Feedstock must be dried at 50 °C in a hot air
oven for 3 days.
3. Feedstock is milled to a particle size of
2–3 mm before subjecting to either chemical
or enzymatic pretreatments (see Note 5).
4. Total solids, ash, nitrogen, and carbohydrate
contents in the biomass sample are analyzed
using the standard methods [22, 26].
478
Commercial Enzymes
1. Cellulolytic and hemicellulolytic enzymes can
be purchased from the leading commercial
suppliers such as Novozymes (Denmark) and
Genencor (USA).
2. Necessary enzyme activities have to be determined [27–32] using the standardized assay
methods (see Note 6).
3. The enzyme solution should be prepared
freshly every time from the stock prior to
saccharification (see Note 7).
4. Enzymes should be kept at 4 °C and brought
to room temperature prior to use.
Enzymatic Saccharification
of Lignocellulosic Biomass
In order to achieve maximum biomass conversion, the reaction conditions for enzymatic
hydrolysis should be optimum. The following
experiments must be performed to determine the
optimal reaction conditions for enzymatic
saccharification.
pH Optimization
1. Prepare buffers at different pH values over the
range 2.6–11. Sterilize the buffer solutions in
an autoclave at 121 °C for 15 min. Allow the
buffer solutions to cool to ambient temperature prior to use (see Note 8).
2. Weigh out 1.5 g of biomass and add 15 mL of
buffer solution to obtain a 10% (w/v) solids
ratio. Prepare all tests in triplicate.
3. Uniformity in the enzyme dosage has to be
maintained since different types of commercial enzymes are used for saccharification.
Therefore, a consistent enzyme dosage should
be used for each test, when comparing different enzymes.
4. Controls in (triplicate) consisting of a denatured sample of the appropriate enzyme should
be run in parallel with tests (see Note 9).
5. Sodium azide is added at 0.02% to prevent
the microbial contamination. If using a
thermostable enzyme preparation at higher
M. Ayyachamy et al.
reaction temperature, preservative may not be
required.
6. Reactions are carried out at 50 °C for 72 h
under shaking conditions (150 rpm).
7. Sampling must be done every 1 h at the start of
the reaction, and every 3–12 h thereafter.
Samples should be centrifuged to remove residual biomass and then assayed for the release of
reducing sugars [31] from the polysaccharides
present in the biomass (see Note 10).
Optimization of Temperature
1. 1.5-g quantity of biomass is weighed and
transferred to a 50-mL polypropylene centrifuge tube. A 15-mL volume of buffer solution
(optimized pH) is added. All tests are prepared
in triplicate.
2. The enzyme should be maintained at an identical dosage level in all tests comparing the
commercial enzymes.
3. A set of controls (in triplicate) with denatured
enzyme must be included in parallel with
test.
4. Sodium azide is added at 0.02% to prevent the
contamination.
5. The reaction is carried out at different temperatures (30–80 °C) for 72 h at 150 rpm.
6. Withdraw samples every 1 h at the start of the
reaction, and every 3–12 h thereafter. Samples
should be centrifuged to remove residual biomass and then assayed for the release of reducing sugars [31] (see Note 10).
Optimization of Enzyme Dosage
1. Add 1.5 g of biomass and transfer to 50-mL
PP centrifuge tube. Add 15 mL of buffer (at
the optimum pH). All tests are prepared in
triplicate.
2. The enzyme should be added at different dosage levels, such as 1, 5, 10, 25, or 50 IU per
gram of biomass (see Note 11).
3. Triplicate controls are prepared with denatured enzyme.
4. Sodium azide is added at 0.02% in all tests to
prevent microbial contamination.
5. The reaction is carried out at the optimum
temperature for 72 h at 150 rpm (see Note 12).
44
Enzymatic Saccharification of Lignocellulosic Biomass
6. Sampling is done every 1 h at the start of the
reaction, and every 3–12 h thereafter. Samples
should be centrifuged to remove residual biomass and then assayed for the release of reducing sugars [31] (see Note 10).
Saccharification of Biomass with Mixtures
of Enzyme Cocktails
1. The conversion of biomass to fermentable
sugars can be further enhanced by mixing different enzyme preparations together at various
proportions (see Note 13).
2. By quantifying the reducing sugars at regular
time intervals, addition of selective accessory
enzymes can be determined.
3. Saccharification should be carried out at optimized reaction conditions (see Note 14).
4. The total reaction volume should be identical
in all tests.
Recovery of Reducing Sugars
(Hydrolysates) from Enzyme
Pretreated Biomass
1. The saccharification process is terminated by
heating the enzyme pretreated biomass samples at 100 °C for 10 min (see Note 15).
2. The biomass sample is filtered through muslin
or centrifuged at 4,000 rpm for 10 min.
3. A 5-mL quantity of buffer is then added to the
residual biomass, and the components mixed
using a vortex for 5 min to recover the sugars
completely from biomass.
4. Store the hydrolysates at 4 °C until required
for analysis.
Analysis of Lignin-Derived Compounds
in the Hydrolysates
1. Test and control samples are centrifuged at
14,000×g for 15 min.
2. Lignin-derived compounds in the supernatant
are observed by measuring the absorption
spectrum between 200 and 465 nm using
UV-Vis spectrophotometer (Varian Cary 100,
USA), or in microassay format using a BioTek
479
Powerwave XS2 microplate reader in the
spectral scan mode, against the control supernatant from biomass pretreated with denatured
enzyme.
3. Compare the absorption values between control and test samples (see Note 16).
Quantification of Reducing and Total
Sugars in the Hydrolysates
1. Centrifuge the hydrolysates from test and control samples at 10,000×g for 10 min and the
supernatant is used for analyzing reducing
sugars.
2. The appropriate dilution of hydrolysates has
to be made.
3. Reducing sugars in the hydrolysates are
quantified by DNS reagent method [31].
4. Total sugars in the hydrolysates is quantified
by phenol-sulphuric acid method [24].
5. Calculate the saccharification (hydrolysis)
efficiency of enzyme cocktails from the
reducing sugars released in relation to the
total sugars present in the lignocellulosic
biomass [32].
Qualitative and Quantitative Analysis
of Reducing Sugars in Hydrolysates
1. The concentrations of mono-, di- and oligosaccharides in the hydrolysates can be determined by HPAEC equipped with UV and
pulsed amperometry ED40 gold electrode
detectors (Dionex, France).
2. Hydrolysate samples at various time intervals
are collected and centrifuged at 10,000×g for
5 min.
3. The supernatant is filtered through 0.2 mm
PDFE
membrane
syringe-less
filters
(Millipore, USA).
4. Elute the samples through the column
CarboPac SD-10 and CarboPac P100 for neutral monosaccharides and oligosaccharides,
respectively, using 50–100 mM NaOH, or
NaOH with increasing sodium acetate for oligosaccharide separation.
480
5. The flow rate is set at 1 mL per minute and the
flow cell temperature is kept at 30 °C.
6. Samples used for HPAEC analysis should be
diluted in such a way that the concentration
lies within the detectable range of the column
(see Note 17).
Notes
1. If specific plant biomass is used for any pretreatment, make sure it is not contaminated
before processing the samples. Rheological
factors may influence the nutrient contents of
biomass and therefore replicate biomass
samples should be collected from the same
sources.
2. Buffers should be freshly prepared each
week. Ionic strength and pH should be
checked carefully, otherwise this will
influence enzymatic hydrolysis.
3. Substrates used for enzyme assays should be
prepared freshly. Make sure that they are not
degraded, otherwise this will affect calculation of the exact activities.
4. Use HPLC-grade water only for sample
preparation. Samples should be filtered
through appropriate filters before analysis,
otherwise this will affect the HPAEC column
life and resolution.
5. If chemically pretreated sample is used for
enzymatic saccharification, they must be
washed properly until it reaches neutral pH.
It is very important to use the same batch
samples for all proximate analyses and
saccharification experiments. This will minimize the variation in the results.
6. Most of the commercial enzyme suppliers will
not give the details about the enzyme activity
and types of enzymes present in their cocktails. Key enzyme activities should be determined using the standard assay methods.
7. It is possible that the diluted enzyme will
lose its activity very quickly. Therefore, fresh
enzyme dilutions should be made whenever
required. Mix the contents properly before
use.
8. The optimal pH may vary for each commercial cocktail. After reviewing the results
M. Ayyachamy et al.
9.
10.
11.
12.
13.
14.
15.
16.
from tests conducted at different pH values,
further optimization studies using an
enzyme cocktail can be conducted at the
optimal pH.
In order to ensure that sugar released is not
due to the potential reducing sugar content of
an enzyme cocktail, a control run with denatured enzymes should be carried out.
Freeze the samples immediately or preferably heat at 100 oC for 15 min to terminate the
enzymatic action.
Key enzyme activities in the cocktail should
be determined to ensure uniform enzyme
dosage in all cases. If necessary, appropriate
dilutions have to be made using the appropriate buffers.
The optimal temperature and pH may vary
between two enzymes. If a big difference
does not exist in the conversion rate between
two temperatures, it is better to perform the
reaction at the lower temperature.
Some of the cocktails may not have the
required accessory enzymes. In this case it is
better to include accessory enzymes. This
will not only increase the conversion
efficiency but also potentially reduce the
reaction time.
It is expected that the optimal conditions will
not be same for two different enzymes.
Therefore, a compromise may be required
such that the best reaction conditions for one
of the enzymes is used preferentially, or to
compromise, reaction temperature is used
that will work for both enzyme preparations.
It is difficult to recover all of the released
sugars from the biomass. After filtering the
hydrolysates, the residual biomass has to be
washed with a minimum amount of buffer,
generally one third of the total reaction
volume.
Lignin-derived compounds in the hydrolysates can be compared with control samples.
Generally, lignin is directly or indirectly
removed due to enzymatic action. Increase in
the absorbance values of the tests in the wavelength range within which lignans and phenolics absorb light is an indication that the
enzyme has acted on lignocellulosic
biomass.
44
Enzymatic Saccharification of Lignocellulosic Biomass
17. Sufficient column washes should be carried
out between HPAEC analyses. Internal sugar
standards should be included in samples, as
appropriate to ensure that separation and
quantification are optimum.
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Protoplast Fusion Techniques
in Fungi
45
Annie Juliet Gnanam
Abstract
Traditionally, genetic manipulations of fungi were mostly dependent on
the conventional mutation method, which has severe limitations. In comparison, protoplast fusion technique offers great potential as a biotechnological tool for improvement in industrial strains, establishing genetic
systems, and overcoming incompatibility barriers in relative and nonrelative fungi. This technique has been successfully used to create recombinant strains with desired properties by fusion and also allowed for
transformation of desired gene/genes and viruses.
Keywords
Protoplast • Regeneration • Reversion • Fusion • KCl • Polyethylene
glycol
Introduction
The protoplast fusion technique and recombinant DNA technology have opened doors for
applied genetics in fungi. Protoplast fusion is an
important tool for gene manipulation because it
can break down the barriers to genetic exchange
imposed by conventional mating systems [1].
Removing the wall and exposing the protoplast
membrane allow for manipulation involving
fusion, or uptake of nucleic acids, processes that
A.J. Gnanam (*)
University of Texas at Austin, Institute for Cellular
and Molecular Biology, College of Natural Science,
2400 Inner Campus Drive, Austin, TX 78712, USA
e-mail: anniegnanam@yahoo.com
are less achievable or impossible with intact
cells [2]. Through protoplast fusion it is possible
to transfer desired traits such as disease resistance, nitrogen fixation, rapid growth, frost hardiness, drought resistance, herbicide resistance
and heat and cold resistance from one species to
another [3].
The isolation of protoplasts from microbial
cells involves the total digestion or localized
puncturing of the walls by enzymes, allowing the
cell contents enclosed by the plasma membrane
to escape. For survival as intact structures, the
protoplasts must be released into a hypertonic
solution to provide osmolarity [2]. Several commercial lysing enzymes are available for use in
the release of protoplasts in fungi. Lytic enzymes
singly or in combinations could be used for effective lysis of cell walls. Interestingly, several lytic
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_45, © Springer Science+Business Media, LLC 2013
483
484
enzymes originally used for lysis of cell walls in
filamentous fungi were made from culture supernatants of several fungal species [4–7], so making lytic enzymes in the laboratory for small-scale
use is a possibility.
The age of the mycelia markedly influences
the protoplast yield, as the susceptibility of the
cell walls to lytic enzyme/lenzymes is dependent
on it. Most frequently, cultures of the early to
mid-exponential phase have been used for good
protoplast yields [8–14]. However, stationary
phase cultures, spores and germ tubes have also
been used for protoplast preparations. The culture media used also has some effect on protoplast release [15]. Pretreatment with compounds
such as thiols has been shown to render the cell
walls susceptible to lysis [16–18].
The lysed protoplasts have to be released into
an osmotic buffer to keep them from bursting and
also to maintain their structural integrity. Several
inorganic salts and sugars in varying concentrations and pH have been used for this purpose.
The most commonly and successfully used inorganic salts, sugars and sugar alcohols include
potassium chloride (KCl), sodium chloride
(NaCl), magnesium sulphate, sorbitol, mannitol,
sucrose and glucose [8, 9, 19–25].
Not all protoplasts have the capability for cell
wall regeneration and subsequent reversion into
normal mycelial forms. The regeneration and
reversion of protoplasts are influenced by the
osmotic stabilizer in the medium, the carbon
source and the type of medium. In most cases, the
regeneration frequencies are higher than reversion frequencies. The ideal conditions for reversion could be determined before the use of
protoplasts in fusion experiments.
The protoplast fusion and transformation systems developed in the past several years have
led to our current understanding of phenomenon
such as genetic incompatibility and to exploit
economically important fungi used in the food
and pharmaceutical industries and also in agriculture. The technique has been widely used for
increased antibiotic production, antagonistic
potential and multipesticide resistance, to name
a few applications [26–28]. The most important
A.J. Gnanam
development in fusion of protoplasts happened
following the introduction of polyethylene glycol (PEG) as a fusogen, which leads to aggregation of the protoplasts. The concentration of
PEG and the type used play a very important
role in the successful fusion of protoplasts. The
general recommended concentrations are
between 20 and 50 %. Below 20 % the stabilizing effect is lost, whereas above the recommended concentration the protoplast starts
shrinking and results in lower fusion frequencies. The addition of the cation, Ca++, to PEG
stimulates the fusion of protoplasts and has been
widely used in the fusion of protoplasts of several fungi [9, 19, 20, 22–25, 28].
Development of fungal genetic transformation
systems was used to bring the power of genetic
analysis to species lacking sexual and parasexual
cycles [29]. Protoplast-mediated transformation
could be the optimal method when heterologous
integrations of multiple copies of a gene of
interest is envisaged to integrate at random sites
in the genome. This technique has been used to
increase protein yields considerably in several
fungi [30–33]. Transmission of double-stranded
RNA mycoviruses via protoplast fusion has
been reported in plant-pathogenic fungi such as
Aspergillus nidulans, Rosellinia necatrix and
Fussarium boothi [25, 34, 35].
Several selection procedures are employed to
identify and screen for protoplast fusants. The
most common methods of selection include
identification based on the difference in colony
morphology from the parent strains, using auxotrophic mutants, resistance/ susceptibility to
heavy metal, catabolite repressors and antifungal
agents [23, 24, 28]. The regeneration of the
fusants is generally carried out in solid or in liquid media under non-selective or selective conditions. The regeneration and reversion of the
protoplast to mycelia state could take between
several hours to days.
The literature on protoplast isolation, regeneration and fusion and the reasons for making
them varies depending on the organisms. By this
technique it is possible to manipulate genomes of
the organisms to a very high level.
45
Protoplast Fusion Techniques in Fungi
Materials
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
25.
26.
Inoculating loop.
Sterile disposable rod/glass rod.
Glass slide.
Cover slips.
Hemocytometer.
1-mL micropipettor.
1-mL sterile disposable plastic tips.
Incubator set at 28 °C.
Bunsen burner.
Orbital shaker set at 28 °C.
Vortexer.
Sterile Erlenmeyer flasks, 25- and 500-mL
capacity.
Microcentrifuge, e.g., Eppendorf, Centrifuge
5417C.
Sterile disposable polypropylene microcentrifuge tubes: 1.5-mL conical.
Sterile distilled water.
Potato dextrose yeast extract agar medium
(PDYEA) (peeled potato, 200 g; dextrose,
20 g; yeast extract, 3 g; agar, 20 g/L of distilled water at pH 6.5).
PDYE agar plates.
PDYE broth.
PDYEA plates with 0.6 M KCl, pH adjusted
to 5.5.
PDYEA plates with 0.6 M KCl, 0.5 mg/mL
benomyl and 100 mg/mL griseofulvin, pH
adjusted to 5.5.
Lytic enzymes—Novozym 234 (5 mg/mL),
chitinase (3 mg/mL), pectinase (2 mg/mL),
lysozyme (1 mg/mL) and cellulose (3 mg/
mL).
0.6 M KCl pH 5.5.
1 M CaCl2, autoclaved or filter sterilized.
PEG (polyethylene glycol, MW 3,500).
Sterile cheese cloth.
Phase contrast microscope, e.g., Carl Leitz
photo microscope.
Methods
The methods given below describe general procedures for protoplast release, fusion, regeneration and reversion in filamentous fungi. Different
485
enzyme combinations and concentrations may be
needed for effective release of protoplasts. The
age of the mycelium plays an important role in
protoplast release and may have to be altered for
different fungal species. The PEG used for fusion
of protoplast has to be picked up from a range of
3,000–6,000 MW and the concentration used
may have to be varied between 25 and 50 %. The
regeneration media has to be made as per the
requirement of the individual organisms used. In
most cases the complete media with the osmotic
stabilizers added is used. However, in some cases
minimal media could also be used. Suitable markers have to be developed for screening of the
fusant colonies. In case there is no defined marker
available, colony morphology, estimation of the
yield of desired product, etc., could be used for
selection.
Protoplast Isolation from Young
Hyphae (e.g., Trichothecium roseum)
The following method has been used to isolate
high number of protoplasts from the filamentous
fungi Trichothecium roseum, Trichoderma harzianum, Trichoderma reesi, Trichoderma longibrachiatum, Trichoderma viride, Dreschlera
oryzae and Venturia inaequalis [8–13, 36].
1. The conidia were collected from 7-day-old
plate cultures by adding 3–5 mL sterile water
to the agar plate and gently scraping the surface with a sterile inoculation loop. The conidial suspension was drawn with a 1-mL
micropipettor and transferred into a sterile
1.5-mL microcentrifuge tube.
2. The conidial suspension was then washed
twice with sterile water (1.2 mL) by centrifugation at 500×g for 10 min at room temperature. The final pellet was resuspended in
appropriate volume of water to bring the concentration of the conidia to 1 × 106/mL.
3. This conidial suspension was grown in 100 mL
PDYE broth at 28 °C with shaking conditions
for 24 h at 200 rpm.
4. The young mycelia (100 mg wet weight) were
harvested by filtration through triple-layer
sterile cheese cloth. It was then washed thrice
with 5 mL sterile distilled water, followed by
486
3–4 washes with 5 mL sterile osmotic stabilizer (see Note 1).
5. The washed mycelia were scooped with a sterile loop and added to the lytic enzyme mix
(see Note 2) and shaken at room temperature
(28 ± 2 °C) for 4 h at 200 rpm (see Note 3).
6. After 4 h, the undigested hyphal material was
removed by filtering the solution through 6
layers of sterile cheese cloth. To the resultant
filtrate equal volume of 0.6 M KCl was added
and the contents transferred into two sterile
1.5-mL microfuge tubes and centrifuged at
500×g for 5 min at room temperature. The
supernatant was carefully removed with a
micropipette and the resulting protoplast pellet was resuspended in 1 mL 0.6 M KCl.
PEG-Meditated Protoplast Fusion,
Regeneration, Reversion and Selection
(e.g., Trichothecium roseum Isolates
AJ102 and AJ 210)
The following method has been widely used for
self-, inter- and intra-specific protoplast fusions
in filamentous fungi and yeasts [9, 20, 22–24, 27,
28]. The success of this method lies in the ability
of the protoplasts to regenerate and revert into
normal vegetative forms after fusion.
1. Follow the steps in Protoplast Isolation from
Young Hyphae (e.g., Trichothecium roseum),
from steps 1–6, for both isolates. Adjust the
concentrations of protoplasts to 1 × 106 with the
osmotic stabilizer, 0.6 M KCl (see Note 4).
2. 1 mL of each of the protoplast suspension was
mixed with 2 mL of 80 % PEG solution (see
Note 5) and 0.2 mL of 1 M CaCl2 and incubated
at room temperature for 15 min (see Note 6).
3. The suspension was then diluted with 6 mL of
0.6 M KCl and incubated again for 15 min at
room temperature and vortexed for 5–10 s (see
Note 7).
4. The protoplast suspension was then transferred to 1.5-mL sterile microfuge tubes
and harvested by centrifugation at 500×g for
5 min at room temperature. The supernatant
was carefully removed with a micropipettor
and the protoplast pellets were pooled into a
1.5-mL sterile microfuge tube.
A.J. Gnanam
5. The pooled protoplasts were washed twice
with 1.2 mL of 0.6 M KCl and resuspended in
the same.
6. The fused protoplasts were then spread plated
on selective and non-selective agar plates at a
concentration of 1 × 107 (see Notes 8–10).
7. The plates were incubated with the right side
up at ±28 °C for 24 h and then inverted the
next day.
8. The fusant colonies appeared after 3 days.
Single colonies were isolated from the selection media. The fusion frequency was calculated based on the ratio of the number of
colonies on selective and non-selective agar.
The colonies were subcultured several times
on selective and non-selective agar before
further studies.
Notes
1. The osmotic solution, 0.6 M KCl, pH 5.5,
was autoclaved, cooled, and stored at room
temperature. Care was taken to keep the
concentration accurate, by ensuring that the
liquid was not lost due to evaporation during
autoclaving.
2. The lytic enzyme solution was made less
than 20 min before use and filter sterilized
with a 0.22-mm filter into a sterile 25-mL
Erlenmeyer flask, the top of which was
plugged with a cotton plug.
3. At every 30-min interval, samples were
drawn and observed under the microscope to
determine the maximum protoplast release.
Ten microlitres of the sample were placed on
a clean glass slide and a cover slip was gently
placed over it for viewing under the microscope. A hemocytometer was used for counting the number of protoplasts.
4. The protoplasts were washed thoroughly to
remove any traces of the lytic enzymes and
the PEG, for effective regeneration and
reversion of the fusants. The samples have to
be handled carefully and gently, as the protoplasts are extremely fragile.
5. For the 80 % PEG solution, PEG 3,500 MW
was dissolved in 0.6 M KCl solution, the pH
adjusted to 5.5 and autoclaved. Before use
45
6.
7.
8.
9.
10.
11.
12.
Protoplast Fusion Techniques in Fungi
the solution was filter sterilized with a 0.22mm filter. The solution containing the protoplasts and the PEG solution were mixed in
equal volumes to bring the concentration of
PEG to 40 %.
The incubation period with PEG could be
variable for different organisms and protoplast preparations.
Step 3 was performed to dislodge the loosely
bound protoplasts and the aggregates that did
not fuse.
Selective agar plates were usually regular
growth media, with the osmotic stabilizer and
also markers such as heavy metals, fungicides
or aminoacids (in the case of auxotrophic
mutants) at appropriate concentrations.
Isolates Trichothecium roseum AJ102 was
sensitive to benomyl and resistant to griseofulvin; AJ210 was resistant to benomyl and
sensitive to griseofulvin. The concentrations
required for selecting the fusants was determined as 0.5 mg/mL benomyl and 100 mg/mL
griseofulvin.
Selective medium (PDYEA plates with
0.6 M KCl, 0.5 mg/mL benomyl and 100 mg/
mL griseofulvin, pH adjusted to 5.5); nonselective medium (PDYEA plates with 0.6 M
KCl, pH adjusted to 5.5). For the selective
medium PDYEA medium was autoclaved
and cooled down to 55 °C. The osmotic stabilizer (KCl) and the antifungal compounds
benomyl and grisefulvin were added to it at
0.6 M, 0.5 mg and 100 mg/mL concentrations,
respectively, mixed thoroughly and poured
into Petri plates. The Petri plates were
allowed to cool and left at room temperature
for 24–48 h before use.
Plating usually involved using 100 mL protoplasts suspension per plate that was spread
gently over the agar plate with a sterile disposable rod or glass rod. Extreme care was
taken during plating as the protoplasts tend
to be very fragile at this stage.
It might take several days for the fusant colonies to be visible on the selection plates. So it
is better to leave the plates for several days in
the incubator.
487
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Murugesan K, Lalithakumari D (2006) Self-fusion of
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Technol 97:2330–2334
10. Mrinalini C, Lalithakumari D (1996) Protoplast
fusion: a biotechnological tool for strain improvement
of Trichoderma sp. Curr Trend Life Sci 21:133–146
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regeneration of protoplasts from mycelium of
Dreschlera oryzae. J Plant Dis Prot 98:197–204
12. Karpagam, S. (1994) Variations in the protoplast
regenerated isolates of Trichoderma harzianum. M.
Phil. Thesis, University of Madras, Chennai, India.
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the formation of protoplasts in Aspergillus niger.
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16. Fawcett PA, Loder PB, Duncan MJ, Beesley TJ,
Abraham EP (1973) Formation and properties of protoplasts from antibiotic-producing strains of
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biotechnological tool. Oxford/IBH, New Delhi
Large-Scale Production
of Lignocellulolytic Enzymes
in Thermophilic Fungi
46
Manimaran Ayyachamy, Mary Shier,
and Maria G. Tuohy
Abstract
A semi-pilot scale process for the production of lignocellulolytic
enzymes in the thermophilic fungus Talaromyces emersonii is presented.
Each step involved in the scale-up process is described in detail.
Keywords
Lignocellulolytic enzymes • Thermophilic fungi • T. emersonii
• Thermozymes • Fermenter • Submerged cultivation
Introduction
Talaromyces emersonii, a thermophilic,
filamentous euascomycete fungus, has been
reported as one of the potential candidates for
lignocellulolytic enzymes production [1, 2].
T. emersonii produces highly specific thermozyme cocktails with efficient catalytic and
long-term storage properties [3–6]. These thermostable fungal enzymes usually work 10–20 °C
M. Ayyachamy (*) • M.G. Tuohy
Molecular Glycobiotechnology Research Group,
Department of Biochemistry, School of Natural Sciences,
National University of Ireland, University Road,
Galway, Ireland
e-mail: manimaran.ayyachamy@nuigalway.ie
M. Shier
Department of Biochemistry, National University
of Ireland, University Road, Galway, Ireland
e-mail: mary.shier@nuigalway.ie
higher than commercially available Trichoderma
sp. enzymes [7].
T. emersonii has the ability to produce a wide
range of extracellular polysaccharide degrading
enzymes in higher amounts than bacteria or yeast
[3]. Owing to the fact that enzymes are secreted
into the culture medium, the downstream processing of enzymes is relatively simple. Enzyme systems from this fungus have been reported already
and patents have been developed for significant
applications [2, 5, 6]. Plant polymer hydrolysis
studies using these crude thermozymes are usually carried out at high temperatures, which
decrease reaction time, minimize microbial contamination and lower hydrolysate viscosity. The
aforementioned catalytic properties of thermozymes are desirable characteristics for several
biotechnological and industrial applications.
Thermozymes production by T. emersonii
using different plant polymeric substrates and
low-cost approaches has already been reported
[2, 8]. Thermozymes have also been evaluated
for food and biomass conversion applications
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_46, © Springer Science+Business Media, LLC 2013
489
490
[7, 9–11]. These promising results undoubtedly
indicate that lignocellulolytic thermozymes from
T. emersonii have potential applications in food,
biofuels and biorefineries sectors.
Materials
1. Fungal strain: Talaromyces emersonii
IMI393751 (see Note 1).
2. Sarbouraud dextrose agar (SDA) from Oxoid
(Basingstoke, UK).
3. TG medium: 2% (w/v) Glucose and 201 mL
of 5× nutrients solution in 80 mL water.
4. Nutrient solution (5×) for TG (g/L): (NH4)2 SO4,
75; KH2PO4, 25; Corn steep liquor, 25 mL;
Yeast extract, 25; FeSO4.7H2O, 0.312; ZnSO4.
7H2O, 0.0625; H3BO3, 0.0625; MnSO4.4H2O,
Na2MoO4, 0.0625; CoCl3.6H2O, 0.0625; KI,
0.0625; MgSO4.7H2O, 2.5; CaCl2.2H2O,
2.5 g; Na2SO4, 5. Stir well and adjust to a
final pH of 4.5 with 1 N NaOH.
5. EI medium: 2% (w/v) wheat bran in water.
6. Inexpensive carbon source: Wheat bran was
collected from a local supplier.
7. Antifoam: Proflo Oil (Traders Protein,
Tennessee, USA).
8. Buffers: pH 4.0, 7.0 and 11.0.
9. Electrolytes for pO2 probe.
10. Steam
generator
(Fibrimatic/Camptel
Dispositivo H2 48 kW minimum).
11. Air compressor (Bambi VT300D, oil-free
350 L/min, 8 bar, 3HP compressor, fitted
with water condenser unit).
12. Cooling water supply pipeline.
13. Erlenmeyer flasks: 500 mL and 1 L.
14. Thermostatically controlled NewBrunswick
Scientific Innova 44 platform, shaking incubator (New Brunswick Scientific, USA).
15. BIOSTAT® B-plus 10 L Fermenter (Sartorius
BBI Systems, Germany).
16. BIOSTAT® D-plus 150 L Fermenter, equipped
with control cabinet, touch screen controller
and SCADA Fermenter Supervisory Control
and Data Acquisition software (Sartorius
BBI Systems, Germany).
17. Magnetic stirrer (Fisher Scientific, Dublin,
Ireland).
M. Ayyachamy et al.
18. Arium 61315 Reverse Osmosis (RO) water
system with 360 L/day capacity, equipped
with RO modules (Sartorius BBI Systems,
Dublin, Ireland).
19. RO water 150-L capacity storage tank
(Sartorius BBI Systems, Dublin, Ireland).
20. Trolley mounted Westfalia HSD® SD1 centrifuge with integrated control panel
(Westfalia Separator AG, Germany).
21. Sartocon2 Sartoflow® membrane filtration
unit (MF/UF crossflow cassette system),
equipped with a Sartojet® pump and
fitted with either 5 kDa or 10 kDa Hydrosart
membranes (Sartorius BBI Systems,
Germany).
22. BioTek Powerwave XS2 microplate reader
with incubation facilities to 50 °C (BioTek,
USA).
23. Filters: Sarstedt (0.2-mm and 0.4-mm filters).
24. Substrates: wheat arabino xylan (Megazyme
International Ireland Ltd., Bray, Co.
Wicklow), oat spelts xylan (SigmaAldrich, Dublin, Ireland), birch wood xylan
(Sigma-Aldrich), b-glucan (from barley;
Megazyme Intl. Ireland Ltd.), and carboxy
methyl cellulose and 4-nitrophenyl a- or
b-glycosides (Sigma-Aldrich). Unless otherwise stated, all chemicals and reagents were
purchased from Sigma-Aldrich (Dublin,
Ireland).
25. High-quality 20-L and 100-L plastic storage
containers (Nalgene, UK, and Fisher
Scientific, Ireland).
26. Laboratory plastics, i.e., disposable
micropipette tips, microcentrifuge tubes
and flat-bottomed, high-optical-quality, lowprotein-absorbing microtitre plates (Sarstedt,
Sinnottstown, Co. Wexford, Ireland).
27. Safety apparel (Caulfields Ltd., Galway,
Ireland).
Methods
This section has been divided into seven subsections. The overall steps involved in the scale-up
process are elaborated.
46
Large-Scale Production of Lignocellulolytic Enzymes in Thermophilic Fungi
Pre-Inoculum Preparation
1. A ~1 cm2 piece of T. emersonii IMI393751,
either from a routinely sub-cultured SDA agar
plate or a glycerol stock, is transferred aseptically onto a fresh SDA agar plate and incubated at 45 °C for 48–72 h (see Note 1).
2. To prepare a submerged (liquid) culture of the
fungus, take 3–4 ~1 cm2 pieces of T. emersonii
IMI393751 mycelial mat (small section) from
SDA agar plate transferred into 500-mL
Erlenmeyer flasks containing 100 mL TG
medium.
3. Incubate the culture flasks at 45 °C with shaking (200 rpm) for 48 h.
3.
4.
5.
6.
Inoculum Development in a 10-L
Fermenter
7.
1. Prepare and sterilize 6 L of TG medium in an
autoclave (Sanyo Labo autoclave, Sanyo,
USA) at 121 ° C for 15 min (see Notes 2
and 3).
2. Prepare a seed culture of T. emersonii (36- to
48-h old) by inoculating this medium with T.
emersonii mycelial suspension taken from the
pre-inoculum shake flasks culture under aseptic conditions; a 5% (v/v) inoculum is used.
3. Agitation, aeration rate and temperature are
maintained at 200 rpm, 1.5 vvm and 45 ° C,
respectively.
4. Dissolved oxygen should be maintained at
20% using the cascade mode (see Note 4).
5. Foaming is controlled by automatic addition
of antifoaming agents.
6. At the end of 36–48 h of cultivation, the fungal cultures are harvested into a sterile container in a laminar air-flow chamber.
Fungal Cultivation in a 150-L Fermenter
1. Utilities (steam generator, air compressor
and water pipe lines) were thoroughly
checked prior to start of the fermenter run.
2. All utilities’ valves should be opened. Make
sure that the pressure bar values meet the
8.
9.
10.
11.
12.
13.
491
technical specifications of the fermenter (see
Notes 5–7).
Calibrate the pH and pO2 probes according to
the instructions given by the manufacturer.
Fill the fermenter with 100 L water from the
storage tank (Sartorius Arium 61315), which
is connected to a Sartorius RO purification
system.
Add wheat bran at 2% (w/v) and mix it thoroughly with the water by setting the agitation
rate at 400 rpm.
Before initiating the sterilization process, a
pressure hold test for the vessel should be
performed to ensure that there is no leakage
in the vessel. This is achieved by closing the
air inlet and gas vent valves manually and
checked to see whether the pressure in the
vessel is maintained over a holding period of
30–45 min (see Notes 8 and 9).
Set the cultivation and sterilization temperatures and cultivation period using the DCU
(digital control unit) cocabinet, which is
connected to an MS Windows-based
MFCS (Multi Fermenter Control System)
SCADA, and allows data acquisition and
batch management.
Sterilize the contents of the fermenter (in
situ) at 105 °C for 30 min.
At the end of sterilization process, allow the
fermenter to cool down automatically until it
reaches the cultivation temperature; the cultivation temperature has been set already
during the sterilization process.
The T. emersonii cultures (36- to 48-h old),
developed in a 10-L reactor, are used to inoculate the 150-L fermenter at 5% (v/v) under
aseptic conditions (see Note 10).
Antifoaming agent (20 mL) is added soon after
inoculating the cultures by setting the antifoam
pump into manual mode. The antifoam pump
is then set back to automatic mode.
Cultivation parameters are the same as
described in the section Inoculum
Development in a 10-L Fermenter.
Samples should be collected from two different sampling ports every 24 h and the desired
enzyme activities and total sugar analysis
determined (see Notes 11–13).
492
14. At the end of cultivation, the culture broth is
mixed thoroughly at 300 rpm for 15–20 min
before harvesting.
M. Ayyachamy et al.
3.
Harvesting and Concentrating
the Enzyme Cocktails
1. Because the target enzymes produced during
fermentation are secreted (i.e., extracellular),
fungal broth cultures are harvested by separating the mycelia (fungal biomass) from the fermentation broth in a high-speed Westfalia
SD1 discharge centrifuge, with integrated
control panel. The contents of the 150-L fermenter are pumped to the centrifuge using a
Sartojet (or other) pump, by a connecting line
(Sartorius BBI Systems). The mycelial mass
and left-over substrate are trapped in the centrifuge bowl and subsequently removed (see
Note 14).
2. The supernatant from the HSD centrifuge is
pumped into a Sartoflow microfiltration (MF)
unit using a Sartojet pump, to remove residual
mycelia and particulate material. The microfiltered crude enzyme is then concentrated
using the Sartoflow system, this time operating the ultrafiltration mode. Fold concentrations vary from 20- to 50-fold, depending on
the desired concentration factor and the viscosity of the enzyme product. The unconcentrated and concentrated enzyme samples are
then assayed for protein content and enzyme
activity.
3. All unconcentrated and concentrated enzyme
samples should be stored at 4 °C (or at −20 °C
for longer-term storage) until required.
4.
5.
6.
7.
specified [2, 4–10]. Each test reaction and all
controls are prepared in triplicate.
The hydrolysis of wheat arabino xylan
(1% w/v, 15 min), oat spelts xylan (1% w/v,
15 min), birch wood xylan (1% w/v, 15 min),
b-glucan (from barley; 1% w/v, 15 min) and
carboxy methyl cellulose (1% or 3% w/v;
30 min) are determined as reducing sugars
released by the dinitrosalicyclic acid (DNS)
reagent method [2, 4–8, 12–14].
Absorbance values are measured at 550 nm in
a 96-well microplate reader equipped with the
appropriate filters (BioTek Powerwave XS2
microplate reader, BioTek, USA).
Exo-acting glycosyl hydrolase activities,
including acetyl esterase (4-nitrophenyl
acetate), b-glucosidase (4-nitrophenyl-b-Dglucopyranoside), a-glucosidase (4-nitrophenyl
a-D-glucopyranoside),
b-xylosidase
(4-nitrophenyl b-D-xylopyranoside) and a-Larabinofuranosidase
(4-nitrophenyl-a-Larabinofuranoside), were determined using
the appropriate 1 mM 4-nitrophenyl a- or
b-glycoside as substrate [2, 4, 5].
Suitably diluted enzyme is incubated with the
appropriate substrate for 15–30 min at 50 °C
and the reaction is stopped using 1 M Na2CO3.
The increase in absorbance at 405 nm on
release of the nitrophenolate anion is measured in a 96-well microplate reader equipped
with the appropriate filters (BioTek, USA).
Enzyme activity is expressed as IU, i.e., m mol
of reducing equivalents (xylose, glucose or
nitrophenol) released/mL of enzyme/min reaction time, under standard assay conditions.
Total Protein and Sugar Analysis
Determination of Enzyme Activities
1. The crude enzyme samples are centrifuged
prior to assay at 1,000 × g for 10 min and the
clarified supernatant is used as crude enzyme
for enzyme assays, protein and reducing sugar
analysis (see Note 8).
2. All enzyme assays are conducted at a working
assay temperature of 50 °C in sodium acetate
buffer (100 mM, pH 5.0), unless otherwise
1. The protein content in the crude extracellular
enzyme was determined preferably by the
Bensadoun and Weinstein modification of the
method of Lowry et al. [15] or by the Bradford
method [16], using bovine serum albumin
(BSA, fraction V) as standard.
2. The total sugars in the culture broth is
quantified using the phenol-sulfuric acid
method; [17] reducing sugars are quantified
by DNS reagent method [14].
46
Large-Scale Production of Lignocellulolytic Enzymes in Thermophilic Fungi
Monitoring the Fungal Cultivation
Temperature, pH and dissolved oxygen are monitored by the Biostat D+150, which is equipped
with a DCU-control cabinet system. The DCU
system is a local control system for the automation of the fermentation process. Thus, growth
parameters for T. emersonii submerged cultures
are automatically monitored in the fermenter.
9.
Notes
1. T. emersonii is routinely sub-cultured on
SDA plate at 45 °C.
2. All ports in the fermenter should be clamped
properly. Make sure that one of the ports is
plugged with non-absorbent cotton wool, if
the fermenter is not autoclaved with gas
cooling exit device.
3. The pH probe must be calibrated prior to
sterilization. The pO2 probe should be calibrated after sterilization. All probes and the
motor must be wrapped with aluminium foil
while sterilizing the fermenter.
4. Dissolved oxygen is one of the critical
parameters for fungal cultivation. Make sure
that the set agitation rate values during the
cascade mode operation do not rupture the
fungal mycelia.
5. Turn on the steam generator first and allow it
to reach 5 bar. Open the steam valve and set
the supply pressure value to 3–3.5 bar. Steam
filters should be replaced after three fermenter runs.
6. The cooling water supply pressure should be
maintained between 3 and 3.5 bar.
7. The water condensate from the air compressor should be removed completely before
passing the air into the fermenter. All automatic valves in the pipeline are manufactured
in such a way that they operate only in
the presence of air. It is important that the
pressure should be maintained between 5 and
6 bar throughout the fermenter run.
8. A leakage test is important to make sure that
all of the ports are sufficiently tightened. If
leakage occurs in the vessel, it will be difficult
10.
11.
12.
13.
14.
493
to attain the desired temperature. First, the
fermenter gas vent should be closed manually and then air passed into the vessel until it
reaches 1 bar in pressure. The pneumatic and
air inlet valves are then closed manually.
After performing the leak test, all valves
should be returned to the automatic mode setting. Before starting the sterilization, steam
in the pipelines should be released through
the drain valves to ensure that high-quality
steam only is used for sterilization. Gas cooling exit pipes should be kept open so that liquid loss in the vessel can be prevented.
The inoculation port and nearby surface
should be cleaned with 75% ethanol. A multitorch or spirit lamp can be used to create
sterile conditions in and around the inoculation port. Minimize the exposure time when
opening the inoculation port during
inoculation.
Before and after collecting the samples from
the fermenter, the sampling port should be
sterilized with steam for 5–10 min. This will
prevent microbial contamination.
All substrates used for enzyme assays
should be freshly prepared and the appropriate standards curves should be conducted in
parallel.
The enzyme cocktails used for the assay
should be diluted appropriately with buffer
prior to enzyme assay.
When the HSD centrifuge bowl becomes
clogged, the resulting supernatant becomes
dirty and unclear. To clear the bowl, a full
discharge is required.
References
1. Stolk AC, Sampson RA (1972) The genus Talaromyces:
studies in mycology, 2nd edn. Centraal bureau Voor
Schimmelcultures Publishers, Baarn, The Netherlands
2. Tuohy MG, Murray PG, Gilleran CT, Collins CM,
Reen FJ, McLoughlin L et al. (2007). Talaromyces
emersonii enzyme systems. Patent WO/2007/091231.
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(2003) Catalytic properties and mode of action of
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Panfungal PCR Method for
Detection of Aflatoxigenic Molds
47
Malik M. Ahmad, Pravej Alam, M.Z. Abdin,
and Saleem Javed
Abstract
Aflatoxigenic molds contaminate the processed or raw agricultural
commodities during their storage and transportation. Traditional methods
offer a laborious and tedious process; also, detection can be performed
only after toxin production. Molecular methods provide a rapid, sensitive,
and specific detection. The major advantage of molecular methods is that
the detection can be observed prior to the toxin production. Molecular
methods for fungal detection can be applied to distinguish the aflatoxinsecreting fungi from non-aflatoxigenic molds. The DNA-based PCR
amplification of a consensus sequence can make it possible to monitor the
range of aflatoxigenic fungi from contaminated products.
Keywords
Aflatoxigenic molds • DNA • Consensus sequence • Panfungal PCR
• Molecular detection
Introduction
The past two decades have witnessed various
scientific innovations in diagnostic microbiology
owing to technological advances in molecular
M.M. Ahmad • P. Alam • M.Z. Abdin (*)
Department of Biotechnology, Hamdard University,
New Delhi, 110062, India
e-mail: mzabdin@rediffmail.com
S. Javed
Department of Biochemistry, Hamdard University,
New Delhi, 110062, India
microbiology. For example, speedy detections of
nucleic acid-based amplification process, with its
mutual characterization through automated and
user-friendly bio-computational tools, have
significantly widened the area of diagnostic
detections for clinical microbiologists [1]. The
accessible traditional methods for aflatoxin detection are time-consuming, labor-intensive, difficult
to standardize, have low sensitivity and nonspecificity, and require days to weeks before the
results are achieved. Serological methods work
on the “one substance one assay” concept [2, 3].
With such performance, researchers have diverted
their scientific thoughts toward molecular
advancements [4]. Most molecular diagnostic
assays use polymerase chain reaction (PCR).
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_47, © Springer Science+Business Media, LLC 2013
495
496
It has become one of the easiest methods, as it
rapidly amplifies the target DNA into millions
of copies, for detection of microorganisms
[4]. Variations in the technique increased the
sensitivity of the detection method (nested PCR),
while multiplex PCR detects a number of
pathogens simultaneously. Nested PCR, however, employs two primer sets (outer and inner),
whereas multiplex PCR uses a series of primer
sets for identification. The basic problem with
both these PCR variants is that the annealing
temperature for each primer sets is first optimized
to work correctly within a single reaction. This
problem can be resolved by having a single consensus sequence of multiple microorganisms for
primer design and its amplification by conventional PCR. In a similar way, phytopathogenic
molds can also be detected by this method, especially those secreting mycotoxins such as
aflatoxins. Aflatoxigenic molds can directly damage crops and indirectly impair human beings, as
the latter ingest mycotoxin-contaminated agricultural products. These are aflatoxin-secreting
fungi group, particularly from Aspergillus species [4]. Aspergillus flavus and A. parasiticus are
the two prominent candidates that secrete the
extrolites, although there are other aflatoxinsecreting species, such as A. tamarii, A. pseudotamarii, A. bombycis, A. oryzae, A. nomius,
A. parvisclerotigenus, A. minisclerotigenes,
A. toxicarius, A. versicolor, and so forth, with
some other genus like Emericella astellata and
E. nidulans [4]. The following protocol can be
used to detect aflatoxigenic molds from agricultural commodities, including foods and feeds.
Materials
Chemicals and Glasswares
1. Nuclease-free water
2. Agarose (molecular biology grade, HiMedia)
3. TAE buffer: 50 mM Tris (HiMedia), 50 mM
acetic acid (HiMedia), 1 mM EDTA (HiMedia).
Dilute when needed from a 50 × stock to 1 ×
for use (see Note 1)
4. Ethidium bromide (EtBr; Sigma-Aldrich):
5.0 mg/mL stock (see Note 1)
M.M. Ahmad et al.
5. Gel-loading dye: 60 % glycerol, 60 mM
EDTA, 0.3 % bromophenol blue, 0.3 % xylene
cyanol, 10 mM Tris–HCl (pH 7.6) (MBI
Fermentas)
6. Molecular marker 100 bp: ready to use (MBI
Fermentas)
7. Disposable polypropylene micropippet tips
(Tarsons): 200 mL and 20 mL (see Note 1)
8. Disposable polypropylene microcentrifuge
tubes (Tarsons): 1.5 mL, tight-fitting snapcapped (see Note 1)
9. PCR tubes (Tarsons) (see Note 1)
Equipment
1. Biofuge (Heraeus Pico)
2. Thermal cycler (GStorm)
3. Horizontal electrophoresis equipment (BioRad
Wide Mini Sub Cell)
4. Gel documentation system (UVI Tech)
PCR Reagents
1. Template DNA of aflatoxigenic and nonaflatoxigenic fungi
2. Oligonucleotide primers (custom-made,
Sigma-Aldrich, USA) re-suspended to a concentration of 100 mM using nuclease-free
water and stored at −20 °C
3. dNTPs: a mixture of dATP, dCTP, dGTP,
and dTTP (2.5 mM each, Merck), stored at
−20 °C
4. Thermostable DNA polymerase (Taq) and
reaction buffer supplied by manufacturer
(Merck). We typically use 10 × reaction buffer
of 100 mM Tris pH 9.0 (25 °C), 500 mM KCl,
15 mM MgCl2, and 0.1 % gelatin. Separate
stocks of MgCl2 (25 mM) are also supplied for
adding to the reaction. All of these reagents
are stored at −20 °C
Method
The method given as follows describes the general procedure for primer designing of a consensus sequence from aflatoxin-secreting fungi
47
Panfungal PCR Method for Detection of Aflatoxigenic Molds
followed by its amplification and subsequent
testing with non-aflatoxigenic molds. The volumes and number of tubes used per sample may
vary, depending on the type of sample and the
number of fungi being processed.
Multiple Sequence Alignment
and Primer Design
The most commonly used target for molecular
fungal detection is DNA encoding the ribosomal
RNA genes including ITS regions, but other
targets include housekeeping genes such as
b-tubulin, calmodulin, actin, and cytochrome b
[5–9] for consensus sequence selection and
primer design. Thus, the primers amplifying these
regions show conserved sequences in nature and
that is why they are known as internal controls.
However, to detect the aflatoxigenic molds at the
molecular level, one has to look for a consensus
region present in all the aflatoxin-secreting species. The following steps are involved in the
selection of consensus sequence and its primer
designing:
1. To identify a consensus region, all the genes
of A. flavus and A. parasiticus involved in
aflatoxin biosynthesis were collected in
FASTA format from the NCBI Genbank. For
example, collect the sequences of aflP gene
from all strains of A. flavus and A. parasiticus
(see Note 2).
2. Compare their similarities through multiple
sequence alignment (ClustalW2, EBI tools)
using default settings. Gene sequences of aflP
of both fungal strains are matched for
similarity.
3. Deduce a consensus sequence from the alignment that provides maximum identity. For
example, aflP gives 96 % similarity. So, a
sufficient consensus sequence can be obtained
for primer design. Mark the mutations present in between the consensus sequence (see
Note 3).
4. Design a primer set for the consensus sequence
for PCR amplification considering all parameters (see Notes 4 and 5).
497
PCR Amplification
After DNA isolation, quantification, and primer
design, PCR amplification of the aflatoxigenic
molds can be performed. Detection of aflatoxin
production before its secretion into food products
is critical for maintaining the quality and economic value of those products. Thus, PCR method
by amplification can forecast the prior production
of aflatoxins and thus the quality of products can
be taken care of. The following steps are involved
in PCR amplification:
1. Keep all reagents and DNA samples on ice
until used. Prepare adequate reaction mixture
for the number of samples to be tested. To
detect aflatoxigenic molds, for example, take
four fungal species (two known for aflatoxinproducing Aspergillus species, A. flavus, and
A. parasiticus; one non-aflatoxigenic
Aspergillus species; and the other Fusarium
species). Prepare PCR “master mix” of 92 mL
containing 64 mL nuclease-free water, 8 mL
10 × PCR buffer, 8 mL 10 mM dNTP mix,
4 mL each primer, and 4 mL Taq polymerase
(5 U/mL). Mix reagents properly through
micropipette and finally vortex and centrifuge briefly to remove the air bubbles before
dispensing 23 mL aliquots into four PCR
tubes and place on ice (see Note 6).
2. Add template DNA (100 ng) of 2 mL to each
of the tubes, using microtip and mix the content briefly (see Note 7).
3. Set a PCR for positive control reaction with
18S ribosomal amplifying region as internal
control and a separate multiplex PCR by
mixing 18S ribosomal and omtA primers in
another PCR master mix for all four fungal
species with same cycling programme (see
Note 8).
4. Put the PCR tubes in thermal cycler and run
the appropriate cycling program: initial denaturation of 1 min at 95 °C followed by 30
amplification cycles of 1 min at 94 °C, 30 s at
56.7 °C and 1 min at 72 °C, with final extension of 2 min at 72 °C.
5. As the amplification completes, analyze 5 mL
of each sample by agarose gel electrophoresis.
498
M.M. Ahmad et al.
Fig. 47.1 Detection of aflatoxigenic and nonaflatoxigenic molds by PCR amplification. Lane M1,
100 bp marker; Lanes 1, 4, 7, 10, 13, amplification of
internal control (18S ribosomal gene) of A. flavus, A. parasiticus, plant-root isolated A. flavus, A. versicolor,
Fusarium sp.; Lanes 2, 5, 8, multiplex PCR-positive
control; Lanes 11, 14, multiplex PCR-negative control;
Lanes 3, 6, 9, PCR of omtA gene from A. flavus, A. parasiticus, and plant-root isolated A. flavus-positive detection; Lanes 12, 15, PCR of omtA region in non-aflatoxigenic
mold (A. versicolor and Fusarium sp.)—negative detection; Lane M2, 1 Kb marker
6. Prepare a gel, add 1.2 % agarose to 1 × TAE
buffer, and heat to dissolve (e.g., using a
microwave oven) (see Note 9).
7. Add EtBr of 3.5 mL per 100 mL of concentration 5 mg/mL, in agarose, and allow to cool
at approximately 40 °C or till when your skin
feels suitable (see Note 10).
8. Pour the agarose gel into a suitable gel casting tray with teflon comb and allow to set for
approximately 20 min.
9. Remove the comb and place the gel in a horizontal electrophoresis tank containing 1 ×
TAE, so that the wells are just covered with
buffer.
10. Add 2 mL of gel-loading dye in 5 mL of each
PCR samples, transfer the mixture into wells
of agarose gel.
11. Pour 2 mL of each 100 bp and 1 kb molecular
marker to the first and last wells and, finally,
electrophores (see Note 11).
12. Visualize the agarose gel on gel documentation system and capture the photograph after
proper separation of markers and amplicons
(Fig. 47.1).
Notes
1. All the microtips, microcentrifuge tubes,
PCR tubes, and necessary reagents should be
autoclaved properly.
2. Select a gene whose product converts penultimate precursor or immediate precursor to
aflatoxin. Thus, all the aflatoxins (B1, G1, B2,
and G2) are covered for the detection purpose.
3. While selecting consensus sequence, it would
be better to consider exonic region for primer
design. Exons are the transcribed regions of the
any genes and thus mostly conserved in nature.
Therefore, these regions have higher similarity
47
4.
5.
6.
7.
8.
9.
10.
11.
Panfungal PCR Method for Detection of Aflatoxigenic Molds
and have large enough consensus region. Thus,
all the aflatoxigenic molds that secret aflatoxin
can be included for detection while those which
are non–aflatoxin-secreting will not be
amplified. These steps will finally enhance the
specificity and sensitivity of the detection.
While designing the primer, do not select the
primer region where the mutations are present in consensus region, particularly at 3¢ end
of primer.
Design the primer with GC content of
50–60 %, Tm difference within 4–5 °C, no
repetition of single bases and 3¢ ends with G,
C, GC, or CG.
After the addition of all reagents, mix them
properly with microtips. Mild vortexing and
centrifugation are necessary to remove the
air bubbles. If present, they can hinder the
amplification of the target DNA.
DNA isolated should be pure, and prior to
amplification, its quantity should be accessed.
If the DNA quantity is high, it should be
diluted in nuclease-free water.
A positive reaction of internal control reduces
the possibility of false-positive PCR
amplification and, thus, enhances the
specificity of the PCR reaction.
Agarose should be melted until bubbles come
from the bottom of the flask ends.
Gloves should be worn throughout these procedures, and particular care should be taken
while handling EtBr (consult safety data
sheets). It is a suspected mutagen.
Electrophoresis of agarose gel should usually be done at 8–10 V/cm. The gel should be
run for 1.5 h for proper separation of marker
and amplicons.
499
Acknowledgements M. M. Ahmad is thankful to receive
RFSMS grant-aided support from UGC, Government of
India, India.
References
1. Ahmad MM, Ahmad M, Ali A, Hamid R, Javed S and
Abdin MZ (2010) Molecular methods for detecting
pathogenic fungi. In: Gupta VK, Tuohy M and Gaur
RK (eds) Fungal Biochemistry and Biotechnology,
Germany: Lap Lambert Academic Publishing AG &
Co. KG. pp. 154–169
2. Konietzny U, Greiner R (2003) The application of PCR
in the detection of mycotoxigenic fungi in foods. Braz
J Microbiol 34:283–300
3. Yong RK, Cousin MA (2001) Detection of moulds producing aflatoxins in maize and peanuts by an immunoassay. Int J Food Microbiol 65:27–38
4. Abdin MZ, Ahmad MM, Javed S (2010) Advances in
molecular detection of Aspergillus: an update. Arch
Microbiol 192:409–425
5. Mauchline T, Kerry BR, Hirsch PR (2002)
Quantification in soil and the rhizosphere of the nematophagous fungus Verticillium chlamydosporium by
competitive PCR and comparison with selective plating. Appl Environ Microbiol 68:1846–1853
6. Fraaije BA, Lovell DJ, Rohel EA, Hollomon DW
(1999) Rapid detection and diagnosis of Septoria
tritici epidemics in wheat using a polymerase chain
reaction/Pico green assay. J Appl Microbiol 86:
701–708
7. Carbone I, Kohn LM (1999) A method for designing
primer sets for speciation studies in filamentous ascomycetes. Mycologia 91:553–556
8. Foster SJ, Singh G, Fitt BDL, Ashby AM (1999)
Development of PCR based diagnostic techniques
for the two mating types of Pyrenopeziza brassicae (light leaf spot) on winter oilseed rape (Brassica
napus ssp. oleifera). Physiol Mol Plant Pathol 55:
111–119
9. Fountaine JM, Shaw MW, Napier B, Ward E, Fraaije
BA (2007) Application of real-time and multiplex PCR
assays to study leaf blotch epidemics in barley.
Phytopathology 97:297–303
Protocols for the Quantification
of dsDNA and Its Fragmentation
Status in Fungi
48
Ioannis Papapostolou, Konstantinos Grintzalis,
and Christos Georgiou
Abstract
The presented protocols describe methodologies for the accurate
quantification of dsDNA concentration and fragmentation status of fungal
DNA (and that of any organism). The protocols can be used to quantify
dsDNA concentration, even up to picogram level, using the PicoGreen
fluorescent dye, and to discriminate the fragmentation status of a DNA
sample as fragmented (0–23 kb) and intact (> 23 kb) or as small size
(0–1 kb) fragmented DNA, using DNA samples as low as 2.5 mg mL-1.
Keywords
dsDNA quantification • DNA fragmentation • Small size (0–1 kb)
fragmented DNA • PEG precipitation • Fluorescence • PicoGreen • Fungi
Introduction
In many biological applications it is necessary to
accurately quantify both dsDNA concentration as
well as its fragmentation status. DNA damage
originating from oxidative attack of reactive oxygen species to DNA [1] may be non-repairable
(e.g., fragmentation) or repairable (e.g., nicks,
sugar/base modifications). Traditionally, fragmented DNA damage was estimated qualitatively
by the Comet assay [2] and by the DNA smearing
I. Papapostolou • K. Grintzalis • C. Georgiou (*)
Department of Biology, University of Patras,
University Campus, Patras 25600, Greece
e-mail: c.georgiou@upatras.gr
observed in an electrophoresis agarose gel [3],
visualized by ethidium bromide staining in both
methods. Other approaches frequently used are
the semi-qualitative micronuclei detection assay
that estimates the % of micronuclei [4], and the
DNA non-specific diphenylamine assay.
Furthermore, other protocols use fluorescent dyes
(e.g., ethidium bromide) to measure DNA concentration without taking into account the
underestimation of DNA concentration due to
the fragmentation degree of the DNA (i.e., the
fluorescence of dsDNA is decreased even up to
70 % when it is fragmented £ 23 kb) [5]. An additional disadvantage of the traditionally used dsDNA
fragmentation protocols is that they do not offer
any specific discrimination according to the size
of the fragmented segments. The present protocols resolve these problems by quantifying accurately any dsDNA sample and by discriminating
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_48, © Springer Science+Business Media, LLC 2013
501
502
its fragmentation status as fragmented (0–23 kb)
and intact (> 23 kb) dsDNA fraction (replacing
the DNA agarose electrophoresis gel method)
and as the small size (0–1 kb) fragmented dsDNA
(replacing the Comet assay) after its fractionation
by polyethylene glycol (PEG), the later being an
index for both necrotic and apoptotic dsDNA
damage [6, 7].
Materials
1. Balance (Kern, 770/65/6 J)
2. Centrifugal vacuum concentrator (Savant,
model SPD111V), connected to a vacuum
pump (KNF, N 820.3 FT.18)
3. Centrifuge tubes, 15 mL (ISC BioExpress,
catalog no. C-3394-1)
4. DNA genomic, from calf thymus, unsheared,
(Sigma-Aldrich, catalog no. D-4764)
5. DNase I from bovine pancreas, 580 units/mg
solid (Sigma-Aldrich, catalog no. DN-25)
6. Dimethyl sulfoxide, DMSO, anhydrous
(Sigma-Aldrich, catalog no. 276855).
Caution: Harmful
7. Ethylenediaminetetraacetic acid, Na2EDTA
(Merck, catalog no. 34033918). Caution:
Irritant, dangerous for the environment
8. Glass tubes, 15 mL
9. Hydrochloric acid, HCl, ³ 37 % (Fluka, catalog no. 84415). Caution: Corrosive
10. Manganese chloride (Merck, catalog no.
5833.0250)
11. Micropipettes (adjustable volume pipettes),
2.5 mL, 10 mL, 20 mL, 100 mL, 200 mL, 1 mL,
and tips (Eppendorf Research)
12. Microcuvette for fluorescence measurements, quartz (SOG/Q) 45 × 4 × 4 mm
(0.5 mL) with its FCA4 adaptor (Starna,
England)
13. NucleoSpin® Extract II kit (by MachereyNagel, Duren, Germany)
14. Sodium phosphate, Na2HPO4 (Merck, catalog no. 30412)
15. Sonicator, model Dr. Hielscher UP-50 H,
equipped with a 2-mm-diameter MS2 microtip (Dr. Hielscher GmbH, Teltow, Germany)
16. pH meter (Metrohm, 827 pHlab)
I. Papapostolou et al.
17. Polyethylene glycol 6000, PEG-6000 or PEG
(Serva, catalog no. 33137)
18. Refrigerated microcentrifuge (Eppendorf,
model 5417R)
19. Sodium chloride, NaCl (Merck, catalog no.
567440)
20. Sodium hydroxide, NaOH (Merck, catalog
no. 567530). Caution: Corrosive
21. Spectrofluorometer
(Shimadzu,
model
RF-1501)
22. Quant-iT™ PicoGreen (Invitrogen Molecular
Probes, catalog no. P7581). Caution: Very
toxic
23. Tris–HCl (Merck, catalog no. 648313)
24. Water, ddH2O, purified by a Milli-Q system
(Millipore Corp)
Methods
Solutions and Standard Curves
For applying the presented protocols, the following stock solutions and a standard curve need to
be performed.
Tris-EDTA (TE) Buffer
Prepare 100 mL TE buffer (10 mM Tris–HCl and
1 mM EDTA, pH 7.5), by dissolving 0.121 g
Tris–HCl and 0.037 g EDTA in ddH2O, and adjust
pH to 8.
Fragmented DNA Stock Solution
Dissolve without stirring overnight at 4 °C 1 mg
unsheared calf thymus genomic DNA in 4 mL TE
buffer. Place 0.5 mL of the 0.25 mg mL-1 calf thymus DNA standard solution in a 1.5-mL Eppendorf
tube, and sonicate it for 15 s at 350 W cm-2.
PicoGreen Stock Solution
Prepare fresh by mixing 10 mL of the commercial
reagent (made in 100 % anhydrous DMSO) with
0.99 mL TE buffer. This solution is sufficient for
15 tests.
PEG-NaCl Stock Solution
Prepare fresh 20 mL, by weighing accurately
2.822 g PEG and dissolving it in 12.344 mL ddH2O.
48
Protocols for the Quantification of dsDNA and Its Fragmentation Status in Fungi
Then, add to it 1.47 mL 0.2 M phosphate buffer,
pH 7.0, and 4.041 mL 4 M NaCl made in 10 mM
phosphate buffer, pH 7.0. The resulting solution
is somewhat viscous.
Construction of Totally Fragmented dsDNA
Standard Curve
Prepare a series of final volume 0.225 mL TE
solutions containing different concentrations of
fragmented dsDNA (up to 200 pg mL-1) and to
each of them add 75 mL PicoGreen stock solution. Measure the fluorescence units (F.U.) of the
above DNA solutions against a reagent blank
(without DNA) at ex/em 480/530 nm (in a 0.5 mL
quartz microcuvette) with a spectrofluorometer
(e.g., a Shimadzu RF-1501 spectrofluorometer,
set at 10 nm excitation/emission slit width and at
high sensitivity).
Fungal dsDNA Isolation
The dsDNA samples to be analyzed by the presented protocols can be isolated from any type of
fungi by the appropriate standard methods and
commercial kits.
Note: Artificial fragmentation of the isolated
dsDNA during its isolation should be avoided.
Note: Any buffers and equipment used for dsDNA
isolation should be sterilized.
Protocol for dsDNA Concentration
Quantification
The concentration of the isolated fungal dsDNA
sample is estimated after its complete fragmentation (via sonication) from the fragmented dsDNA
standard curve, and is performed as follows:
1. Dilute the previously isolated dsDNA sample
and sonicate it (as in the section Fragmented
dsDNA Stock Solution).
2. Measure the F.U. of the sonicated dsDNA
sample (as in the section Construction of
Totally Fragmented dsDNA Standard Curve)
and convert its F.U. to dsDNA concentration
using the totally fragmented dsDNA standard
curve.
503
Protocol for the Discrimination
of dsDNA as Fragmented and Intact
This protocol is used to quantify the fragmented
(0–23 kb) and intact (> 23 kb) dsDNA fractions in
any sample. There is a precaution that should be
taken into account, since the discrimination by
fluorescence is limited to ³ 23 kb, the analyzed
DNA sample should be above 23 kb in its native
condition (given the fact that DNA can also exist in
some organisms in sizes < 23 kb) [8, 9]. Regarding
the sensitivity limit for discrimination of dsDNA
fragmentation status, this protocol requires at least
dsDNA concentration of 135 pg mL-1. The protocol
is performed as follows:
1. Initially, the dsDNA sample is diluted appropriately and divided in two 0.3-mL portions.
2. One 0.3-mL portion, designated as “fragmented,” is sonicated (as stated in the section
Fragmented dsDNA Stock Solution) and the
F.U. (using 0.225 mL of it and 75 mL PicoGreen
stock solution) is measured (F.U.sonicated), as
stated in the section Construction of Totally
Fragmented DNA Standard Curve. From this
F.U.sonicated the concentration of the DNA sample is estimated using the totally fragmented
DNA standard curve.
3. The fluorescence of the other 0.3-mL portion
(using 0.225 mL of it and 75 mL PicoGreen
stock solution), designated as “unsonicated,”
is measured (F.U.unsonicated, i.e., without subjecting it to sonication).
4. By taking into account any appropriate dilutions, the above F.U. measurements are compared and the percentage of fragmented/intact
DNA is expressed as follows:
a. If F.U.sonicated and F.U.unsonicated are equal (with
S.E. ±3 %), the dsDNA sample is totally
fragmented.
b. If F.U.sonicated and F.U.unsonicated differ by
67–73 %, the dsDNA sample is composed
only of intact DNA (as previously mentioned, the difference in F.U. of a DNA
sample if it is totally intact and totally fragmented is 70 %).
c. If F.U.sonicated and F.U.unsonicated differ below
67 %, the dsDNA sample is partially
fragmented, and exists as a mixture of
504
I. Papapostolou et al.
fragmented and intact fractions, which are
quantified by the following mathematical
treatment of F.U.sonicated and F.U.unsonicated:
[5, 7].
i. To estimate the F.U. the dsDNA sample
would have if it were intact (F.U.intact),
multiply the F.U.sonicated value by 3.333
(=100/30 %) and set the resulting value
as 100 %. Then use the resulting
value in order to convert the F.U.unsonicated
value to %, designated as y %
(=100 % F.U.unsonicated/F.U.intact).
ii. Enter this y % value in the equation
x % = (100 % – y %)/0.7, which calculates the percentage of the fragmented
DNA fraction expressed as %x of the
total dsDNA concentration (which is
determined using the F.U.sonicated in step
1). The concentration of the intact
dsDNA fraction, expressed as 100-x %,
can be determined accordingly.
Protocol for the Isolation and
Quantification of Small Size (0–1 kb)
Fragmented dsDNA
This protocol quantifies the small-size (0–1 kb)
fragmented DNA after its isolation by PEG precipitation which is very important as an index of
necrotic or apoptotic events. The protocol can be
applied to any previously isolated DNA sample
and is performed as follows:
1. Initially, the concentration of the isolated fungal dsDNA sample is determined as in
“Protocol
for
dsDNA
concentration
quantification.” For applying this protocol the
limitation in dsDNA concentration of the sample for accurate PEG precipitation is 0.2 mL
2.5 m g mL-1 [6].
2. In a 1.5-mL Eppendorf tube add 0.16 mL of the
isolated dsDNA sample and 0.34 mL PEGNaCl stock solution. Mix gently and incubate
the resulting DNA-PEG-NaCl mixture for minimum 12 h (or overnight) in ice-water bath.
Note: Accurate pipetting is required due to the
small volumes used in this step.
3. Centrifuge in a refrigerated microcentrifuge
(4 ºC) at 15,000 g for 5 min and collect the
supernatant in a new Eppendorf tube. The
supernatant contains the 0–1 kb fragmented
DNA fraction, and the pellet the intact plus the
> 1 kb fragmented dsDNA.
4. For measuring small-size fragmented dsDNA
concentration in the PEG supernatant in step 3
by fluorescence, the interfering PEG is
removed and the dilute small fragment dsDNA
is concentrated as follows. Dilute the supernatant 2 × with TE buffer and concentrate the
0–1 kb fragmented dsDNA with the Nucleospin
Extract II kit (following manufacturer’s
instructions) in 0.05 mL of the kit’s elution
buffer.
5. Dilute the eluate appropriately and determine
the dsDNA concentration of the isolated
small-size (0–1 kb) fragmented DNA as in the
section Protocol for dsDNA Concentration
Quantification.
References
1. Halliwell B, Gutteridge CMJ (1999) Free radicals in
biology and medicine, 3rd edn. Oxford University
Press, Oxford, UK
2. Olive PL, Banath JP (2006) The comet assay: a method
to measure DNA damage in individual cells. Nat Protoc
1:23–29
3. Sambrook J, Fritsch FE, Maniatis T (1989)
Molecular cloning: a laboratory manual. Cold
Spring Harbor Laboratory Press, Cold Spring Harbor,
New York
4. Fenech M (2000) The in vitro micronucleus technique.
Mutat Res 455:81–95
5. Georgiou DC, Papapostolou I (2006) Assay for the
quantification of intact/fragmented genomic DNA.
Anal Biochem 358:247–256
6. Georgiou DC, Patsoukis N, Papapostolou I (2005)
Assay for the quantification of small-sized fragmented
genomic DNA. Anal Biochem 339:223–230
7. Georgiou CD, Papapostolou I, Grintzalis K (2009)
Protocol for the quantitative assessment of DNA concentration and damage (fragmentation and nicks). Nat
Protoc 4:125–131
8. Brown TA (2001) Gene cloning and DNA analysis: an
introduction. Blackwell Publishing, Malden, MA
9. Griffiths AJF, Miller JH, Suzuki DT, Lewontin RC,
Gelbart WM (2000) An introduction to genetic analysis. W. H. Freeman, New York, NY
Rapid Identification and Detection
of Pathogenic Fungi by Padlock
Probes
49
Clement K.M. Tsui, Bin Wang, Cor D. Schoen,
and Richard C. Hamelin
Abstract
Fungi are important pathogens of human diseases, as well as to agricultural
crop and trees. Molecular diagnostics can detect diseases early, and
improve identification accuracy and follow-up disease management. The
use of padlock probe is effective to facilitate these detections and pathogen
identification quickly and accurately. In this chapter we describe three
diagnostic assays that utilize padlock probes in combination with various
technologies for the detection of pathogenic fungi.
Keywords
Molecular diagnostics • Padlock probes • Pathogenic fungi • Single nucleotide polymorphisms • Rolling circle amplification
Introduction
C.K.M. Tsui (*)
Department of Forest Sciences, The University
of British Columbia, 2424 Main Mall, Vancouver,
BC, Canada V6T 1Z4
e-mail: clementsui@gmail.com
B. Wang
Centre of Virus Research, Westmead Millennium
Institute, University of Sydney, Westmead Hospital,
Darcy Road, Westmead, NSW 2145, Australia
C.D. Schoen
Department of Bio-Interactions and Plant Health,
Plant Research International B. V, Droevendaalsesteeg 1,
Wageningen 6708PB, The Netherlands
R.C. Hamelin
Department of Forest Sciences, The University
of British Columbia, 2424 Main Mall, Vancouver, BC,
Canada V6T 1Z4
Natural Resources Canada, Laurentian Forestry Centre,
1055 rue du P.E.P.S, Quebec, QC, Canada G1V 4C7
Fungi represent the greatest eukaryotic diversity
on earth and they are the primary decomposers in
the ecosystems. Of the estimated 1.5 million
species, more than 400 species have been associated with human diseases and about 100 species
can cause infection to normal individuals [1].
Representatives of genera such as Candida
(Saccharomycetes, Ascomycota), Cryptococcus
(Tremellales, Basidiomycota) and Aspergillus
(Eurotiales, Ascomycota) are prevalent human
diseases that cause superficial, systematic and
lung infections [1]. Fungi also represent one of the
most important groups of pathogens to plants and
they cause significant economic losses to agriculture and forestry business [2, 3]. In addition, some
fungal pathogens can cause ecosystem-wide
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_49, © Springer Science+Business Media, LLC 2013
505
506
disturbances that can endanger other plant and tree
species. Fusarium (Hypocreales, Ascomycota),
Cronartium
(Pucciniales,
Basidiomycota),
Grosmannia (Ophiostomatales, Ascomycota),
and Phytophthora (Peronosporales, Oomycota
(fungus-like Stramenopiles)) are some examples
of genera that comprise serious pathogens of
crops and trees [2, 3].
Rapid and accurate detection and identification
of fungal pathogens at species and subspecies
level in the clinical setting and natural environment, or on plant materials are the keys to proper
patient treatment and disease/pathogen surveillance, containment and eradication [1, 4].
However, many fungal pathogens exist as species-complexes or they have very low abundance
in the clinical specimen and natural environment.
Different molecular types/varieties also exist
within species and they may have different pathogenic profiles and virulence level to the hosts.
These issues pose increasing difficulties to accurate pathogen genotyping/identification [1, 4].
Culture-based morphological features are the
most predominant approach used for fungal
pathogen identification. But the major drawback
of this approach is that many pathogenic species
do not produce fruiting bodies readily in culture
that can be useful for morphological identification
[5]. In addition, identification procedures are
time-consuming or laborious, or require taxonomical or specialized expertise in interpretation
[5]. Other approaches commonly used in clinical
pathogen identification are histopathological,
biochemical or serological information and
molecular analyses such as polymerase chain
reaction (PCR) fingerprinting, amplified fragment length polymorphism (AFLP) and restriction fragment length polymorphism (RFLP) [1].
Similarly, these techniques may be time-consuming and sensitive to contamination, which can
result in low accuracy and specificity.
Routine DNA sequencing of the internal transcribed spacer (ITS) and large subunit (LSU)
regions of rRNA gene followed by comparative
sequence analysis to resources in GenBank facilitate the design of novel molecular method(s)
for rapid species identification. This approach
C.K.M. Tsui et al.
includes the selection of genomic regions with
low sequence polymorphisms such as single
nucleotide polymorphisms (SNPs) because many
pathogenic molecular types or subspecies differ
by only a few base pairs in various regions of the
genome. Although detection and characterization
of SNPs is becoming increasing popular for
pathogen identification, it was considered as a
major challenge for conventional real-time qPCR
using regular oligos and probes targeting SNPs
detected by fluorescent dyes (e.g., SYBR green
or TaqMan probes). In order to recognize a single
SNP among different genotypes, techniques other
than conventional qPCR are required.
Padlock probes could be an alternative platform to the conventional approaches [6]. Padlock
probes comprise two target-complementary
sequence regions at both ends for hybridization
to specific DNA sequence, as well as a non
target-complementary segment. The sequence
regions at both ends of the probe are designed
that they are joined together after target hybridization and DNA ligation (Fig. 49.1). The helical
nature of double-stranded DNA (dsDNA) enables
the probe to topologically bind to the target
strand. The advantage of padlock probes is that
the sequences at both ends of the probe are sensitive to mismatches [7].
Padlock probes were initially introduced for
in situ DNA localization and detection [6]. They
were developed for the discrimination of centromeric sequence variation in human chromosomes
[8]. However, the method has been applied to
detection of genetically modified organisms [9].
In addition, this technology also has multiplexing
potential as the interaction between padlock
probes does not give rise to circular molecules,
which can be easily removed from the detection
system [10].
The sensitivity of a padlock probe is comparable to a conventional oligonucleotide probe, but
padlock probes have greater specificity with less
noise background [8]. To improve the detection
sensitivity, various technologies have been introduced to detect and amplify the signals from the
circularized probes targeting various fungal
pathogens (Table 49.1) [11, 18].
49
Rapid Identification and Detection of Pathogenic Fungi by Padlock Probes
Fig. 49.1 Overview of the padlock probe technology
coupled with hyperbranching rolling circle amplification
(H-RCA) method for detection of the single nucleotide
polymorphisms (SNPs). (a) Hybridization of padlock
probe to a target template with a mismatch at 3¢ end.
(b) Padlock probe matches perfectly to target template.
(c) High-fidelity DNA ligase is unable to ligate the mismatched template-probe complex. (d) In perfectly
matched template-probe complex, the 5¢ and 3¢ ends of
padlock probe can be joined, forming a circular molecule.
(e) Templates and linear probes are removed by exonu-
Materials
1.
2.
3.
4.
Vortexer.
Water bath.
Microcentrifuge.
Disposable polypropylene microcentrifuge
tubes (1.5-mL conical, 2-mL screw-capped).
507
cleolysis reaction (Exonuclease I and III). (f) Only the
circularized probe remains in the assay system. (g) H-RCA
is performed using two primers and Bst DNA polymerase;
however, no amplification will take place without a circular probe. (h) In the presence of a circular template, the
two primers generate a self-propagating, ever-increasing
pattern of alternating strand displacement, branching, and
DNA fragment release events. (I) Exponential accumulation of product during isothermal H-RCA of DNA
minicircles can be detected by real-time PCR instrument
with the addition of SYBR green
5. PCR strips (flat cap, 0.2-mL thin-walled).
6. PCR Thermal cycler (e.g., Applied
Biosystems GeneAmp® 9700).
7. Real-Time PCR Thermal cycler (e.g., Corbett
RotorGene 3000).
8. Sterile distilled water (PCR grade)/Sigma
water.
508
Table 49.1 Summary of padlock probe investigations in the detection of fungal pathogens
Type of samples
DNA from cultures
Gene target
ITS rRNA
Detection system
Hyperbranched rolling
circle amplification
DNA from cultures
ITS rRNA
Hyperbranched rolling
circle amplification
DNA from cultures
and clinical specimens
(sputum, blood)
DNA from cultures
ITS rRNA
Hyperbranched Rolling
circle amplification
Lanosterol 14a-demethylase
(ERG11p)—enzyme in
fungal ergosterol synethesis
ITS, LSU rRNA
Hyperbranched rolling
circle amplification
Candida albicans
Wang et al. [16]
Hyperbranched rolling
circle amplification
Real-time PCR using
Taqman probe
Grosmannia clavigera, Leptographium longiclavatum
Tsui et al. [3]
Phytophthora nicotianae, P cactorum, P. infestans,
P. sojae, Pythium ultimum (Oomycetes), Rhizotonia
solani, Fusarium oxysporum f. sp. radicis-lycopersici,
Myrothecium roridum, Myrothecium verrucaria,
Verticillium dahlia, Verticillium alboatrum
Phytophthora nicotianae, P cactorum, P. infestans,
P. sojae, Pythium ultimum (Oomycetes), Rhizotonia
solani, Fusarium oxysporum f. sp. radicis-lycopersici,
Myrothecium roridum, Myrothecium verrucaria,
Verticillium dahlia, Verticillium alboatrum
Phytophthora nicotianae, P cactorum, P. infestans,
P. sojae, Pythium ultimum (Oomycetes), Rhizotonia
solani, Fusarium oxysporum f. sp. radicis-lycopersici,
Myrothecium roridum, Myrothecium verrucaria,
Verticillium dahlia, Verticillium alboatrum
Szemes et al. [13]
DNA from cultures
and bark beetles
DNA from cultures
ITS rRNA
ITS rRNA
Real-time PCR on
OpenArrays™ system
DNA from cultures,
and the filters in which
horticultural re-circulation
water spiked with various
pathogens has passed
through
DNA from cultures and
clinical specimens
(urine, charcoal swabs,
sputum, vaginal swab
culture broth)
ITS rRNA
A desthiobiotin moiety
connected to the linker
sequence
ITS rRNA
Rolling circle amplification
and Luminex™
Aspergillus flavus, A. fumigatus, A. nidulans, A. niger,
Candida albicans, C. glabrata, C. tropicalis,
C. parasilopsis, Cryptococcus neoformans,
Pneumocystis jirovecii
Including Oomycetes, which are fungus-like organisms classified under Stramenopiles. Traditionally, they have been studied by fungal biologists
a
References
Kong et al. [14]
Zhou et al. [15]
Kaocharoan et al. [17]
van Doorn et al. [25]
van Doorn et al. [27]
Eriksson et al. [18]
C.K.M. Tsui et al.
DNA from cultures
Genera or species identifieda
Trichophyton rubrum, T. tonsurans, T. soudanense,
T. violaceum, T. mentagrophytes, Epidermophyton
floccosum, Microsporum canis, M. gypseum
Candida albicans, C. glabrata, C. krusei, C. tropicalis,
C, dubliniensis, C. guilliermondii, Aspergillus fumigatus,
A. flavatus, Scedosporium apiospermum, S. prolifercans
Cryptococcus neoformans var. grubii, Cryptococcus
neoformans var. neoformans, Cryptococcus gattii
49
Rapid Identification and Detection of Pathogenic Fungi by Padlock Probes
9. dNTPs, a mixture of dATP, dCTP, dGTP and
dTTP 400 mM each (Promega, Australia),
stored at −20 °C.
10. DMSO (Dimethyl sulfoxide)—used to minimize the secondary structure formed in DNA
and RNA (Sigma-Aldrich, Australia).
11. Thermostable DNA polymerase (e.g., Taq,
and reaction buffer supplied by manufacturer). We typically use Taq from MBI
Fermentas (York, UK), which is supplied
with a 10× reaction buffer of 100 mM Tris–
HCl pH 8.8 (25 °C), 500 mM KCl and 0.8 %
Nonidet P40. Separate stocks of BSA (20 mg/
mL) and MgCl2 (25 mM) are also supplied
for adding to the reaction. All of these
reagents are stored at −20 °C.
12. TBE buffer—50 mM Tris, 50 mM boric acid,
1 mM EDTA. Dilute when needed from a
10× stock.
13. Padlock probe (custom-made, SigmaAldrich); page purified, re-suspended to a
stock concentration of 100 mM and diluted to
1 mM working concentration.
14. Pfu DNA ligase with 10× reaction buffer
(Stratagene) 20 mM Tris–HCl (pH 7.5),
20 mM KCl, 10 mM MgCl2, 0.1 % Igepal,
0.01 mM rATP and 1 mM DTT. All of these
reagents are stored at −20 °C.
15. Exonuclease I and III with 10× reaction buffer 1 (New England Biolabs, Ipswich, MA,
USA), stored at −20 °C. (Note: Degraded
nucleic acids and nonspecific products due to
cross reactions from the 5¢ and/or 3¢ ends are
degraded in order to remove any unreacted
probes, while preserving reacted endless
probes.)
16. Bst DNA polymerase with 10× ThermoPol
Reaction Buffer (New England Biolabs,
Ipswich, MA, USA). All of these reagents
are stored at −20 °C.
17. Oligonucleotide primers targeting the linker
sequence (custom-made, Sigma-Aldrich),
re-suspended to a concentration of 10 mM.
18. SYBR Green I (Sigma-Aldrich, St Louis,
MO, USA), stored at −20 °C
19. A confocal ScanArray 4000 laser scanning system (Packard GSI Lumonics)
containing a GreNe 543-nm laser for PE and
20.
21.
22.
23.
24.
509
a HeNe 633-nm laser for Cy5 fluorescence
measurement in microarrays analysis.
OpenArrays® Real-Time PCR Instrument
(Applied Biosystems).
The ZipCodes were chosen from the GeneFlex
TagArray set (Affymetrix). Potential for secondary structures and ZipCode specificity
was examined with Visual OMP 6.0 software
(DNA Software Inc.).
The cZipCode oligonucleotides carrying a
C12 linker and a 5¢ NH2 group were synthesized and spotted onto Nexterion MPX-E16
epoxy-coated slides by Isogen B.V. according to the manufacturer’s instructions (Schott
Nexterion).
Enzymes EcoRI, HindIII and BamHI (New
England Biolabs Inc., Ipswich, USA).
MyOne™ Streptavidin C1 Dynabeads®
(Dynal Biotech ASA, Oslo, Norway).
Design of Padlock Probes
1. To select a gene with enough resolution as
target for the padlock probe design.
2. To improve the efficiency of hybridization, the
padlock probes should be designed with minimum secondary structure, and the melting
temperature (Tm) of 5¢ end probe complementary sequence will be close to or above ligation temperature (60 °C).
3. To increase 3¢ end complementary sequence
specificity, Tm will be optimized with 10–15 °C
below the ligation temperature.
4. The genetic linker region will be designed/
created to minimize its similarity to any known
species or sequences by performing BlastN
search.
Differentiation of Fungal Pathogens
by Padlock Probe and Rolling Circle
Amplification
Rolling circle amplification (RCA) is based on
the rolling replication of short single-stranded
DNA (ssDNA) circular molecules [12, 19]. RCA
involves a single forward primer complementary
510
to the linker region of the padlock probe and
a DNA polymerase with strand displacement
activity in an isothermal condition [20]. As a
result, the padlock probe signal can be amplified
several thousand-fold because the polymerase
extends the bound primer along the padlock
probes for many cycles and displaces upstream
sequences, producing a long ssDNA molecule
comprising multiple repeats of the probe sequence.
While two primers; a first forward primer that
binds to the padlock probe and initializes RCA,
and a second primer that targets the repeated
ssDNA sequence of the primary RCA product,
can generate large numbers of copies of the DNA
fragments. This is called hyperbranching RCA
(H-RCA) (see Fig. 49.1) [12].
Padlock probe coupled with H-RCA offers a
significant advantage for the detection of SNPs.
The formation of circular probes via ligation
happens when both ends of the padlock probes
perfectly hybridize to the target at juxtaposition.
The subsequent H-RCA amplification of target
probe can be carried out when circularized
probes become available. These two strict conditions create an ideal detection platform for highly
sensitive and specific SNPs detection. By increasing the hybridization temperature and shortening
the 3¢ complementary sequence (below the reaction temperature), the discrimination of SNP can
be further improved [20, 21]. This method for
SNPs detection has been developed for various
groups of pathogenic microorganisms [3, 14–17,
22–24].
Below we describe the general procedures to
perform assays using padlock probes coupled
with H-RCA amplification of the probe signal.
Modifications would be required to adjust for differences in probe annealing temperature. The
size of tubes used for genotyping may be varied,
depending on the type of real-time PCR
machines.
1. Mix 1011 copies purified PCR amplicons,
1 pmol of padlock probe (Sigma-Aldrich), 2 U
of Pfu DNA ligase (Stratagene) and 1 mL 10×
reaction buffer with a total reaction volume of
10 mL in a 0.2 mL thin-walled PCR tube (see
Note 1).
2. Centrifuge briefly and place the tube in a
thermal cycler with a heated lid.
C.K.M. Tsui et al.
3. The ligation reaction included one cycle of
5 min at 94 °C to denature the target template
followed by 5 cycles of 94 °C for 30 s and
4 min ligation at 65 °C.
4. After the ligation reaction, quickly transfer the
tube to 4 °C.
5. Add 10 U each of Exonuclease I and III (New
England Biolab, Ipswich, MA, USA), 2 mL of
10× reaction buffer 1 and sterile distilled water
to PCR tube containing ligation mixture to
make up the final volume to 20 mL.
6. Incubate the PCR tube at 37 °C for 30 min in
thermal cycler with heated lid followed by
94 °C for 3 min to inactivate Exonuclease.
7. Prepare H-RCA master mixture by adding 8 U of
Bst DNA polymerase (New England Biolabs,
Ipswich, MA, USA), 5 mL reaction buffer,
400 mM dNTP mix, 10 pmol of each of two
H-RCA primers [P1: 5¢ ATGGGCACCG
AAGAAGCA3¢;P2:5¢CGCGCAGACACGATA
3¢], 5 % of DMSO (v/v) and 1× SYBR green I to
a final volume of 30 mL.
8. Add 30 mL of H-RCA master mixture to the
PCR tube; incubate at 65 °C for 30 min in
Corbett RotorGene 3000 Real-Time thermal
cycler. Collect florescence signal every minute
for up to 30 min.
Quantitative Multiplex Detection
by the Padlock Lock System Coupled
with Universal, High-Throughput
Real-Time PCR on OpenArray®
Instrument
This assay enables specific, high-throughput,
quantitative detection of multiple pathogens over
a wide range of target concentrations. The ligation padlock probes (also known as PRI-lock
probes) are long oligonucleotides with target
complementary regions at their 5¢ and 3¢ ends
[25]. Upon target hybridization, the PRI-lock
probes are circularized via enzymatic ligation,
subsequently serving as templates for individual amplification via unique probe-specific
primers. Adaptation to the OpenArray, which
can accommodate up to 3,072 ×33 nL PCR
amplifications, allowed high-throughput realtime quantification (Fig. 49.2) [26].
49
Rapid Identification and Detection of Pathogenic Fungi by Padlock Probes
511
Fig. 49.2 (a) OpenArray™ architecture. The OpenArray™
has 48 subarrays and each subarray contains 64 microscopic through-holes of 33 nl volume. The primers are preloaded into the holes. The sample combined with the
reaction mix is auto-loaded due to the surface tension, provided by the hydrophilic coating of the holes and the
hydrophobic surface of the array. (b) PRI-lock probe
design. T1a and T1b indicate target complementary
regions. Unique primer sites ensure specific amplification
(forward: F1 and reverse: R1) and each PRI-lock contains
a universal sequence (US) and a desthiobiotin moiety
(dBio). (c) Multiple target specific PRI-lock probes are
ligated on fragmented DNA samples. T1a and T1b bind to
adjacent sequences of the target and in case of a perfect
match, the probe is circularized by a ligase. The probes are
captured via the desthiobiotin moiety using magnetic
streptavidin-coated beads. The PRI-lock probes are washed
and quantitatively eluted from the beads. Unreacted probes
are removed by exonuclease treatment. (d) Circularized
probes are loaded and independently amplified on the
Biotrove OpenArray™ platform using PRI-lock probe
specific primers. The amplification is monitored using
SYBR-Green and the ligated PRI-lock probes are quantified
based on the threshold cycle number (CT)
The following protocol uses the PRI-lock detection system targeting plant pathogens at different
taxonomic levels. The nucleic acid targets can be
reliably quantified over 5 orders of magnitude
with a dynamic detection range of more than 104
copies/mL. Pathogen quantification is equally
robust in single target versus mixed target assays.
1. Genomic DNA (i.e., 500 pg) (Note 2) is fragmented by digestion using EcoRI, HindIII
and BamHI (New England Biolabs Inc.,
Ipswich, MA, USA) for 15 min at 37 °C.
2. Add 250 pM of the individual PRI-lock
probes (Seraing, Belgium) [Note 3], 20 mM
Tris–HCl, pH 9.0, 25 mM KCH3COO,
10 mM Mg(CH3COO)2, 1 mM NAD, 10 mM
DTT, 0.1 % Triton X-100, 20 ng sonicated
salm sperm DNA and 20 U Taq ligase (New
England Biolabs Inc., Ipswich, MA, USA)
to the fragmented DNA sample to an end
volume of 10 mL (Note 4).
3. Place the tubes in the PCR machine and run
the appropriate ligation cycle programme.
Samples are denatured at 95 °C for 5 min and
subsequently subjected to 20 cycles of 30 s at
95 °C and 5 min at 65 °C, followed by
enzyme inactivation at 95 °C for 15 min.
512
4. When the ligation is complete 30 mL distilled
water is added to each reaction.
5. The desthiobiotin moiety of the PRI-lock
probes is captured through the addition of
40 mL solution containing 2 M NaCl, 10 mM
Tris–HCl, pH 7.5, 1 mM EDTA, 0.2 M NaOH
and 200 mg magnetic MyOne™ Streptavidin
C1 Dynabeads (Dynal Biotech ASA, Oslo,
Norway).
6. Rotate at 4 °C for 1 h.
7. Centrifuge the samples at 2,000×g for 10 s,
and collect the Dynabeads via application of
a magnetic field.
8. The Dynabeads are washed with 100 mL
100 mM Tris–HCl, pH 7.5 and 50 mM NaCl.
9. The Dynabeads are re-suspended in 10 mL distilled water and incubated at 95 °C for 10 min,
allowing quantitative elution of the PRI-lock
probes from the Dynabeads (Note 5).
10. Samples are transferred onto ice and the
empty magnetic streptavidin beads removed
via application of a magnetic field, leaving
the washed PRI-lock probes in the solution.
11. 10 mL of Exonuclease mixture (10 mM Tris–
HCl, pH 9.0, 4.4 mM MgCl2, 0.1 mg/mL
BSA, 0.5 U Exonuclease I (USB Europe
GmbH, Staufen, Germany) and 0.5 U
Exonuclease III (USB Europe GmbH,
Staufen, Germany)) is added to each reaction, and incubated at 37 °C for 30 min.
12. When exonuclease step is complete the
enzyme is inactivated at 95 °C for 2.5 h.
13. Amplification of ligated PRI-lock probes is
followed in real-time using an OpenArray
Real-Time PCR Instrument (Applied
Biosystems, Foster City, USA) (Note 5).
Each subarray is loaded with 5.0 mL mastermix containing 2.5 mL ligated padlock mixture and reagents in a final concentration of
1× LightCycler® FastStart DNA Master
SYBR Green I mix (Roche Diagnostics
GmbH, Mannheim, Germany), 0.2 %
Pluronic F-68 (Gibco, Carlsbad, USA), 1 mg/
mL BSA (Sigma-Aldrich, St Louis, USA),
1:4000 SYBR Green I (Sigma-Aldrich),
0.5 % (v/v) Glycerol (Sigma-Aldrich), 8 %
(v/v) deionized formamide (Sigma-Aldrich)
and 1.0 pg PCR control template (Note 6).
C.K.M. Tsui et al.
14. The PCR OpenArray thermal cycling protocol is fixed and consist a 90 °C step for
10 min, followed by 27 cycles of 28 s at
95 °C, 1 min at 55 °C and 70 s at 72 °C
(imaging step).
15. The OpenArray Real-time PCR Software is
used for data analyses. The positive
amplification reactions are analyzed for
amplicon specificity by studying the individual melting curves (Note 70).
Detection and Identification
of Multiple Microorganisms
in Environmental Samples by a
Cleavable Padlock Probe-Based
Ligation Detection Assay
The ligation detection (LD) system uses a single
compound detector probe per target. The padlock
probes contain asymmetric target complementary regions at both their 5¢ and 3¢ ends that confer specific target detection and have both a
desthiobiotin moiety and an internal endonuclease IV cleavage site [27]. PCR amplification
of universal phylogenetic target genes (e.g., 16S,
18S, and 23S rRNA genes), a number of
microorganism-specific genetic markers [28–30],
or random amplification of genomic DNA
(gDNA) fragments [31] serve as potential targets
for PRI-lock probe ligation. Upon target hybridization, the PRI-lock probes are circularized via
enzymatic ligation, captured, and cleaved, allowing only the originally ligated padlock probes to
be visualized on a universal microarray
(Fig. 49.3) (Note 8). Unlike previous procedures,
the probes themselves are not amplified, thereby
allowing a simple padlock cleavage to yield a
background-free assay.
The following LD protocol enables specific
detection and identification of multiple pathogens
over a wide range of target concentrations and is
adaptable to a variety of applications.
1. Targets for ligation are generated by PCR
preamplification of 10 ng extracted gDNA
(Notes 2 and 9). Preamplification is performed in a 2× Master Mix (Applied
Biosystems) containing AmpliTaq Gold
49
Rapid Identification and Detection of Pathogenic Fungi by Padlock Probes
513
Fig. 49.3 (a) PLP design. T1a and T1b are asymmetric
target complementary regions. Each PLP contains a unique
ZipCode sequence for universal array hybridization, two
spacer sequences (S1 and S2), a desthiobiotin moiety
(dBio) for probe capture, a polyoligo(dT) linker sequence,
and a polydeoxyuracil sequence for probe cleavage. (b)
Multiple target-specific PLPs are ligated to PCRpreamplified DNA samples. T1a and T1b bind to adjacent
sequences of the target, and in the case of a perfect match,
the probe is circularized by enzymatic ligation. The PLPs
are reversibly captured and washed via the desthiobiotin
moiety with magnetic streptavidin-coated beads. Next, the
washed probes are cleaved at the polydeoxyuracil
sequences with UNG and endonuclease enzymes. The
sample containing the cleaved PLPs is hybridized on a universal microarray. Finally, only the hybridized PLPs that
were originally ligated can be labeled and visualized with
streptavidin R-PE by using the desthiobiotin moiety
DNA polymerase, deoxynucleoside triphosphates with dUTP, 0.12 mL UNG (Applied
Biosystems), 10 ng gDNA extract and
300 nM of each primer (Note 11). The reaction mixture is incubated at 50 °C for 2 min,
followed by 10 min denaturation at 95 °C
and 40 cycles consisting of a 30 s incubation
at 95 °C, an annealing step at 60 °C for 30 s
(ITS), and an elongation step at 72 °C for
60 s (Note 9). After the last cycle, the reaction is immediately cooled to 4 °C.
2. The total PCR-amplified products are fragmented by digestion with EcoRI, HindIII,
and BamHI (New England BioLabs, Inc.) for
15 min at 37 °C.
3. Add 5 mL of the fragmented DNA sample to
20 pM of the individual LD padlock probes,
20 mM Tris–HCl (pH 7.5), 20 mM KCl,
10 mM MgCl2, 0.1 % Igepal, 0.01 mM rATP,
1 mM dithiothreitol, 20 ng sonicated
salmon sperm DNA, and 4 U Pfu DNA
ligase (Stratagene) with an end volume of
10 mL.
4. Place the tubes in the PCR machine and run
the appropriate ligation cycle programme.
Samples are denatured at 95 °C for 5 min
and subsequently subjected to 20 cycles
of 30 s at 95 °C and 5 min at 65 °C, followed by enzyme inactivation at 95 °C for
15 min.
514
5. When the ligation is complete 30 mL distilled
water is added to each reaction.
6. The desthiobiotin moiety of the LD probes is
captured through the addition of 40 mL containing 2 M NaCl, 10 mM Tris–HCl, pH 7.5,
1 mM EDTA, 0.2 M NaOH, and 200 mg magnetic MyOne™ Streptavidin C1 Dynabeads
(Dynal Biotech ASA, Oslo, Norway).
7. Rotate at 4 °C for 1 h.
8. Centrifuge the samples at 2,000 × g for 10 s,
and collect the Dynabeads via application of
a magnetic field.
9. Wash the Dynabeads with 100 mL 100 mM
Tris–HCl, pH 7.5, and 50 mM NaCl.
10. Re-suspended the Dynabeads in 10 mL distilled water and incubated at 95 °C for
10 min, allowing quantitative elution of the
LD probes from the Dynabeads [32].
11. Transfer the samples onto ice and remove the
empty magnetic streptavidin beads via application of a magnetic field, leaving the washed
LD probes in the solution.
12. 10 mL of cleavage mixture (10 U uracilN-glycosylase (UNG; Applied Biosystems),
10 U endonuclease IV (New England
BioLabs), 2× NEBuffer 3 (New England
BioLabs), 2× bovine serum albumin) is
added to each reaction mixture and incubated
at 37 °C for 1.5 h.
13. 2 ml of 1.1 M NaOH is added and incubated
at 95 °C for 10 min. Finally, 8 mL of 0.5 M
Tris buffer (pH 6.8) is added to the samples
for neutralizing the solution.
14. The cleaved ligation samples are heated for
10 min at 95 °C and then cooled rapidly on
ice (Note 12).
15. 40 mL of the array hybridization mixture is
added to each well (Note 10); the chambers
are sealed and the arrays are hybridized at
55 °C overnight under high humidity.
16. The array is washed once at 55 °C for 5 min
in preheated 1× SSC (0.15 M NaCl plus
0.015 M sodium citrate), 0.1 % sodium dodecyl sulfate and twice for an additional 1 min
at room temperature in 0.1× SSC-0.1 %
sodium dodecyl sulfate and in TNT (0.1 M
Tris–HCl (pH 7.6), 0.15 M NaCl, 0.05 %
Tween 20), respectively.
C.K.M. Tsui et al.
17. The array is incubated in blocking solution
(0.1 M Tris–HCl (pH 7.5), 0.15 M NaCl,
0.5 % blocking reagent (Perkin-Elmer)) for
10 min at high humidity at room temperature
and washed for 1 min in TNT.
18. 20 ml of staining solution (15 mg/mL streptavidin R-phycoerythrin (PE; Qiagen) in 20 mL
of blocking solution) is added to each well.
19. The array is incubated in the dark at room
temperature for 15 min. The silicon structures
are removed from the slides, washed three
times for 1 min in TNT and twice for 1 min in
0.1× SSC, respectively. Finally, the slides are
dried by spinning at 250 × g for 3 min.
20. Microarrays are analyzed with a confocal
ScanArray 4000 laser scanning system
(Packard GSI Lumonics) containing a GreNe
543-nm laser for PE and a HeNe 633-nm
laser for Cy5 fluorescence measurement
(Note 13).
Notes
1. Gloves should be worn throughout these
procedures.
2. Genomic DNAs from all microorganisms are
isolated using the Puregene Genomic DNA
isolation kit (Gentra/Biozym, Landgraaf, the
Netherlands) according to the manufacturer’s
instructions; other methods to prepare gDNA
from organisms can be used as well.
3. The padlock probe target complementary
regions are engineered according to previously described design criteria [13] and are
connected by a 60 bp compound linker
sequence. The linker sequence contains a
20 bp, generic sequence and two, unique
primer binding sites for specific PCR
amplification. All primer pairs have equal
melting temperatures to allow universal
SYBR-Green-based detection in real-time
PCR. The primer pairs are chosen from
GeneFlex™ TagArray set (Affymetrix) in a
way to minimize padlock probe secondary
structures and optimize both, primer TM and
primer specificity. Potential for secondary
structures, primer TM and primer specificity
49
4.
5.
6.
7.
Rapid Identification and Detection of Pathogenic Fungi by Padlock Probes
are predicted using Visual OMP 6.0 software (DNA Software Inc., Michigan, USA).
The prediction parameters are set to match
ligation
([monovalent] = 0.025
M;
[Mg2+] = 0.01 M; T = 65 °C [probe] = 250 pM)
and PCR conditions ([monovalent] = 0.075 M;
[Mg2+] = 0.005 M, T = 60 °C). When necessary, PRI-lock probe arm sequences are
adjusted to avoid strong secondary structures
that might interfere with efficient ligation as
described previously [13].
Between the primer sites a thymine-linked
desthiobiotin molecule is introduced for
specific capturing and release with streptavidin-coated magnetic beads. The rationale of
using desthiobiotin instead of biotin is the
approximately 1,000 times lower affinity for
streptavidin [33, 34], which permits the
reversible release of the padlock probes.
Reaction mixtures were prepared on ice and
rapidly transferred into a thermal cycler.
OpenArray® real-time PCR amplification of
ligated PRI-lock probes can be followed in
real-time using an OpenArray® Real-Time
PCR Instrument (Applied Biosystems, Foster
City, USA). OpenArray® subarrays are preloaded by Applied Biosystems with selected
primer pairs. Each primer pair is spotted in
duplicate to a final assay concentration of
128 nM.
A PCR control is developed to monitor differences in PCR efficiency within and between different OpenArrays®. To this end, a 99 bp PCR
control template (5¢-CTAACGAATCTGGGAC
G T G C AT C C G G T C T C AT C G C T G
AATCGCTCGTGAGGGCAGGGCCGGG
AGGGGGGTCCGCAGGCGCAACACTGT
AGTCGGTGCTA-3¢) was amplified using the
forward5¢-CTAACGAATCTGGGACGTGC-3¢
and reverse 5¢-TAGCACCGACTACAGTG
TTG-3¢ primer pair.
The OpenArray® Real-time PCR Software
uses a proprietary calling algorithm that estimates the quality of each individual CT value
by calculating a CT confidence value for the
amplification reaction. In our assay, CT values with CT confidence values below 700
were regarded as background signals.
515
8. The cZipCode oligonucleotides carrying a
C12 linker and a 5¢ NH2 group were synthesized and spotted onto Nexterion MPX-E16
epoxy-coated slides by Isogen B.V. according to the manufacturer’s instructions (Schott
Nexterion).
9. The internal transcribed spacer (ITS) regions
of the fungal and oomycetal rRNA genes
are amplified using the primers ITS1 and
ITS4 [35].
10. Sixteen-well silicon structures (Schott
Nexterion) were attached to the arrays to create 16 separate subarray chambers. Before
hybridization, the arrays are washed and
blocked according to the manufacturer’s
instructions.
11. Preamplification reactions are performed in
the presence of 0.1 aM internal amplification
control (IAC) oligonucleotide (5¢TCCGT
AGGTGAACCTGCGGCGGATCGTTA
C A A G G G T C T C C A A C TA C G T C TA G
CGCATAGACCACGTATCGAAGCTAGG
T GCATATCAATAAGCGGAGGA 3¢). It is
often observed that DNA extracted from
environmental samples contain PCRinhibiting compounds [36], which may lead
to false negatives. In order to monitor PCR
inhibition during the preamplification reaction, an IAC was developed. This IAC consist a single-stranded oligonucleotide
containing a random nonsense sequence
flanked by ITS1-ITS4 primer regions, which
is added to each preamplification reaction
mixture. The IAC LD probe containing ligation arms targeting the reverse complement
of the IAC oligonucleotide is added to each
ligation mixture. In the case of PCR inhibition, the complementary strand of the IAC
oligonucleotide will not be generated.
Therefore, the target of the IAC padlock
probes will not be present during the ligation
reaction and, consequently, no IAC padlock
probe signal will be observed on the array.
12. The array hybridization mixtures are made
up of 15 mL of sample and 1 mL of 0.1 mM
Cy5-labeled corner oligonucleotide in 1×
tetramethylammonium chloride in a final
volume of 48 mL.
516
13. Laser power is fixed at 70 % for both lasers,
while the photomultiplier tube power ranged
from 50 to 70 %, depending on the signal
intensity. Fluorescence intensities is
quantified by using QuantArray1 (Packard
GSI Lumonics), and the mean signal minus
the mean local background (mean PE-B) is
used. The absolute signal intensity is defined
as the mean PE-B minus the assay background. The assay background for each subarray is defined as the mean PE−B + 2
standard deviations (SDs) of the fluorescence
of the unused cZipCodes. The cZipCode oligonucleotides are spotted in threefold triplicates (nine parallels) or twofold quadruplicates
(eight parallels), depending on the array
batch used. After exclusion of the outliers
(mean PE−B ± 2 SDs), signals are averaged
for the probes, and SDs calculated.
Acknowledgements We are grateful to Drs. N. Saksena,
S. Chen, and F. Kong (Westmead Hospital, Australia) for
assistance and advice. Part of the funding for this research
has been provided by Genome Canada and Genome
British Columbia in support of The Tria I and Tria II
Projects http://www.thetriaproject.ca.
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Drug-Induced Permeabilization
in Fungi
50
Maria D. Mayan, Alexandra McAleenan,
and Priscilla Braglia
Abstract
One common problem for researchers working with yeast is the difficulty
of efficiently treating whole cells with drugs or chemicals, as their uptake
from the growth medium is very limited. Several methods have been
described to increase drug penetration. However, most of them do not
allow yeast growth under normal conditions. This chapter describes two
protocols for permeabilizing whole yeast cells using either detergents at
4°C, to allow in vitro assays to be executed, or low amounts of the sesquiterpene dialdehyde polygodial, which permits experiments to be performed
in vivo, without affecting yeast morphology or growth.
Keywords
Permeabilization • Sesquiterpene dialdehyde polygodial • Budding yeast
• Detergent
Introduction
M.D. Mayan (*)
Biomedical Research Center-INIBIC,
Xubias de Arriba, 84, A Coruña, Galicia 15006, Spain
e-mail: MA.Dolores.Mayan.Santos@sergas.es
A. McAleenan
Imperial College London, Clinical Sciences Centre,
Hammersmith Hospital Campus, Du Cane Road,
London W12 0NN, UK
P. Braglia
Sir William Dunn School of Pathology, University
of Oxford, South Parks Road, Oxford OX1 3RE, UK
Yeast cells are surrounded by a cell wall and
are not normally permeable to many drugs,
including the transcription inhibitors a-amanitin
and DRB (5,6-Dichloro-1-b-d-ribofuranosylbenzimidazole), which are commonly used to
treat mammalian cells. Digestion of the cell wall
can be achieved using cell lytic enzyme or zymolyase, but handling of the spheroplasts can be
tricky and does not allow studies to be performed
under physiological conditions. Different protocols have been described to increase drug uptake
in yeast cells. For example, the ERG6 gene has
been mutated or deleted to make yeast permeable
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_50, © Springer Science+Business Media, LLC 2013
519
520
to inhibitors of the 26S proteasome. ERG6 mutations affect ergosterol biosynthesis and hence
alter the membrane lipid composition [1–3].
Another example is the use of the antibiotic
amphotericin B which has been used to allow
treatment of yeast cells with the transcription
inhibitors rifampicin and actinomycin D.
However, amphotericin B binds to ergosterol disrupting the fungal cell membrane [4–7]. Different
organic solvents, including ethanol, have also
been used for cell permeabilization [8]. In this
chapter, we describe the use of detergents at 4°C
to perform in vitro assays and polygodial under
normal growth conditions to perform in vivo
assays. Low concentrations of the sesqueterpene
dialdehyde polygodial increase the penetration
of components of the medium into the cells without affecting either the cell membrane or growth
[9–12]. It is therefore of particular interest for
the treatment of living cells under standard
conditions.
Materials
Yeast Permeabilization Using
Detergents
1. Yeast growth medium: prepare the medium to
grow S. cerevisiae as required for the
experiment.
2. Sterile H2O, ice-cold.
3. 10% solution sodium N-lauroyl sarcosine
(Sigma-Aldrich).
4. 2.5x Transcription Buffer: 50 mM Tris/Cl pH
7.7; 500 mM KCl; 80 mM MgCl2.
Yeast Permeabilization Using
Polygodial
1. Polygodial stock solution 3,200×: 1.25 mg
of polygodial (Santa Cruz Biotechnology)
dissolved in 20 mL of absolute ethanol.
The stock solution can be stored for up to
3 months at −20°C, according to manufacturer’s instructions.
M.D. Mayan et al.
Methods
Yeast Permeabilization Using
Detergents
The use of detergents to permeabilize yeast cells
is best exemplified by the protocol commonly
used for transcriptional run-on [13]. TRO assays
allow the density of elongating RNA polymerases
over a desired target gene to be directly measured,
providing a measure of nascent transcription at a
set time-point [14, 15]. Here whole cells are used
to perform in vitro transcription for a short period
of time after providing radioactively labelled
NTPs. The hot RNA is then extracted and hybridized to single-stranded DNA probes, immobilized
on a filter. RNA labelling takes place after the
yeast are permeabilized with sodium N-lauroylsarcosine; this also allows the effective treatment
of cells with a-amanitin (AM), to which yeast
cells are normally impermeable, to measure polymerase sensitivity to this inhibitor.
1. Grow 100 mL cultures to OD600 between 0.05
and 0.2. If, for example, a temperature shift is
required, start it at OD600 = 0.05 and allow the
cells to grow for two generations, to OD600 = 0.2
(see Note 1).
Working at 4°C:
2. Harvest the cells by centrifugation, 4 min at
3,000 rpm.
3. Discard the supernatant and wash the cell pellet with 5 mL ice-cold sterile H2O. Resuspend
by pipetting.
4. Spin again 4 min at 3,000 rpm, discard the
supernatant and resuspend the cell pellet in
950 mL ice-cold sterile H2O. Transfer to
eppendorf tubes.
5. To the cell suspension add 50 mL of 10%
sodium N-lauroyl sarcosine. Mix by inverting
the tubes 5–6 times.
6. Incubate 20 min on ice.
7. Pellet the cells in microcentrifuge, 1 min at
6,000 rpm. Remove supernatant.
8. Resuspend the cell pellet in 60 mL 2.5× transcription buffer and proceed with run-on protocol [14–16].
50
Drug-Induced Permeabilization in Fungi
521
Fig. 50.1 AM and DRB efficiently inhibit transcription
by RNAP-II in permeabilized yeast using low concentrations of polygodial. (a) Analysis of RNA levels by RT-PCR
in rpb1-1 cells at 25°C and following incubation at 37°C.
The temperature shift inactivates RNAP-II in this mutant
strain. Data were normalized to the mature form of the U2
small nuclear RNA. Note that mature snRNAs have been
shown to be stable following RNAP-II inactivation [20].
(b) In cells permeabilized with polygodial, ACT1 mRNA
(ACT1 mRNA, exon1) is stable after 1 h of treatment with
the transcription inhibitors AM or DRB. However the
level of primary transcript (ACT1p, exon1/intron1 junction) decreases by ~90% after treatment with the inhibitors. (c) Non-coding transcripts at the ribosomal locus
(labelled 14, 15, 19, 20 and 21) are transcribed by
RNAP-II. The data relative to the rpb1-1 strain, a temperature sensitive mutant in a key subunit of RNAPII, are
shown in parallel with the data obtained and in the presence of the transcription inhibitors AM and DRB. Data are
presented as mean ± SEM, n = 2
Yeast Permeabilization Using
Polygodial
transcription drops to 10–20% of the initial level
after only 1 h in the presence of 10 mg/mL AM or
45 min in the presence of 200 mM DRB (Fig. 50.1).
Our experience indicates that adding polygodial
to the yeast culture only a few minutes before the
treatment is enough to promote the entry of different drugs.
1. Grow the yeast as desired (see Note 2).
2. Add 0.39 mg/mL polygodial resuspended in
absolute ethanol. Mix the culture (see Note 3).
3. Split the culture into two flasks, untreated and
treated culture.
4. Add the desired drug to the medium at an
appropriate concentration.
5. Grow the cells for the desired treatment time.
6. Collect the cells as required for subsequent
protocols.
Adding polygodial to a yeast cell culture a few
minutes before treatment with different drugs,
immediately promotes uptake of the drug into the
cell [11, 12]. Polygodial should not be used at
concentrations higher than 1 mg/mL as higher
levels of polygodial affect the integrity of the
membrane and therefore the cell metabolism,
interfering with the results obtained. A good
example of the use of polygodial is provided by
the treatment with the RNA polymerase II
(RNAP-II) inhibitors AM or DRB. AM and DRB
inhibit transcription elongation by different
mechanisms [17–19]. After treating the cells with
0.39 mg/mL polygodial, the level of RNAP-II
522
M.D. Mayan et al.
Notes
1. When comparing different strains/conditions,
ensure cultures are grown to similar OD600.
This might require dilution of the fastergrowing strain [16]. Cultures grown to a higher
OD600 have empirically resulted in lower TRO
signal.
2. Temperatures from 23 to 39°C have been used
in the presence of 0.39–1 mg/mL of polygodial
without affecting cell membrane structure.
3. For 200 mL of culture add 62.4 mL of polygodial 1.25 mg/mL. Using this stock, the ethanol
is diluted 3,200 times in the culture. No further incubation time with polygodial is needed:
the 5–10 min spent dividing the culture into
two different flasks, is enough to promote the
entrance of the drugs after adding polygodial.
7.
8.
9.
10.
11.
12.
13.
Acknowledgements The experimental work was performed in Professor Luis Aragon’s lab. We thank R. Young
for the Z118 strain (rpb1-1). This work was supported by
the Medical Research Council of the United Kingdom.
Maria Mayán is currently funded by the Xunta de Galicia.
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Extraction and Characterization
of Taxol: An Anticancer Drug
from an Endophytic and Pathogenic
Fungi
51
M. Pandi, P. Rajapriya, and P.T. Manoharan
Abstract
The basic characteristic feature of cancer is the transmissible abnormality
of cells that is manifested by reduced control over growth and cell division, leading to serious adverse effects on the host through invasive growth
and metastases. Abnormal development of cells leads to the growth of
tumor; when the tumor is malignant in nature, it is termed as cancer.
Cancer is one of the most common causes of premature death in the world.
The most recent estimate of cancer indicates that 8.1 million new cases are
diagnosed worldwide each year. Breast cancer is the second most prevalent cancer worldwide and its incidence is gradually increasing. Paclitaxel
(taxol) is the most effective antitumor agent developed in the past three
decades. Taxol has been hailed by many in the cancer community as a
major breakthrough in the treatment of cancer. Taxol was originally isolated from the bark of the Pacific yew Taxus brevifolia in 1971. The
increased demand for taxol, coupled with its limited availability from the
protected Pacific yew, has had researchers scrambling for alternate sources.
The endophytic and pathogenic fungi can produce taxol as a cheaper and
more widely available product, eventually via industrial fermentation.
This chapter deals with the isolation and identification of endophytic and
pathogenic fungi and the extraction and characterization of taxol, an anticancer drug from endophytic and pathogenic fungi.
M. Pandi (*)
Department of Molecular Microbiology,
School of Biotechnology, Madurai Kamaraj University,
Madurai, Tamil Nadu 625 021, India
e-mail: an_pandi@rediffmail.com
P. Rajapriya
Department of Microbiology, Srinivasan College
of Arts and Science, Perambalur,
Tamil Nadu 621 212, India
P.T. Manoharan
Department of Botany, Vivekananda College, Madurai,
Tamil Nadu 625 217, India
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_51, © Springer Science+Business Media, LLC 2013
523
524
M. Pandi et al.
Keywords
Cancer • Paclitaxel (taxol) • Antitumor • Endophytic fungi • Pathogenic
fungi
Introduction
Taxol is a chemotherapic drug specifically effective against prostate, ovarian, breast, and lung
cancer. Its primary mechanism of action is
related to the ability to stabilize the microtubules
and to disrupt their dynamic equilibrium [1–4].
Taxol inhibits cell proliferation by promoting
the stabilization of microtubules at the G-M
phase of the cell cycle, by which depolymerization of microtubules to soluble tubulin is blocked
[5–7]. Taxol was originally isolated from the
bark of the Pacific yew, Taxus brevifolia [8]. The
limited availability of mature yew trees, slow
growth rate of cultivated plants, and the low
yield of the taxol has resulted in its high cost and
also has raised concerns about environmental
damage from excessive exploitation of wild
trees. This makes taxol a financial burden for
many patients. The search for an alternative
source of taxol other than the bark of the yew
trees (Taxus sp.) has been carried out by scientists all over the world to meet the demand in
clinics. The most significant finding in the last
decade might be the discovery of endophytic
taxol-producing fungus in Gymnosperm, particularly in yew trees. It is remarkable that the taxol
produced by the endophytes is identical to that
produced by Taxus spp., chemically and biologically [9–16].
The production of taxol by using fungi has
given rise to the possibility of reduced cost and
wider availability, as taxol may eventually be
available via large-scale industrial fermentation.
The fungus can serve as a potential material for
genetic engineering to improve taxol production.
The purpose of this chapter is to focus on the isolation and identification of endophytic and pathogenic fungi and the extraction and characterization
of taxol, an anticancer drug from endophytic and
pathogenic fungi.
Materials
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
Sterile distilled water.
Leaves of selected plants.
Ethanol (75% V/V).
Sodium hypochloride (2.5% V/V).
Potato dextrose agar medium (PDA).
Ampicillin (200 mg/mL) and streptomycin
(200 mg/mL).
Hand lens.
Clean glass slides.
Petri plates and blotting papers.
Stereomicroscope.
Lactophenol and DPX mountant.
Carl Zeiss Axiostar Plus—Photomicroscope.
0.1% mercuric chloride solution.
5 mg/g chloramphenicol.
MID medium supplemented with 1 g soy
tone/L.
0.25 g of Na2Co3 (0.025% W/V).
Dichloromethane.
Rotary evaporator.
1% vanillin sulfuric acid (w/v).
100% methanol.
Beckman DU-40 UV-Spectrophotometer.
IR grade potassium bromide (KBr).
Methanol/acetonitrile/water (25:35:40, by
vol.).
Methods
Isolation and Identification
of Endophytic Fungi
The leaves of selected plants were collected from
different places. The plant materials were subjected to endophytic isolation within 3 h after
harvest. The endophytic fungal cultures were
separated from the healthy leaves according to
the general mycological procedure [11–14, 16].
51
Extraction and Characterization of Taxol…
1. The leaves were washed with running tap
water, sterilized with ethanol (75% V/V) for
1 min and sodium hypochloride (2.5% V/V)
for 5 min, then rinsed in sterile water for three
times and cut into 1 cm long segments.
2. Plant segments were then transferred to
potato dextrose agar containing Petri plates
amended with ampicillin (200 mg/mL) and
streptomycin (200 mg/mL) to inhibit bacterial
growth.
3. After 2 days of incubation, mycelia of fungi
were observed growing from the inner leaf
segments in the plates.
4. Individual hyphal tips of the various fungi
were carefully removed from the agar plates
with the help of an inoculation loop; then they
were placed on new PDA medium and incubated at 25 °C for at least 7–10 days.
5. Each fungal culture was checked for purity
and subcultured to another agar plate by the
hyphal tip method.
6. Fungal identification was based on the morphology of the fungal culture, the mechanism
of spore production, and the characteristics of
the spores [17].
Isolation of Plant Pathogenic Fungi
1. The pathogenic infected leaves of various
plants with symptoms showing Coelomycetes
fungi were collected from different places in
Tamilnadu, India.
2. Coelomycetes fungi are found on dead twigs,
leaf litter, bark, and infected leaves.
3. The fungal infection was confirmed by the
presence of conidiomata on the substrate using
a hand lens in the field.
4. Then the specimen is collected. The data should
be recorded on the envelope used to transport
the material to the laboratory. It is essential that
the identity of the substrate should be accurately known for the identification of many species is still based on a host basis.
5. Unless the material is examined immediately
it should be dried thoroughly to prevent the
growth of molds and unwanted saprophytes.
525
Examination of the Specimens
The following methods were used to study
Coelomycetes fungi on the specimen.
Direct Examination of the Specimen
Once the selected conidioma has been removed
from the substrate, it is transferred to a clean
glass slide and mounted with water, for microscopic examination.
Moist Chamber Incubation Method
1. This technique was used to induce
sporulation.
2. The specimen was incubated in a 15-cm diameter sterilized Petri plate lined with moist blotting paper.
3. The plates were kept moist by adding sterile
distilled water periodically, but the blotting
paper was never flooded with water.
4. The specimens were examined after a week
under a stereomicroscope and the conidiomata
on them were studied.
5. The fungi found in sporulating conditions
were isolated, examined, and identified down
to species level [17].
Hand Section
1. Cutting vertical section of conidiomata is essential to confirm the nature of the fructification.
2. Specimens with fruit bodies were sectioned
with the help of razor blade.
3. Good sections were selected through stereomicroscope; then they were mounted in lactophenol and viewed under a light microscope.
Preparation of Permanent Slides
For the preparation of permanent slides, water
and lactophenol were used. The slides were
sealed by DPX mountant.
Illustration and Photomicrographs
Photomicrographs of conidia were taken with
the help of Carl Zeiss Axiostar Plus—
Photomicroscope (Phase contrast) with Nikon
FM 10 camera and Nikon HF X Labophot (bright
field) with Nicon Fx—35A by using Konica
films.
526
Isolation of Single Pathogenic Fungi
1. The infected plant parts of approximately
4 mm2 were sterilized by immersing for 1 min
in 0.1% mercuric chloride solution and washed
by successive transfer through sterile distilled
water three times.
2. These were then incubated in sterile distilled
water containing 5 mg/g chloramphenicol.
3. A few drops of spore suspensions were spread
over Potato dextrose agar plates and incubated
at 25 °C for 6–48 h.
4. Germinated spores were transferred to oatmeal agar or Potato dextrose agar plates.
Subcultures were also made from spores
extruding from pycnidia produced in culture.
5. Individual hyphal tips of the various fungi
were removed from the agar plates, placed on
new PDA medium and incubated at 25 °C for
at least 2 weeks.
6. Each fungal culture was checked for purity
and sub cultured to another agar plate by the
hyphal tip method.
7. Fungal identification methods were based on
the morphology of the fungal culture, the
mechanism of spore production, and the characteristics of the spores [17].
Cultivation and Extraction
of Taxol from Selected Fungal
Isolates
1. The selected endophytic or pathogenic isolates were inoculated into a 2,000-mL Hopkins
flask of MID medium supplemented with 1 g
soy tone/L, incubated for 12 h under light and
dark cycle at temperature between 22 and
25 °C for 21 days.
2. After 21 days, the cultures were passed
through four layers of cheesecloth and 0.25 g
of Na2CO3 (0.025% W/V) was added to the
culture filtrate to avoid fatty acid
contamination.
3. The culture filtrate was further extracted with
twice the volume of dichloromethane and the
organic phase was taken to dryness under
reduced pressure at 50 °C using a rotary
evaporator.
M. Pandi et al.
4. Then the dry solid residue was re-dissolved in
methanol for the subsequent separation.
5. The presence of taxol was confirmed in the
crude extracts, which were analyzed using different chromatographic and spectroscopic
methods.
Thin-Layer Chromatography
1. TLC analysis was carried out on Merck 1-mm
(20 × 20 cm) silica gel precoated plates.
2. The plates were developed by the solvent system reported by Strobel et al. [18, 19].
3. The taxol was detected with 1% vanillin sulfuric acid (w/v) and heating. It appears as a bluish spot that faded to dark grey after 24 h.
4. Then the area of the plate containing putative
taxol was carefully removed by scraping off
the silica at the appropriate Rf value and eluted
with methanol. Then they were further analyzed by UV, IR, and HPLC to confirm the
production of taxol.
Ultraviolet Spectroscopic Analysis
1. After chromatography, the area of the TLC
plate containing putative taxol was carefully
removed by scrapping off the silica at the
appropriate Rf and exhaustively eluting it with
methanol.
2. The purified sample of taxol was dissolved in
100% methanol and analyzed by Beckman
DU-40 UV-Spectrophotometer and compared
with authentic taxol.
Infrared Spectroscopic Analysis
1. The IR spectra of the compound were recorded
on Shimadzu FTIR 8000 series instrument.
2. The purified taxol was ground with IR-grade
potassium bromide (KBr) (1:10) pressed into
discs under vacuum using spectra lab Pelletiser
and compared with authentic Taxol.
3. The IR spectrum was recorded in the region
between 4,000 and 5,000 cm.
51
Extraction and Characterization of Taxol…
High-Performance Liquid
Chromatography Analysis
1. To confirm the presence of taxol, the fungal
extract was subjected to high performance liquid chromatography (HPLC).
2. Taxol was analyzed by HPLC (Shimatzu 9A
model) using a reverse phase C18 column with
a UV detector.
3. Twenty microliter of the sample were
injected each time and detected at 232 nm.
The mobile phase was methanol/acetonitrile/
water (25:35:40, by vol.) at a flow rate of
1.0 mL/min.
4. The sample and the mobile phase were filtered
through 0.2 mm PVDF filter before injecting
into the column.
5. Fungal taxol was confirmed by comparing the
peak area of the samples with authentic taxol.
Acknowledgement I thank the University Grants
Commission, New Delhi, India, for the financial support
of the Research Grant.
References
1. Horwitz SB (1992) Mechanism of action of taxol.
Trends Pharmacol Sci 13:131–6
2. Rao S, Orr GA, Chaudhary AG, Kingston DGY,
Horwitz SB (1995) Characterization of the taxol binding site on the microtubule, 2-(m-Azidobenzoyl)
Taxol photolabels a peptide (amino acids 217–231) of
beta-tubulin. J Biol Chem 270:20235–8
3. Jordan MA, Wilson L (1998) Microtubules and actin
filaments: dynamic targets for cancer chemotherapy.
Curr Opin Cell Biol 10:123–30
4. Caplow M, Shanks J, Ruhlen R (1994) How taxol
modulates microtubule disassembly. J Biol Chem
38:23399–402
5. Horwitz SB (1994) Taxol (paclitaxel): mechanisms of
action. Ann Oncol 6:3–6
6. Nicolaou KC, Yang Z, Liu JJ, Ueno H, Nantermet PG,
Guy RK (1994) Total synthesis of taxol. Nature
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molecular genetics, and biotechnological applications. Appl Microbiol Biotechnol 57:13–19
8. Wani MC, Taylor HL, Wall ME, Coggon P, McPhail
AT (1971) Plant antitumor agents VI. The isolation
and structure of taxol, a novel antileukemic and antitumor agent from Taxus brevifolia. J Am Chem Soc
93:2325–2327
9. Stierle A, Strobel G, Stierle D (1993) Taxol and taxane production by Taxomyces andreamae, an endophytic fungus of Pacific yew. Science 260:214–6
10. Wang J, Li G, Lu H, Zheng Z, Huang Y, Su W (2000)
Taxol from Tubercularia sp. strain TF5, an endophytic
fungus of Taxus mairei. FEMS Microbiol Lett
193:249–53
11. Gangadevi V, Muthumary J (2007) Taxol, an anticancer drug produced by an endo-phytic fungus Bartalinia
robillardoides Tassi, isolated from a medicinal plant
Aegle marmelos Correa ex Roxb. World J Microbiol
Biotechnol 24:717–24
12. Gangadevi V, Muthumary J (2009) Taxol production
by Pestalotiopsis terminaliae, an endophytic fungus
of Terminalia arjuna. Biotechnol Appl Biochem
52:9–15
13. Gangadevi V, Muthumary J (2008) Isolation of
Colletotrichum gleosporioides, a novel endophytic
Taxol-producing fungus from the leaves of a medicinal plant Justicia gendarussa. Mycol Balcanica 5:1–4
14. Senthil Kumaran R, Hur BK, Muthumary J (2008)
Production of taxol from Phyllosticta spinarum, an
endophytic fungus of Cupressus sp. Eng Life Sci
8(4):1–10
15. SenthilKumaran R, Muthumary J, Hur B-K (2008)
Taxol from Phyllosticta citricarpa, a leaf spot fungus
of the angiosperm Citrus medica. J Biosci Bioeng
106(1):103–6
16. Pandi M, Senthil KR, Choi Y-K, Kim HJ, Muthumary
JP (2011) Isolation and detection of Taxol, an anticancer drug produced from Lasiodiplodia theobromae, an
endophytic fungus of the medicinal plant Morinda citrifolia. Afr J Biotechnol 10(8):1428–1435
17. Sutton BC (1980) The coelomycetes. Fungi imperfecti with pycnidia, acervuli and stromata.
Commonwealth Mycological Institute, Kew, Surrey
18. Strobel G, Yang X, Sears J, Kramer R, Sidhu RS, Hess
WM (1996) Taxol from Pestalotiopsis microspora, an
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(1996) Taxol from fungal endophytes and the issue of
biodiversity. J Ind Microbiol 17:417–423
Identification of Mycotoxigenic
Fungi Using an Oligonucleotide
Microarray
52
Eugenia Barros
Abstract
Mycotoxins are secondary metabolites produced by fungi; they can play a
role as food contaminants and have the ability to negatively influence
human and animal health. To improve food safety and to protect consumers from harmful contaminants, numerous detection tools have been developed for the detection and analysis of various mycotoxigenic fungi. These
include PCR-based assays and microarrays targeting different areas of the
fungal genome depending on its application. This chapter describes the
development of an oligonucleotide microarray specific for eleven mycotoxigenic fungi isolated from different food commodities in South Africa.
This array is suitable for the detection and identification of cultures of
potential mycotoxigenic fungi in both laboratory samples and commodityderived food samples.
Keywords
Mycotoxins • Fungi • Oligonucleotide microarray • Food contaminants
Introduction
Fungi can grow on many food commodities but
when these fungi are able to produce health
threatening substances, such as mycotoxins, they
then become a threat to human and animal health.
Although the presence of a mycotoxigenic fungus in a food commodity does not necessarily
E. Barros (*)
Department of Biosciences, Council for Scientific
and Industrial Research (CSIR), Meiring Naude Road,
Brummeria, Pretoria 0001, South Africa
e-mail: ebarros@csir.co.za
indicate the production of the respective mycotoxin it is however necessary to control the presence of fungal contaminants in the food production
chain and develop quicker methods to identify
food-borne fungi.
Many molecular biology techniques have been
developed as alternative approaches to detect and
quantify fungal growth so as to prevent contaminated commodities from entering the food chain.
Most of the developed molecular techniques used
restriction enzymes and the polymerase chain
reaction (PCR) to assess the genetic variability
within and among fungal species taking advantage of the polymorphisms that occur naturally in
the DNA of a given species. Successful assays
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_52, © Springer Science+Business Media, LLC 2013
529
530
have utilized the highly conserved ribsosomal
RNA gene sequences to design species-specific
PCR primers [1] and gene-specific PCR detection assays for genes involved in the biosynthesis
of mycotoxins [2, 3].
The microarray technology provides a tool to
potentially identify and quantify levels of gene
expression for all genes in an organism. Small
spots of DNA are fixed to a matrix; this can be
either glass or a nylon membrane. Microarrays can
be constructed using cDNAs, genomic sequences
or oligonucleotides synthesized in silico. Typically,
cDNAs used to construct microarrays are partial
gene sequences that are derived from coding DNA
and generally have a high degree of sequence conservation. Genomic sequences include sequences
that target specific genes of interest like toxin producing genes in the case of fungal biology and,
oligonucleotides include those synthesized from
polymorphic sequences that occur naturally in the
DNA of a given species.
In fungal biology microarrays can be used for
a variety of applications. Some applications
include identification of fungal species;
identification of potential mycotoxigenic fungi;
to study gene expression and thus identify differentially expressed genes; and identification of
toxin genes. However the design of the microarray is dependent on its application and the most
common arrays are cDNA microarrays [4] and
oligonucleotide microarrays [5]. There are different types of oligonucleotide microarrays and they
include oligonucleotides synthesized from (1)
polymorphic fragments identified by molecular
marker techniques, like for example, amplified
fragment length polymorphisms (APFLs); (2)
conserved regions like the internal transcribed
spacer (ITS) regions and (3) gene sequences like
fungal toxin genes, among others.
The present contribution focuses on the generation of probes that showed polymorphisms
within the internal transcribed spacer (ITS)
regions of rRNA of eleven mycotoxigenic fungi
and the development of an oligonucleotide
microarray that can detect and identify these fungi
from different food commodities. The eleven
mycotoxin producing fungi are amongst the most
prevalent in South African food commodities [5].
E. Barros
Furthermore, the technique used to label the target
DNA was random labelling and it does not involve
amplification of target DNA prior to microarray
hybridization. This technique avoids amplification
bias, diminishes secondary structures and ensures
a more efficient target [5, 6].
Materials
Fungal Strains
Isolates of eleven food-borne fungi known to produce mycotoxins were obtained from the
Agricultural Research Council (ARC) culture
collection in Pretoria, South Africa. These
included Aspergillus carbonarius, Aspergillus
clavatus, Aspergillus niger, Alternaria alternata,
Eurotium amstelodami, Penicillium corylophilum, Penicillium expansum, Penicillium fellutanum, Penicillium islandicum, Penicillium
italicum, and Stenocarpella maydis.
Culture Media
Fungal strains were grown on Malt Salt Agar
(MSA) at 25 °C for 1–2 weeks (see Note 1). MSA
was prepared by dissolving 90 g NaCl in 360 mL
dH2O in a Schott bottle (solution 1); 24 g of malt
extract (Merck, South Africa) and 24 g of agar
(Merck, South Africa) were added to 840 mL of
dH2O in a Schott bottle (solution 2). The solutions were autoclaved for 20 min at 121 °C.
Solution 1 was then added aseptically to solution
2 and the medium poured into 90-mm petri dishes
and allowed to settle.
DNA Extraction Buffer
Genomic DNA was extracted from the eleven
fungal cultures using the method of Raeder and
Broda [7]. The DNA extraction buffer contained
200 mM Tris–HCl (pH 8), 150 mM NaCl, 25 mM
EDTA (pH 8), 0.5% SDS and 1% PVP. Just before
use, 0.2% (v/v) 2-Mercapto-ethanol was added to
the extraction buffer.
52
Identification of Mycotoxigenic Fungi Using an Oligonucleotide Microarray
Methods
Fungal DNA Extraction
1. Fungal mycelium was gently scrapped off
the cultures grown on MSA media and placed
into 1.5-mL eppendorf tubes together with
metal yellow tungsten carbide beads (3 mm)
and in 500 mL DNA extraction buffer, as
described in the section DNA Extraction
Buffer, and according to the method of
Raeder and Broda [7].
2. The tubes were span in a FastPrep® machine
(MP Biomedicals, Cambridge, UK) at speed
4 for 5 s (see Note 2).
3. The metal beads were removed and the contents transferred to a 1.5-mL eppendorf tube
to which 50 mL 1 M Tris–HCl (pH 8.0),
100 mL phenol and 170 mL chloroform were
added and the tubes placed on ice for 5 min.
4. The suspension was then centrifuged at
20,817 × g for 15 min and the aqueous phase
transferred to a new eppendorf tube.
5. The aqueous phase was cleaned once again
with 50 mL 1 M Tris–HCl (pH 8.0), 100 mL
phenol and 170 mL chloroform as described
in the previous step and the cleaning procedure repeated until the interface was clean.
6. Chloroform, 1× volume, was added to the
clean aqueous phase and the samples centrifuged at 10,621 × g for 15 min.
7. The aqueous phase was transferred to a new
eppendorf tube to which 2.5× volume of ethanol and 0.5 M ammonium acetate were
added and the solution allowed to precipitate
overnight at −20 °C.
8. The suspension was centrifuged at 10,621 × g
for 10 min, the supernatant discarded and the
pellet washed with 2.5× volume of 70% ethanol followed by centrifugation at 10,621 × g
for 10 min.
9. The supernatant was discarded, the pellet allowed to air dry and then resuspended
with 50 mL of ddH2O.
10. The DNA concentration and purity were
assessed from the absorbance measurements
with the Nanodrop 1000 instrument.
531
PCR Amplification of Fungal DNA
with ITS1 and ITS4 Universal Primers
The ITS regions of the eleven fungi were
amplified using universal fungal primers for ITS1
(5¢-TCCGTAGGTGAACCTGCGG-3¢) and for
ITS4
(5¢-TCCTCCGCTTATTGATATGC-3¢)
according to White et al. [8]. The PCR
amplifications were carried out using the following reaction mixture in a total volume of 25 mL:
8 ng fungal template DNA, 1.5 mM MgCl2,
0.2 mM of each dNTP, 0.5 U Taq polymerase
(Bioline), 1× PCR reaction buffer (Bioline) and
0.4 mM of ITS1 primer and 0.4 mM of ITS4
primer. The PCR amplification consisted of an
initial denaturation step of 94 °C for 5 min; followed by 35 cycles of denaturation at 94 °C for
30 s, primer annealing at 50 °C for 45 s and
primer extension at 72 °C for 1 min; and a final
extension at 72 °C for 5 min.
Amplicon Sequencing, Identification
of Polymorphisms and Probe Design
Aliquots of the PCR products were separated on an
agarose gel (1.4%) for quality control. The remainder of each PCR product was precipitated in a
NaAc/EtOH solution made up of 90% ethanol and
0.9 mM NaAc (pH 5.2). The precipitate was collected by centrifugation at 3,600 × g for 30 min. The
pellets were washed in 70% ethanol, dried, and
then resuspended in 50 mL dionized H2O. Aliquots
of the resuspended amplicons were sequenced by
Inqaba Biotec (Pretoria, South Africa).
The sequenced fragments were aligned using
ClustlX software as described by Thompson et al.
[9] and polymorphisms were identified. These
sequences were then used to design genusspecific and species-specific probes of various
lengths, ranging from 14 to 25 bases, and within
a narrow range of melting temperature, 56 ± 5 °C.
The oligonucleotide probes were designed using
the Primer Designer 4 Package, Version 4.2
(Scientific and Educational Software, Cary, NC).
The probe sequences generated for each of the
eleven fungi as well as the specific annealing
temperatures are shown in Table 52.1.
532
Table 52.1 Fungal isolates, potential mycotoxins produced, and probe sequences generated to construct the oligonucleotide microarray
Probe name and reference
AR1 [5]
ACIF [5]
ANIG [5]
Aaf AaR [5]
EurAF EurAR [5]
PenCorF PenCorR [5]
PenExF PenExR [5]
PenFeF PenFeR [5]
PenIsF PenIsR [5]
PenltF PenltR [5]
4 F 4R [11]
Probe sequence (5¢→3¢)a
ATCTGCTGCACAGTTGGCT
ATTCGGAAACCUGCTCAGTACG
ACGTTATCCAACCAT
GACCGCTTTCGTGGTATGCA
TGGCGGCACCATGTC
TGGTTAAAAGATTGGTTGCGA
GTCCAAACCCTCCCACCCA
GTCAGACTTGCAATCTTCAGACTGT
TTACCGAGTGAGGCCGT
GCCAGCCTGACAGCTACG
CTGAGTGCGGGCCCTCT
CGCCGAAGCAACACTGTAAG
CGAGTGCGGGTTCGACA
GGCAACGCGGTAACGGTAG
CTCCCACCCGTGTTTATTTATCA
TCACTCAGACGACAATCTTCAGG
CAAACGTCGGGTCAGAAGAAGCGAC
AGGAACCGTCCCCGCCGACGTTTG
Fungal isolates
Aspergillus carbonarius
Aspergillus clavatus
Aspergillus niger
Alternaria alternata
Eurotium amstelodami
Mycotoxins produced
Ochratoxin A
Cytochalacin E; Patulin
Ochratoxin A
Tenuazonic acid
Sterigmatocystin
Annealing temperature
(°C) for PCR amplification
56
58
55
56
58
Penicillium corylophilum
Cyclopiazonic acid
55
Penicillium expansum
Patulin
58
Penicillium fellutanum
Patulin
55
Penicillium islandicum
57
Penicillium italicum
5,6, dihydro-4-methoxy-2 H-pyran-2-one
Patulin
Stenocarpella maydis
Diplosporin
57
57
Locked nucleic acids (LNAs) that were used to increase the specificity of a probe are in bold and italic
a
E. Barros
52
Identification of Mycotoxigenic Fungi Using an Oligonucleotide Microarray
Ensuring Uniqueness and Specificity
of Probes
The specificity of each oligonucleotide probe was
further assessed by subjecting the sequence/s to
similarity searches in public databases; the databases used included NCBI (http://www.ncbi.nlm.
nih.gov) and EMBL (www.ebi.ac.uk/embl).
BLAST searches were used and only unique oligonucleotide probes were selected for printing on
the array. In cases where probes had similar
sequences their specificity was enhanced by substituting an oligonucleotide with a high affinity
DNA analogue known as locked nucleic acid
(LNA) according to the method of Johnson,
Haupt, and Griffiths [10]. This technique was
also used in the design of some of the probes to
ensure that the set of oligonucleotide probes that
were printed on the array had similar hybridization efficiencies. This approach was used for the
probes specific for A. alternata and P. expansum
and the LAN is indicated in bold and italic in
Table 52.1. All the probes, including the designed
probes, were synthesized by Inqaba Biotech
(Pretoria, South Africa).
Construction of the Array
The 18 uniquely designed species-specific oligonucleotide probes were used to construct the
array together with three control probes consisting of ITS1, ITS3 and ITS4 fragments. Equal
volumes (10 mL each) of 100 pmol/mL oligonucleotide and 100% DMSO were transferred into a
384-well plate (Amersham Pharmacia Biotech).
Sixteen replicates of each of each oligonucleotide
were arrayed onto Vapour Phase-coated Glass
Slides (Amersham Pharmacia Biotech) using an
Array Spotter Generation III (Molecular
Dynamics, Sunnyvale, CA, USA) at the
Microarray Facility of the African Centre for
Gene Technologies (ACGT), University of
Pretoria, South Africa (http://fabinet.up.ac.za/
microarray). Following printing the slides were
allowed to dry overnight at 45–50% relative
humidity. Spotted DNA was bound to the slides
by UV cross-linking at 250 mJ and then baked
533
for 2 h at 80 °C. The control ITS fragments were
spotted at concentrations of 50 ng/mL, 100 ng/mL,
150 ng/mL, and 200 ng/mL.
Preparation of Labelled Target DNA
DNA was extracted from the eleven fungal cultures following the DNA extraction protocol
described in the section Fungal DNA Extraction.
Two micrograms of DNA were labelled with redfluorescent dye Cyanine 5 (Cy5) using the Cy™
Dye Postlabelling Reactive Dye Pack (GE
Healthcare, UK). For each labelling reaction the
DNA was diluted in 5 mL 0.2 M Na2CO3 (pH 9)
and 2.5 mL Cy5 mono NHS ester 4,000 pmol dye
resuspended in 12 mL DMSO. The reactions were
incubated in the dark, at room temperature, for
90 min. After labelling the dye-coupling reaction
was column-purified using the QIAquick PCR
purification Kit (QIAGEN Germany).
Preparation of Labelled Control Probes
Each of the control probes ITS1, ITS3, and ITS4
were labelled by incorporation of greenfluorescent dye Cyanine 3 (Cy3-dUTP)
(Amersham Biosciences, Buckinghamshire, UK)
using the Klenow fragment DNA polymerase I
(Roche Diagnostics). Each labelling reaction
contained 5 mg of probe DNA, 1.8 mM dNTP
mix (0.3 mM dATP, 0.3 mM dGTP, 0.3 mM
dCTP and 0.8 mM dTTP), 0.1 mM Cy3-dUTP,
1× hexanucleotide mix (Roche Diagnostics) and
8 U Klenow enzyme (Roche Diagnostics). The
reaction was incubated at 37 °C overnight. After
labelling the dye-coupling reaction was columnpurified using the QIAquick PCR purification Kit
(QIAGEN Germany).
Hybridization of Array Slides
The Cy5-labelled target DNA and the Cy3labelled control probes (0.3 pmol) were resuspended in 40 mL of hybridization mixture
containing 50% formamide (Sigma-Aldrich),
534
25% 2× hybridization buffer (Amersham
Pharmacia Biotech) and 25% dionized H2O. The
mixture was denatured at 95 °C for 5 min and
stored on ice for hybridization. The hybridization
mixture was then pipetted onto a glass slide
(24 × 60 mm, No. 1, Marienfeld, Germany), covered with a cover slip and inserted into a custommade hybridization chamber (N.B. Engineering
Works, Pretoria, South Africa) and allowed to
hybridize overnight at 53 °C. The slides were
then washed twice in 2× SSC and 0.2% SDS at
37 °C for 6 min, once in 0.2× SSC and 0.2% SDS
at room temperature for 5 min and then twice in
0.075× SSC at room temperature for 5 min. The
slides were rinsed in de-ionized H2O for 2 s and
dried by centrifugation at 1,000 × g for 5 min.
Scanning and Data Processing
The oligonucleotide arrays were scanned using a
GenePix 4000B Scanner (Molecular Dynamics,
USA) and the mean pixel intensity within each
spot and the local background were determined
using Array Vision, version 6.0 software (Imaging
Research Inc., Molecular Dynamics, USA).
All signal intensities were background corrected
by subtracting the local background from the
raw spot intensity value. The spot intensity data
generated by the control samples were used as a
reference for normalization of all spot intensity
data. Irregular spots were manually flagged for
removal and further data analysis was performed
in the Microsoft Excel software (Microsoft,
Richmond, WA).
Protocols describing the downstream data
analysis confirming the reproducibility of the
array, data processing and statistical analysis fall
outside the scope of this chapter.
Notes
1. Growth of different fungal species on solid
media is variable. However, 2 weeks incubation at 25 °C was found to be the optimum
period to obtain sufficient fungal mycelium
E. Barros
for DNA extractions, but it can be extended if
necessary.
2. If a FastPrep® machine is not available, mix
thoroughly the tubes containing the mycelium,
the metal beads, and the DNA extraction buffer and incubate for 15 min and at 60 °C in an
ultrasonic bath (e.g., S10H from Elma, Singen,
Germany).
References
1. Mishra PK, Fox RTV, Culham A (2003) Development
of a PCR-based assay for rapid and reliable
identification of pathogenic Fusaria. FEMS Microbiol
Lett 218:329–332
2. Paterson RRM, Archer S, Kozakiewicz Z, Lea A,
Locke T, O’Grady E (2000) A gene probe for the patulin metabolic pathway with potential for use in patulin
and novel disease control. Biocontrol Sci Technol
10:509–512
3. Waalwijk C, van der Lee T, de Vries I, Hesselink T,
Arts J, Kema GHJ (2004) Synteny in toxigenic
Fusarium species: the fumonisin gene cluster and the
mating type region as examples. Eur J Plant Pathol
110:533–544
4. Barros E, van Staden C, Lezar S (2009) A microarraybased method for the parallel analysis of genotypes
and expression profiles of wood-forming tissues in
Eucalyptus grandis. BMC Biotechnol 9:51
5. Lezar S, Barros E (2010) Oligonucleotide microarray
for the identification of potential mycotoxigenic fungi.
BMC Microbiol 10:87
6. Lane S, Everman J, Logea F, Call DR (2004) Amplicon
structure prevents target hybridization to oligonucleotide microarrays. Biosensors Bioelectron 20:728–735
7. Raeder U, Broda P (1985) Rapid preparation of DNA
from filamentous fungi. Lett Appl Microbiol 1:17–20
8. White TJ, Bruns T, Lee S, Taylor J (1990) Amplification
and direct sequencing of fungal ribosomal RNA genes
for phylogenetics. In: Gelfand DH, Sninsky JJ, White
TJ, Innis MA (eds) PCR protocols. a guide to methods
and applications. Academic, San Diego, CA, pp
315–322
9. Thompson JD, Gibson TJ, Plewniak F, Jeanmougin F,
Higgins DG (1997) The CLUSTL_X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tool. Nucleic Acids
Res 25:4876–4882
10. Johnson MP, Haupt LM, Griffiths LR (2004) Locked
nucleic acids (LNA) single nucleotide polymorphism
(SNP) genotype analysis and validation using realtime PCR. Nucleic Res 32:e55
11. Barros E, Crampton M, Marais G, Lezar S (2008) A
DNA-based method to quantify Stenocarpella maydis
in maize. Maydica 53:125–129
DNA Microarray-Based Detection
and Identification of Fungal
Specimens
53
Minna Mäki
Abstract
Novel DNA-based molecular methods can be used to detect the fungal
species faster than with the conventional methods. Combination of PCR
and microarray provides rapid, sensitive, and reliable detection of pathogenic fungi. The advantages over other DNA-based methods are that
microarray technologies allow broader coverage of detectable targets and
simultaneous detection of multiple targets in a single assay. Furthermore,
microarray technologies have the potential to discriminate between closely
related fungal species. Although the use of microarray technologies in
clinical diagnostics is still rare, the microarray-based approaches are
believed to have great clinical potential in the field of infectious diseases.
Keywords
DNA • Polymerase chain reaction • Microarray • Pathogen • Fungi
• Identification
Introduction
Conventional microbiological diagnostics of a
fungal infection mainly rely on microscopic and
cultural techniques that are time-consuming,
labor-intensive, and require expertise. These
methods usually yield diagnostic results in days
or in some cases up to weeks after sampling.
Furthermore, cultivation of fungi is not always
M. Mäki (*)
Program Leader, NAT, Orion Diagnostica Oy,
P.O. Box 83, FI-02101 Espoo, Finland
e-mail: Minna.Maki@mobidiag.com;
Minna.Maki@welho.com
successful under laboratory conditions. Such
failures may occur due to unsuitable culturing
media and conditions for the fungal species in
question. Molecular methods based on detection
of nucleic acid (NA) from clinical samples aim to
circumvent these problems. In addition, they aim
to improve the diagnosis of fungal infections by
shortening time to result and increasing sensitivity and accuracy. Polymerase chain reaction
(PCR)-based assays can amplify and detect minute quantities of DNA isolated from a pathogenic
fungus in few hours, having a limit of detection
of only a few genome copies per reaction.
Although the multiplex PCR is slowly gaining
ground in fungal diagnostics, most of the tests are
still amplifying only one or few fungal targets in
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_53, © Springer Science+Business Media, LLC 2013
535
536
a single reaction [1, 2]. Multiplex or broad-range
PCR in combination with microarray allows rapid
detection of microbial DNA and species
identification of multiple microbial targets in a
single assay [3–7]. The simple, array-type technologies with broad target coverage have especially been believed to have great clinical
potential in the field of infectious diseases [8–12].
Rapid clinical diagnostics reduces the use of antimicrobials in addition to allowing a faster switch
to the most optimum treatment, which improves
patient outcome and reduces both side-effects
and treatment costs [13–17].
Breakspear and Momany have reviewed the
use of fungal microarray in research settings,
including studies of fungal metabolisms, development, pathogenesis, symbiosis, and industrial fermentations [18]. Recently, several publications
have also demonstrated the applicability of fungal
microarray in clinical diagnostic purposes [7, 19–
22]. These publications described the use of a
multiplex/broad-range PCR with oligonucleotide
probe array, targeting highly conserved and variable species-specific regions of the internal transcribed spacers (ITS) of rRNA gene complex of
clinically relevant fungal pathogens. More often
the ITS regions have been chosen as targets in fungal microarrays due to their presence in numerous
copies in the fungal genome, which enables highly
sensitive amplification by PCR. The high level of
sequence variability of the ITS regions also allows
reliable differentiation of closely related fungal
taxa and species. Moreover, the comprehensive
rRNA gene complex database is rapidly expanding and, thus, supporting the in silico design of
primers and oligonucleotide probes.
Three commercial PCR and microarray-based
products for fungal diagnostics stand out from
the rest: CLART® SeptiBac+ (Genomica, Madrid,
Spain), MycArray™ (Myconostica, Manchester,
UK), and Prove-it™ Sepsis, v2.0 (Mobidiag,
Helsinki, Finland). All of these assays use similar
methodologies for detection of pathogenic fungal
species from clinical samples. The assays involve
the use of PCR as an amplification method prior
to microarray phase, where the actual identification
of fungal species occurs. The ArrayTube™ or an
M. Mäki
analog microarray is used as a platform for the
oligonucleotide probes. The ArrayTube™ has
been demonstrated to detect and identify viral
and bacterial pathogens or bacterial pathotypes
with a high degree of sensitivity [23–27] and to
be capable of detecting antimicrobial resistance
genes [28–30] from an isolated DNA sample.
Also, Monecke et al. have published a case report
of peritonitis where the ArrayTube™ harboring
the fungal content was used to detect the causative agent, Rhizopus microspores [31].
In contrast to the previously mentioned fungal
microarray publications, in these three commercial platforms, the principle behind the visualization of a positive hybridization on the microarray
is based on a colorimetric reaction instead of
fluorescent-based methods. In the workflow, biotin labeled amplicons are first hybridized with the
specific oligonucleotide probes pre-printed on
the microarray surface and then streptavidinhorseradish peroxidase (HRP) conjugate is
attached to the biotin label. Finally, the presence
of the HRP is visualized in the precipitation reaction by which HRP catalyzes the conversion of
3,3¢,5,5¢-Tetramethyl Benzidine (TMB) substrate
or an analog into a precipitate thus forming a colored spot on the specific microarray position. An
image is then captured from the microarray by
dedicated reader device. The image is analyzed,
and the result of the analysis, typically the name
of the causative agent and signal intensities of
each oligonucleotide probes, are reported by the
software.
The fungal panels of the CLART SeptiBac+,
MycArray, and Prove-it Sepsis assays vary, but
all of them target the clinically relevant Candida
species, that is, C. albicans, C. krusei, and C.
glabrata. The assays aim at identification of fungal species from the positive blood culture used
in sepsis diagnostics. Sepsis necessitates rapid
and accurate diagnostics to improve the chances
of a positive outcome for the patient. The fungal
sepsis is associated with significant mortality and
morbidity, especially when Candida spp. is the
causative agent. Fluconazole is the choice for
first-line therapy in candidemia; therefore, rapid
differentiation between fluconazole-sensitive and
53
DNA Microarray-Based Detection and Identification of Fungal Specimens
537
Fig. 53.1 The images of the Prove-it™ Sepsis TubeArray reader and Prove-it™ TubeArray, which is a plastic
microreaction tube containing a microarray at the bottom
potentially fluconazole-resistant Candida species
is of the essence. Recent studies have shown that
appropriate and early antifungal therapy (treatment started within the 48 h after the onset of
candidemia) is a major factor associated with a
good prognosis in fungal infection [16, 17].
The performance of Prove-it Sepsis assay in
routine clinical settings for sepsis diagnostics has
been recently published [32]. In the multicenter
study, the definitive identification of bacterial
species with the Prove-it microarray platform
and the corresponding assay protocol was considered highly sensitive (95%) and specific
(99%). It was concluded that the assay was faster
than the gold-standard culture-based methods
and it could thus enable earlier evidence-based
management for clinical sepsis. Furthermore, it
was also stated that the microarray platform’s
robust nature, ease of implementation, softwarecontrolled decision support for results, and
portability has potential for successful strategic implementation in low resource settings
(Fig. 53.1). The current generation of the Prove-it
Sepsis v2.0 assay consists of a pathogen panel
that covers the majority of sepsis-causing pathogens, including over 60 g-negative and grampositive bacterial species, the methicillin
resistance marker together with 13 fungal species. The fungal detection is realized by broadrange PCR primers that originate from the
conserved regions of ITS together with specific
oligonucleotide probes located at hyper-variable
regions flanked by the primers. Each probe on the
array matches either a particular pathogen species or higher-level taxon. The turnaround time
of the assay is three hours, excluding DNA
extraction. The fungal pathogen panel of the
assay covered the following clinically relevant
species: C. albicans, C. glabrata, C. parapsilosis, C. tropicalis, C. guilliermondii, C. lusitaniae,
C. dubliniensis, and C. krusei and pan-yeast
identification covering C. pelliculosa, C. kefyr, C.
norvegensis, C. haemulonii, and Saccharomyces
cerevisiae. The protocol below is based on the
procedure of Prove-it Sepsis StripArray
(Figs. 53.2 and 53.3).
538
M. Mäki
Fig. 53.2 The images of the Prove-it™ Sepsis StripArray system and Prove-it™ StripArray, which consists of eight
plastic microreaction vials containing a microarray at the bottom of each well
Fig. 53.3 The image of the Prove-it™ Advisor result.
The top section of the result view presents the end result
of the assay including icons for the identified target(s) and
the analyzed microarray image. The results view has the
following tabs: Summary, Details, Graphs, Images, and
Panel, from which the details can be viewed. Also, the
Materials
1. Prove-it Sepsis v2.0 kit.
2. Prove-it StripArray System.
detailed result of bacterial content and bacterial controls
(pass/fail), the detailed result of fungal content and fungal
controls (pass/fail), and other assay information (from
Sample ID to Software version) that is common for all
contents are shown
3. Distilled water.
4. Nucleic acid and nuclease-free, aerosolresistant pipette tips.
5. Sterile, nucleic acid-free 1.5-mL microfuge
tubes.
53
DNA Microarray-Based Detection and Identification of Fungal Specimens
6. Sterile, nucleic acid-free PCR tubes suitable
for the PCR instrument.
7. Racks for tubes.
8. Disposable gloves and laboratory coats.
9. PCR Thermal Cycler. The performance of
Prove-it Sepsis has been evaluated using
Eppendorf Mastercycler® epGradient S. The
selection of the PCR cycler instrument may
affect the assay protocol duration and the
assay sensitivity.
10. At least two thermal mixers capable of 25,
30, and 66 °C with microtiter plate adapter.
11. A vortex mixer.
12. A spin microfuge.
13. Adjustable micropipettes for pre-PCR and
post-PCR areas.
14. A vacuum suction system.
Methods
Detection and Identification of Fungal
Species Using PCR- and MicroarrayBased Methods
The protocol of the commercial Prove-it Sepsis
v2.0 assay is modified from the protocol
published by Järvinen et al [3]. and Aittakorpi
et al. [33].
Preparing the Fungal PCR
1. Take PCR reagents, except for polymerase, to
room temperature.
2. Vortex and spin down all reagents.
3. Prepare the fungal master mixture to a clean
laboratory tubes. Add 3.1 mL of PCR water
(Mobidiag), 1.5 mL of 10× Buffer (Qiagen,
Hilden, Germany), 1.1 mL of BSA (Mobidiag),
0.3 mL of MgCl2 (Qiagen), 2.3 mL of dNTPmix (Mobidiag), 0.8 mL of Prove-it Fungi
Primer-F (Mobidiag), and 2.0 mL of Prove-it
Fungi Primer-R (Mobidiag) to the master mixture. Make 10% more of the master mixture
than needed.
4. Add polymerase to the master mixture, 0.4 mL
of HotStarTaq® DNA Polymerase (Qiagen)
per reaction. Always store polymerase at
−20 °C.
539
5. Vortex and spin down the master mixture.
Aliquot it to PCR tubes or strips (11.5 mL of
master mixture per tube).
6. Add 2 mL of the internal PCR control to each
tube. It is not recommended to store or handle
the PCR controls in the same facilities where
the PCR master mixes and primers are handled. The PCR control can be added into the
mixture in the same facilities with the DNA
template.
7. Add 1.5 mL of DNA sample.
8. Place all the tubes/strips to PCR machine and
start PCR program: a denaturation step at
95 °C for 15 min, 36 cycles of 10 s at 96 °C,
35 s at 52 °C, 10 s at 72 °C, 5 cycles of 5 s at
96 °C, 30 s at 65 °C, 5 cycles of 5 s at 96 °C
and finally 30 s at 68 °C.
Preparing Fungal
Hybridization onto Prove-it
Sepsis StripArrays
1. Pick up the number of microarrays and seals
needed.
2. Take all hybridization reagents and distilled
water to room temperature and make sure
that they are equilibrated to RT.
3. Switch on the thermal blocks and make sure
that they are at right temperatures of +30 and
+66 °C. Also check correct agitation speed of
550 rpm.
4. Prepare a fresh hybridization buffer by mixing together the 2× hybridization buffer and
Hybridization buffer diluents (1:2).
5. Prepare a fresh conjugate solution by mixing
together conjugate stock and conjugate
diluent (1:80).
6. Prewashing.
(a) Add 200 mL of distilled water into
microarray wells.
(b) Incubate at +30 °C for 10 min with
550 rpm agitation.
(c) After incubation, carefully remove all
liquid from microarray wells.
7. Hybridization.
(a) Add 97 mL of fresh hybridization buffer
to the microarray wells.
(b) Add into the same microarray well 3 mL
of fungal PCR product.
540
8.
9.
10.
11.
12.
M. Mäki
(c) Incubate at +66 °C for 20 min with
550 rpm agitation.
(d) After incubation, carefully remove all
liquid from microarray wells.
Washing.
(a) Add 200 mL of washing buffer into
microarray wells.
(b) Incubate at +30 °C for 1 min with
550 rpm agitation.
(c) After incubation, carefully remove all
liquid from microarray wells.
Conjugation
(a) Add 100 mL of freshly mixed conjugate
solution into microarray wells.
(b) Incubate at +30 °C for 10 min with
550 rpm agitation.
(c) After incubation, carefully remove all
liquid from microarray wells.
Washing
(a) Add 200 mL of washing buffer into
microarray wells.
(b) Incubate at +30 °C for 5 min with
550 rpm agitation.
(c) After incubation, carefully remove all
liquid from microarray wells.
Precipitation staining
(a) Add 100 mL of substrate into microarray wells.
(b) Incubate at +25 °C for 10 min. NO agitation for this incubation!
(c) After Incubation, carefully remove all
liquid from microarray wells.
Analysis with the Prove-it StripArray reader
and Prove-it Advisor software.
DNA Extraction
A prerequisite for a successful DNA-based analysis of fungal specimens is the efficiency of cell
wall disruption step and subsequent recovery of
fungal DNA without putative PCR inhibitors
originated from the specimen. Hence, the most
appropriate sample preparation and DNA extraction method for any particular application
depends also on the specimen type and quantity
used. Khot and Fredericks have reviewed
both in-house and commercial DNA extraction
methods used with various clinical specimens in
PCR-based fungal diagnostics [1]. A variety of
in-house methods are available. Also, many manufacturers are providing DNA purification kits
that are suitable for preparation of DNA from
fungal specimens [34]. However, when adapting
a protocol to be used in clinical diagnostics, it is
of high importance that the used reagents and
materials are free of fungal bioburden. Any risks
for false-positive result reporting due to the
fungal bioburden should be avoided. Since the
contamination of DNA extraction reagents
with fungal DNA is common [35], the upstream
methods to be used in conjunction with PCR
and microarray-based analysis should always
be evaluated carefully. When blood culture samples is used as a specimen type, it should also be
taken into account that blood culture media contains a common additive polyanetholesulfonate
(SPS), which is a potent inhibitor of PCR and
resistant to removal by some DNA purification
methods [36].
No traces of fungal bioburden have been
observed from the current production versions of
three commercially available DNA extraction
methods. These methods are also efficient regarding the disruption fungal cell wall and the removal
of SPS from positive blood culture samples.
The protocols for automated solution of
NorDiag Arrow (Nordiag, Oslo, Norway) and
Nuclisens®easyMAG® (bioMérieux, Marcy
l’Etoile, France), and manual solution of MycXtra
Fungal DNA Extraction Kit (Myconostica,
Manchester, UK) are introduced below.
Extraction of DNA from Blood Culture
Using NorDiag Arrow
The NorDiag Arrow pipetting instrument is recommended to be used according to the manufacturer’s instructions and recommendations with
the Arrow Viral NA kit and Viral NA v.1.0 program (www.nordiag.com). Shortly, NorDiag
Arrow is an automated extraction instrument for
NAs, using a magnetic bead-based method and
running 1–12 samples simultaneously. Arrow
provides cost-efficient purchasing and running
53
DNA Microarray-Based Detection and Identification of Fungal Specimens
costs. The following protocol is to be used in
conjunction with Prove-it Sepsis assay:
1. Switch the NorDiag Arrow instrument on
and on the select protocol menu, select the
protocol dedicated for Arrow VIRAL NA kit
to be run.
2. Load the instrument with the required consumables, i.e., pumps and pump-tips.
3. Place the cartridge containing the extraction
reagents onto the Arrow rack and place the
Arrow rack to the instrument. Note: The foil
on the cartridges must be peeled off prior to
starting a run.
4. For the DNA eluate, place a clean microcentrifuge tube to the appropriate place in the
Arrow rack.
5. Mix 240 mL of blood culture and 10 mL of
proteinase K in a microcentrifuge tube.
6. Place the sample solution to the appropriate
place in the Arrow rack.
7. From the protocol touch screen, choose the
sample input volume of 250 mL.
8. From the protocol touch screen, choose the
sample elution of 100 mL.
9. Start the protocol. The run is carried out
automatically.
10. Run is finished within ~50 min, after which
DNA is ready to be used in PCR
applications.
541
from the silica, after which it is ready to be used
in PCR applications. The following protocol is to
be used in conjunction with the Prove-it Sepsis
assay:
1. Switch the NucliSENS easyMAG instrument
on and select the protocol Generic 2.0.1 to
be run.
2. Start the off-board lysis extraction protocol
Generic 2.0.1 and set the sample material;
i.e., blood culture media and the elution volume of 55 mL www.biomerieux.com.
3. Add 100 mL of blood culture to 2 mL of
NucliSENS Lysis buffer. Vortex thoroughly.
Incubate 10 min at room temperature.
4. Insert aspiration tip sets into the instrument.
5. Load the lysed sample into the 1 well of the
8-well sample vessel.
6. Mix 550 mL of distilled water and 550 mL
magnetic silica together.
7. Add 100 mL of silica mixture to the well
of the sample vessel containing the lysed
sample and mix thoroughly.
8. Insert the vessel into the instrument.
9. Start the run which is carried out automatically. The instrument automatically verifies
if there are sufficient amount of the on-board
reagents.
10. After the run of 40 min, the extraction protocol is completed and the eluted DNA can be
moved from the vessel to a clean laboratory
tube for the use in PCR applications.
Extraction of DNA from Blood
Culture Using NucliSENS easyMAG
NucliSENS® easyMAG® instrument is recommended to be used according to the manufacturer’s instructions and recommendations (www.
biomerieux.com). NucliSENSeasyMAG is an
automated system for total nucleic acid extraction from a variety of sample types and volumes,
capable of running 1–24 samples simultaneously.
NA extraction method is based on the magnetic
silica particles. The target NAs bind to the magnetic silica particles during the incubation of
lysed sample. The magnetic device is then introduced to the silica particles, enabling the system
to purify the NAs trough several washing steps.
After washing, the heating step releases DNA
Extraction of DNA from Clinical
Specimen Using MycXtra Fungal
DNA Extraction Kit
MycXtra kit is recommended to be used according to the manufacturer’s instructions and recommendations
(www.myconostica.com).
The
principle of the kit is to lyse the fungal cells in the
sample by combining the use of a detergent and a
mechanical force against specialized beads. The
cellular components are lysed by a mechanical
action on a vortex. From the lysed cells, the
released DNA is bound to a silica spin filter.
The filter is washed and DNA is harvested in a
buffered solution. The protocol is manual.
542
1. Centrifuge the sample for 20 min at 3,000 × g.
Decant the supernatant and retain it.
2. Resuspend the pellet in 800 mL of the retained
supernatant and transfer it to microcentrifuge
tube.
3. Centrifuge for 2 min at 10,000 × g and discard
the supernatant. Resuspend the pellet in the
solution remaining in the tube and transfer the
entire amount to a 2-mL Bead Solution tube.
4. Gently invert to mix.
5. Add 60 mL of Solution S1 to the Bead
Solution tube and invert several times.
6. Add 200 mL of Solution IRS to the Bead
Solution tube.
7. Vortex at maximum speed for 10 min.
8. Centrifuge the Bead Soltuin tube at 10,000 × g
for 30 s.
9. Transfer 450 mL of supernatant to a clean
microcentrifuge tube taking care not to disturb
the beads. Discard the Bead Solution tube.
10. Add 250 mL of Solution S2 to the supernatant and vortex for 5 s. Incubate at 4–8 °C for
5 min.
11. Centrifuge tubes for 1 min at 10,000 × g.
12. Avoiding the pellet, transfer the entire volume of the supernatant to a clean microcentrifuge tube.
13. Add 1.1 mL of Solution S3 to the supernatant. Mix by inverting.
14. Load approximately 650 mL on to a spin filter
and centrifuge at 10,000 × g for 30 s. Discard
the flow trough. Repeat this step until all
supernatant has passed through the spin
filter.
15. Add 300 mL of Solution S4 to the spin filter
and centrifuge for 30 s at 10,000 × g.
16. Discard the flow through.
17. Centrifuge again for 1 min to remove the last
traces of S4, which will inhibit the PCR
reaction.
18. Carefully place spin filter in a new microcentrifuge tube and add 40 mL of Solution S5 to
the center of the white spin filter membrane.
Leave at room temperature for 2 min.
19. Centrifuge for 30 s at 10,000 × g.
20. Discard the spin filter.
21. DNA in the tube is now ready for use in a
PCR application.
M. Mäki
Notes
1. Specimens should always be considered as
potentially infectious.
2. Store and extract DNA from a specimen separately from the reagents and the pre-PCR area.
3. The procedure should be performed in physically separated areas (pre-PCR area and postPCR area) to avoid contamination with
microbial organisms or nucleic acids or any
other extraneous material or agents; e.g., amplicons from previous PCR runs. In the pre-PCR
area, the preparation of the PCR mixture should
be conducted in an area separate from where
the addition of the DNA sample takes place.
4. Each pre/post-PCR area should have its own
dedicated working materials assigned; e.g.,
pipettes, spin microfuge, and disposable
gloves. Any material in the post-PCR area
must never come into contact with that of the
pre-PCR area.
5. Always follow the unidirectional workflow
from the pre-PCR area to the post-PCR area.
Never reverse the direction.
6. Always wear suitable protective clothing and
gloves during the procedure.
7. Follow the recommendation of the manufacturer of thermal cycler regarding to the quality of PCR plastic ware.
8. Avoid scratching the microarrays in the
bottom.
9. Avoid bubble formation on the microarray
surface. Reverse pipetting is recommended
to avoid bubble formation.
10. Keep the microarray bottom clean at all times
to avoid any interference when detecting the
assay result.
11. Be careful not to let the microarray wells dry
for longer than necessary between the hybridization steps.
12. TMB-based substrate must be protected from
light.
13. Proceed to the PCR step immediately after
the DNA extraction step. Also, proceed to the
microarray step after the PCR step. Storing
the DNA extract or PCR product may affect
the assay result.
53
DNA Microarray-Based Detection and Identification of Fungal Specimens
Acknowledgments I thank my colleagues in Mobidiag,
especially Anne Aittakorpi, for her constructive comments
on the manuscript.
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Bioinformatic Protocols
and the Knowledge-Base
for Secretomes in Fungi
54
Gengkon Lum and Xiang Jia Min
Abstract
Fungal secreted proteins play important roles in cell signaling, metabolism, and regulation of fungal growth and development. The secretome
refers to all secreted proteins in a proteome that are identified from completely sequenced genomes. The majority of secreted proteins are classical, signal peptide-dependent proteins that can be predicted using
bioinformatics tools. In this chapter, we describe some commonly used
tools for secreted protein prediction in fungi and propose a relatively accurate bioinformatic protocol for fungal secretome identification. The protocol combines multiple signal peptide or subcellular location predictors,
including SignalP, WoLF PSORT, and Phobius, with TMHMM for removing transmembrane proteins and PROSITE PS-Scan for removing endoplasmic reticulum (ER) proteins. Applying this protocol, we have built the
fungal secretome knowledge-base (FunSecKB). The utility of FunSecKB
is described in detail. FunSecKB serves the community as a central portal
for search and deposition of fungal secretome information.
Keywords
Secreted proteins • Secretome • Signal peptide • Fungi • Prediction
• Knowledge-base • Database
Introduction
G. Lum
Department of Computer Science and Information
Systems, Youngstown State University,
Youngstown, OH 44555, USA
X.J. Min (*)
Department of Biological Sciences, Center for Applied
Chemical Biology, Youngstown State University,
One University Plaza, Youngstown, OH 44555, USA
e-mail: xmin@ysu.edu
Secreted proteins are proteins which are synthesized within cells and then secreted to extracellular
space and matrix to play their roles. Secreted proteins play important roles in cell signaling, metabolism, and regulation in growth and development
of all organisms. As the genomes have been completely sequenced in many organisms, the proteomes could be predicted using the information in
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_54, © Springer Science+Business Media, LLC 2013
545
546
genomes. The term “secretome” was first used to
include all proteins secreted to extracellular space
and matrix and proteins involved in the secretion
pathway including endoplasmic reticulum, Golgi
apparatus, and transportation vesicles [1–3]; however, more recently, the term was used to include
secreted proteins only [4,5]. In this work, the
secretome only refers the complete set of secreted
proteins in an organism.
Secretomes are an important part of the fungal
proteome. These secreted proteins include
enzymes, growth factors, cell wall proteins, and
other bioactive molecules which play important
roles in host–pathogen interactions. Fungal
secreted enzymes are used to break down potential food sources for transport into the cells. As
there are many types of fungi producing a great
variety of enzymes that are able to break down
lignocelluloses and other biopolymers, fungi
have an important function in the biosphere as
decomposers. Since secreted proteins are useful
in their ability to break down biopolymers, they
have found a role in many applications including
pharmaceutical and industrial [6]. Therefore, the
ability to analyze a protein to determine if it is
secreted and what functions it may have is useful
as a tool in research to more easily focus on or to
eliminate potential targets. Increased understanding of the secretome biology of fungi will further
promote exploration of the potential applications
of fungal secreted proteins in environmental
remediation and industrial processing including
bio-fuel production.
Most of secreted proteins in fungi are classical
secreted proteins, which have a signal peptide on
the N-terminus of protein sequences. A signal
peptide is typically 15–30 amino acids long,
located at the N-terminus of the protein and is
cleaved off during translocation across the membrane. The presence of a signal peptide directs
the protein to the rough endoplasmic reticulum
(ER) and the Golgi complex in preparation for
transport through the secretory vesicles. This is
referred to as the classical secretory pathway.
Although not all proteins excreted extracellularly
contain a signal peptide, it is believed that that
the majority of fungal proteins are secreted in this
manner [6]. The presence of a transmembrane
G. Lum and X.J. Min
domain in the protein sequence, however, indicates that although the protein passes through the
classical secretory pathway, it is not secreted
extracellularly but instead becomes part of the
cell membrane. By combining the results of one
or more predictions for the presence of a signal
peptide along with the absence of a transmembrane domain, the likelihood of the protein being
secreted is very high. Our recent evaluation
reveals that combining the results of multiple
programs increases the accuracy by reducing the
number of false positives and negatives [7].
Two fungal-specific secretome databases are
currently available. The Fungal Secretome
Database1 developed by Choi et al. used nine bioinformatics tools and protein sequences from
completely sequenced genomes including some
work in progress draft genomes [8]. The Fungal
Secretome Knowledge-Base (FunSecKB)2 developed by us used all fungal protein sequences
available in the NCBI RefSeq database and being
linked and supplemented with protein sequences
in the UniProt database. The detailed comparison
of the two databases was described by Lum and
Min [5]. In this work, we focus on how to utilize
FunSecKB.
Materials (Data)
For individual secreted protein identification, the
input is a fungal protein sequence in FASTA format. For a species-specific secretome prediction
from a whole proteome, which often is obtained
from a completely sequenced genome, a set of
proteins in multiple FASTA format are used as
input.3 We will use a glucoamylase enzyme from
Aspergillus niger (gi 145235763) and a
Schizosaccharomyces pombe protein (gi 19115161)
as examples to explain the input and output of the
tools mentioned in Sect. “Methods.”
http://fsd.snu.ac.kr/.
http://proteomics.ysu.edu/secretomes/fungi.php.
3
A description of the FASTA format may be found at
http://www.ncbi.nlm.nih.gov/BLAST/fasta.shtml.
1
2
54
Bioinformatic Protocols and the Knowledge-Base for Secretomes in Fungi
Methods
The programs used for fungal secretome prediction include SignalP 3.0 [9], TargetP 1.1 [10],
TMHMM 2.0 [11], Phobius [12], WoLF PSORT
[13], PS-Scan for PROSITE [14], and FragAnchor
[15]. SignalP and TargetP predict the presence
and location of an N signal peptide and a potential cleavage site. TMHMM predicts the presence
of a transmembrane domain. Phobius is a combined signal peptide and transmembrane topology predictor. WoLF PSORT (WolfPsort) predicts
the subcellular location(s) of a protein. PS-Scan
is a PROSITE scanning tool which predicts
whether or not a protein contains an endoplasmic
reticulum (ER) targeting sequence (Prosite:
PS00014). FragAnchor is used to predict if there
is a glycosylphosphatidylinositol (GPI) anchor in
the protein, which may indicate if the secreted
protein is a cell wall protein or attaches to the
outside of the plasma membrane.
There are two methods of using these tools.
The first one, most often used by a biologist to
process an individual protein, uses the online
Webserver tool. The second one, often used by
bioinformaticians to process proteome-wide
secretome identification, use a standalone package which may be downloaded and run on a
UNIX (Linux) platform.
547
In the Signal-NN results, two different neural
networks are used for each prediction, one for predicting the presence of the signal peptide, the other
for predicting the position of the cleavage site. For
each position in the protein, a C, S, and Y score is
calculated. The C score is the cleavage site score
with values being high at potential cleavage sites.
The S score is reported for every position submitted with high scores for amino acids which are
part of the signal peptide and low score for those
which are part of a mature protein. The Y score is
a derivative of C and S with a likely cleavage
point being when the slope of S is steep and there
is a high C score resulting in a high Y score. The
mean S score is the average of the S scores from
the N-terminus to the highest Y score. The D score
is average of the Y score and the mean S score.
The D score is used to determine whether or not a
protein is predicted to be secreted.
In the Signal-HMM results, the positions are
evaluated to determine the likelihood of being a
part of the n-region, h-region, or c-region. Signal
peptides commonly have a hydrophobic central
core (h-region) surrounded by the N- and
C-terminal hydrophilic regions. The HMM makes
a prediction of a signal peptide, a nonsecretory
protein or a signal anchor. A protein with a signal
anchor passes through the membrane but the
uncleaved signal peptide remains anchored to the
membrane resulting in a type II membrane protein [9]. The results also include a probability for
both a signal peptide and a signal anchor.
SignalP 3.0
SignalP 3.0 uses both neural network (NN) and
hidden Markov model (HMM) algorithms in two
different predictors to predict whether a protein
has a signal peptide and where the most likely
cleavage site would be if one is detected [9].4 For
each protein processed by SignalP 3.0, scores are
calculated and returned in two sections:
SignalP-NN result (Fig. 54.1a) and SignalPHMM result (Fig. 54.1b).
4
http://www.cbs.dtu.dk/services/SignalP/.
Phobius
Phobius is a combined signal peptide and a transmembrane topology predictor 5 [12]. A known
problem with signal peptide and transmembrane
topology predictors is the high similarity of the
hydrophobic regions of both the signal peptide
h-region and transmembrane helix. Due to this
similarity, pure signal peptide predictors and
transmembrane topology predictors sometimes
5
http://phobius.sbc.su.se/index.html.
548
G. Lum and X.J. Min
Fig. 54.1 (a) Neural network results output for SignalP 3.0 Server of Aspergillus niger glucoamylase protein.
(b) Hidden Markov model results output for SignalP 3.0 Server of Aspergillus niger glucoamylase protein
54
Bioinformatic Protocols and the Knowledge-Base for Secretomes in Fungi
549
Fig. 54.2 Results output for Phobius of Aspergillus niger glucoamylase protein
results in false classifications. To this end,
Phobius was designed to do both signal peptide
and transmembrane topology prediction and to
distinguish between the two regions.
The output formats available are long with
graphics, long without graphics, or short format
(Fig. 54.2). The default output (long with graphics) shows the prediction of probable locations
for sections of the protein. Some possible predictions are: SIGNAL for signal peptide, REGION
for N-, H-, and C-regions, TOPO_DOM for
topology (cytoplasmic or non-cytoplasmic) and
TRANSMEM for positions predicted to be within
the membrane. The range of positions is given for
each predicted segment. If the entire sequence is
labeled cytoplasmic or non-cytoplasmic though,
the prediction is that there are no membrane helices and is not an actual prediction of location, but
the most probable location.
The short output format gives TM as the number of predicted transmembrane segments, SP as
the prediction of whether or not there is a signal
peptide, and PREDICTION as the predicted
topology. The format of the predicted topology is
given as a series of numbers and letters. If a signal peptide is detected, it is given in the format:
n#-#c#/# where # represents a position in the
sequence. The numbers between n and c is the
range of the hydrophobic h-region and the #/# is
the cleavage site. Following this is either an “i” if
the loop is cytoplasmic or an “o” if it is on the
non-cytoplasmic side and then numbers in the
550
G. Lum and X.J. Min
Fig. 54.3 Results output for WoLF PSORT of Aspergillus niger glucoamylase protein
Fig. 54.4 Results output for TargetP 1.1 Server of Aspergillus niger glucoamylase protein
format #-# indicating the range of the transmembrane helix. This format is repeated until the end
of the sequence.
WoLF PSORT
WoLF PSORT6 is a program for predicting the
subcellular location of proteins [13]. It takes the
amino acid sequences and converts them into
numerical vectors which are then classified using
a weighted k-nearest neighbor classifier. The predictions are based on known sorting signal motifs
and the content of the amino acids. It requires the
selection of the organism type: animal, plant, or
fungi. For our example protein (Fig. 54.3), the
result was based on using k = 27 nearest neighbors. Of these 27 closest, 26 were extracellular
and the result is displayed as extr: 26.0. A list of
the localization site definitions is available on the
Website and include locations such as golg for
the Golgi apparatus, mito for mitochondria, and
nucl for nuclear.
TargetP 1.1
TargetP is designed to predict the subcellular locations of eukaryotic proteins7 [10]. TargetP predicts
in the N-terminus the presence of any of the
N-terminal presequences such as signal peptide
(SP), chloroplast transit peptide (cTP), or mitochondrial targeting peptide (mTP). The output is
given in Fig. 54.4. Name is the sequence name
truncated to 20 characters. Len is the length of the
sequence. cTP, mTP, SP, other are the final neural network (NN) scores. cTP is only used if the
organism group on the submission page is set to
Plant since it is used to detect a cTP.
TMHMM 2.0
TMHMM 2.0 uses a HMM to predict the presence and topology of transmembrane helices and
their orientation to the membrane (in/out)8[11].
The output shows the results of the prediction
(Fig. 54.5).
7
6
http://wolfpsort.org/
8
http://www.cbs.dtu.dk/services/TargetP/.
http://www.cbs.dtu.dk/services/TMHMM/.
54
Bioinformatic Protocols and the Knowledge-Base for Secretomes in Fungi
551
Fig. 54.5 Results output for TMHMM Server 2.0 of a Schizosaccharomyces pombe protein
Using the default long format: Length is the
length of the sequence submitted. Number of
predicted TMHs is the number of predicted
transmembrane helices. Exp number of AAs in
TMHs is the expected number of amino acids in
transmembrane helices. Exp number, first 60
AAs is the expected number of amino acids in
transmembrane helices within the first 60 positions. Total prob of N-in is the total probability
that the N-terminus is on the cytoplasmic side of
the membrane. Following this section is the prediction of where specific parts of the protein are
likely to be: inside, outside, or TM helix (part of
the transmembrane helix). The structure is the
identifier used, followed by the program name
(TMHMM2.0), the predicted location, then the
starting and ending position of the segment. In
this example, the prediction is that the N-terminus
is outside the membrane and the protein crosses
the membrane seven times and the C-terminus
ends on the inside of the cell. Using the short format: len, length of sequence, ExpAA, expected
number of amino acids in transmembrane helices, First60, expected number of amino acids in
transmembrane helices within the first 60 positions, PredHel, number of predicted transmembrane helices by N-best, and Topology, the
topology predicted by N-best with “o” indicating
sections outside and “i” indicating sections inside
the cell.
552
G. Lum and X.J. Min
Fig. 54.6 Results output for ScanProsite of a Schizosaccharomyces pombe protein
PS-Scan for PROSITE
PROSITE is a database containing protein families, domains, and functional sites. The
ScanProsite Website9 scans the PROSITE database for motifs matching the input sequence [14].
The output from the Website lists any hits found
in their database matching sections within our
sequence. In our FunSecKB database, we used
the standalone program PS-Scan to determine if
there was an ER retention signal (Prosite:
PS00014), which if found could rule out the possibility that the particular protein would be
secreted. The output of a Schizosaccharomyces
pombe protein (gi 19115161) shows an ER targeting sequence detected (Fig. 54.6) at positions
321–324.
potential GPI-anchored sequences and a HMM to
classify those sequences into categories of likelihood. The four categories are highly probable,
probable, weakly probable, and potential false
positive. Our example did not contain a potential
GPI-anchored sequence and thus was rejected
with the HMM classification never being run,
thus the output is not shown here. However, the
detailed information of the GPI-anchored secreted
proteins and the correlations with proteome size
and genome size can be found in Lum and Min
[5]. This Webserver support a batch of sequences,
but no standalone tool is available. There are
some other tools available for GPI anchor prediction, including Big-PI predictor 11 and PredGPI.12
SecretomeP
FragAnchor
FragAnchor10 is a tool to detect the presence of a
glycosylphosphatidylinositol (GPI) anchor [15].
It uses a combination of a neural network to select
SecretomeP13 is a program that uses a sequencebased method for prediction of secreted proteins based on nonclassical secretory pathways.
The original program was trained on bacteria
and support for mammalian proteins was added
http://mendel.imp.ac.at/sat/gpi/gpi_server.html.
http://gpcr.biocomp.unibo.it/predgpi/.
13
http://www.cbs.dtu.dk/services/SecretomeP/.
9
11
10
12
http://expasy.org/tools/scanprosite/.
http://navet.ics.hawaii.edu/~fraganchor/NNHMM/
NNHMM.html.
54
Bioinformatic Protocols and the Knowledge-Base for Secretomes in Fungi
Table 54.1 Linux
commandline summary
for standalone packagesa
Tools
SignalP
Phobius
WoLFPsort
TargetP
TMHMM
PS-Scan
553
Commands
signalp –t euk –f summary input_file>output_file
phobius input_file –short>output_file
runWolfPsortSummaryOnly fungi<input_file>output_file
targetp –c –N input_file>output_file
tmhmm input_file –short –noplot>output_file
ps_scan.pl input_file –p PS00014 –o scan –d prosite.dat>output_file
Input_file is the protein sequences in FASTA format. output_file is the file to save the
results of the program
a
afterward. The Webserver currently has support
for gram-negative and gram-positive bacteria
along with mammalian proteins but its use in prediction of secreted fungal proteins by nonclassical pathways has not been tested. Choi et al. used
this tool to predict nonclassical, signal peptide
independent secreted proteins in constructing
the Fungal Secretome Database14 [8]. However,
as the accuracy of the tool in fungal secretome
prediction was not reported, we did not use this
tool in FunSecKB (see discussion in Sect.
“TMHMM 2.0”).
Commands of Standalone Tools
We described the online Webservers above. The
online Webservers normally have a limit for
the maximum number of sequences allowed to be
submitted at once; therefore, to process a large
number (i.e., a proteome of a whole species)
the standalone tools are needed. For the standalone tools that need to be installed on a Linux
system, the commands of how to run them are
summarized in Table 54.1. Detailed explanations
of how to run each tool often can be found in the
“readme” page in each downloaded package.
Protocol Evaluation
The accuracy of a prediction tool can only be
evaluated using a set of sequence data. Min
reported the accuracy of some of the tools mentioned above in prediction of fungal secretomes
[7]. The tools were evaluated individually and in
combination with others. The dataset contained
241 secreted proteins and 5,992 nonsecreted proteins and the results were measured using sensitivity (Sn) (Equation 54.1), specificity (Sp)
(Equation 54.2), and Mathews’ Correlation
Coefficient (MCC) (Equation 54.3) [16–18],
Sn (% ) = TP / (TP + FN )× 100
(54.1)
Sp (% ) = TN / (TN + FP )× 100
(54.2)
1/ 2
MCC (% ) = (TP × TN − −FP × FN )× 100 / ((TP + FP )(TP + FN )(TN + FP )(TN + FN ))
where TP represents the number of true positives,
FN is the number of false negatives, TN is the
number of true negatives, and FP is the number
of false positives. When tools were combined, a
true positive was counted only when all the tools
http://fsd.snu.ac.kr/.
14
(54.3)
used predicted the protein as positive. The results
were provided in Table 54.2, which was adopted
from Ref. [7]. Based on the results, we used the
combination of SignalP, Phobius, WoLF PSORT,
TMHMM, and PS-Scan, which gave the highest
MCC (83.4 %) result, as the prediction protocol
for fungal secretome prediction in FunSecKB
development [5]. The TargetP 1.1 can be used for
554
G. Lum and X.J. Min
Table 54.2 Prediction accuracies of secreted proteins in fungia
Methods
SignalP
Phobius
TargetP
WolfPsort
SignalP/TMHMM
Phobius/TMHMM
TargetP/TMHMM
WolfPsort/TMHMM
SignalP/TMHMM/WolfPsort
SignalP/TMHMM//WolfPsort/Phobius
SignalP/TMHMM/WolfPsort/Phobius/PS-Scan
SignalP/TMHMM/WolfPsort/Phobius/TargetP/PS-Scan
TP
232
226
228
230
228
224
224
227
226
222
222
218
FP
329
203
583
167
168
200
265
135
86
69
67
66
TN
5663
5789
5409
5825
5824
5792
5727
5857
5906
5923
5925
5926
FN
9
15
13
11
13
17
17
14
15
19
19
23
Sn (%)
96.3
93.8
94.6
95.4
94.6
92.9
92.9
94.2
93.8
92.1
92.1
90.5
Sp (%)
94.5
96.6
90.3
97.2
97.2
96.7
95.6
97.7
98.6
98.8
98.9
98.9
MCC (%)
61.2
68.8
48.6
73.1
72.6
68.6
63.5
75.8
81.6
83.1
83.4
82.6
TP true positives; FP, false positives; TN, true negatives; FN, false negatives; Sn, sensitivity; Sp, specificity; MCC,
Mathews’ correlation coefficient.
a
The table is reproduced with permission from Min [7].
individual secreted protein prediction, however,
adding it to the pipeline for secretome prediction
slightly reduced the accuracy (see Table 54.2).
The Fungal Secretome
Knowledge-Base
The Fungal Secretome Knowledge-Base
(FunSecKB) is a database of fungal proteins collected from NCBI and UniProt on which we have
performed various analyses for prediction of possible extracellular secretion [5].15 From this site
(Fig. 54.7), you can look up specific proteins
using either NCBI’s gi or RefSeq accession or
UniProt’s accession numbers. In addition you can
enter a keyword to search for such as species,
function, or cellular location. You may also search
for secreted proteins of a specific species or perform BLAST (Basic Local Alignment Search
Tool) search against our fungal database. When a
keyword or species secretome search is performed, a list of results will be displayed with an
identifier to the left followed by a description.
The identifier is a link and clicking on it will display the details page for that protein. Similarly,
It is an online resource available at http://proteomics.
ysu.edu/secretomes/fungi.php.
15
searching for a specific protein by gi or accession
will display that particular protein. This page
shows the results of the different tests performed
on the protein along with the sequence in FASTA
format and any available manually curated data.
The Web page is divided up into five main sections: Search individual proteins by ID or
keyword(s), Search secretome information by species, BLAST search, and Community Annotation.
Search by ID or Key Words
This section allows searching for a specific protein by using NCBI’s RefSeq accession or gi
number or UniprotKB’s accession number.
A search by keyword(s) will return a list of proteins containing the keyword(s) based on the
UniProt Protein name. Details of an individual
protein’s results may be found by clicking on the
identifier. For each protein which has been tested
in our database, the results of those tests are displayed. The first area includes the various
identifiers from NCBI and UniProt along with a
clickable direct link to those sites. Also listed are
the species, RefSeq definition, UniProt name,
and a UniProt annotation for subcellular location
(if any). The second area is a summary of the test
results consisting of a yes/no for prediction of
a secreted protein for each test. Also listed is a
54
Bioinformatic Protocols and the Knowledge-Base for Secretomes in Fungi
555
Fig. 54.7 Home page of Fungal Secretome Knowledge-Base
conclusion of whether or not this protein is
belonged to a Secretome: based on our own combination prediction algorithm as mentioned
above, that is, SignalP/Phobius/WoLF PSORT
predicted to have a signal peptide, TMHMM predicted not have a transmembrane domain, and
PS-Scan did not find an ER retention signal. The
third area is the details for each of the tests along
with a link to the original site’s page on how to
interpret the results (if available) or the Web site
for the program. After the test results is listed
the protein sequence used in FASTA format and
if manual curation was done for the particular
protein, the experimental evidence and the
PubMed reference to the paper is given.
Search or Download Secretomes
by Species
This section allows searching by species of
secreted proteins, which are either predicted or
curated. You can either select from a drop-down
menu one of 53 species or manually input a species to search for. When using the drop-down
menu, you may also select a protein set, either
556
Complete Secretome or Curated Proteins. The
complete secretome is all proteins in the species
predicted or curated by UniProt or our curator to
be secretomes. From these options you may either
search or download the FASTA. Search gives a
listing of proteins similar to searching for keyword as above where an individual protein may
be clicked to view details. FASTA download
allows you to download the FASTA for a particular species. When “Curated Proteins” is selected,
a list of available proteins is given on a Web page
which may be copied and pasted. When “Complete
Secretomes” is selected a window will appear
allowing you to download and save a “.fas” file
for that species since usually the entire FASTA
file would be too large to display on screen.
BLAST Search
This section allows either a BLASTP or a
BLASTX search against either of our two fungal
databases. One is the secretome database containing our predicted and confirmed proteins and
the other is for all fungal proteins in our database.
The input format is a sequence or file in FASTA
format. The NCBI BLAST page provides more
information about how to use BLAST.16
Community Annotation
This section is a Community Annotation submission page allowing the user community to submit
a protein for manual curation and addition into
our database. The required entries are email,
RefSeq gi and accession numbers, subcellular
location of the protein, evidence and reference
for the submission. Entries will be curated and if
confirmed, entered into our database. Currently,
we have manually curated secreted proteins from
Aspergillus niger based on Tsang et al. [19] and
A. oryzae based on Oda et al. [20] We would like
to request the fungal secretome research community to submit experimentally verified secreted
16
http://blast.ncbi.nlm.nih.gov/Blast.cgi.
G. Lum and X.J. Min
fungal proteins to FunSecKB using this utility.
Once a protein has been curated, it will be permanently included as part of the database.
Acknowledgments We thank Dr. Gary Walker for his
mentoring support and Jessica Orr for assistance in manual data curation. The work is supported by Youngstown
State University (YSU) Research Council (Grants 200910 #04-10 and 2010-2011 #12-11), YSU Research
Professorship (2009–2011), and the College of Science,
Technology, Engineering, and Mathematics Dean’s reassigned time for research to XJM.
References
1. Tjalsma H, Bolhuis A, Jongbloed JD, Bron S, van Dijl
JM (2000) Signal peptide-dependent protein transport
in Bacillus subtilis: a genome-based survey of the
secretome. Microbiol Mol Biol Rev 64:515–547
2. Greenbaum D, Luscombe NM, Jansen R, Gerstein M
(2001) Interrelating different types of genomic data,
from proteome to secretome: ‘oming in on function.
Genome Res 11:1463–1468
3. Hathout Y (2007) Approaches to the study of the cell
secretome. Expert Rev Proteomics 4:239–248
4. Simpson JC, Mateos A, Pepperkok R (2007) Maturation
of the mammalian secretome. Genome Biol 8:211
5. Lum, G. and Min, X. J. (2011) FunSecKB: the Fungal
Secretome KnowledgeBase. Database 2011:
doi:10.1093/database/bar001.
6. O’Toole N, Min XJ, Storms R, Butler G, Tsang A
(2006) Sequence-based analysis of fungal secretomes.
Appl Mycol Biotechnol 6:277–296
7. Min XJ (2010) Evaluation of computational methods
for secreted protein prediction in different eukaryotes.
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8. Choi J, Park J, Kim D et al (2010) Fungal secretome
database: integrated platform for annotation of fungal
secretomes. BMC Genomics 11:105
9. Bendtsen JD, von Nielsen H, Heijne G, Brunak S
(2004) Improved prediction of signal peptides:
SignalP 3.0. J Mol Biol 340:783–795
10. Emanuelsson O, Nielsen H, Brunak S, von Heijne G
(2000) Predicting subcellular localization of proteins
based on their N-terminal amino acid sequence. J Mol
Biol 30:1005–1016
11. Krogh A, Larsson B, von Heijne G, Sonnhammer
ELL (2001) Predicting transmembrane protein topology with a hidden Markov model: application to complete genomes. J Mol Biol 305:567–580
12. Käll L, Krogh A, Sonnhammer ELL (2004) A combined transmembrane topology and signal peptide
prediction method. J Mol Biol 338:1027–1036
13. Horton P, Park KJ, Obayashi T, Fujita N, Harada H,
Adams-Collier CJ et al (2007) WoLF PSORT: protein
localization
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35:W585–W587 (Web Server issue)
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14. Sigrist CJA, Cerutti L, de Casro E, LangendijkGenevaux PS, Bulliard V, Bairoch A et al (2010)
PROSITE, a protein domain database for functional
characterization and annotation. Nucleic Acids Res
38:161–166
15. Poisson G, Chauve C, Chen X, Bergeron A (2007)
FragAnchor a large scale all Eukaryota predictor of
glycosylphosphatidylinositol-anchor
in
protein
sequences by qualitative scoring. Genomics
Proteomics Bioinformatics 5:121–130
16. Matthews BW (1975) Comparison of the predicted
and observed secondary structure of T4 phage
lysozyme. Biochim Biophys Acta 405:442–451
17. Baldi P, Brunak S, Chauvin Y, Andersen CA,
Nielsen H (2000) Assessing the accuracy of
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18. Menne KM, Hermjakob H, Apweiler R (2000) A comparison of signal sequence prediction methods using a
test set of signal peptides. Bioinformatics
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19. Tsang A, Butler G, Powlowski J et al (2009)
Analytical and computational approaches to define
the Aspergillus niger secretome. Fungal Genet Biol
46:S153–S160
20. Oda K, Kakizono D, Yamada O, Iefuji H, Akita O,
Iwashita K (2006) Proteomic analysis of extracellular
proteins from Aspergillus oryzae grown under submerged and solid-state culture conditions. Appl
Environ Microbiol 72:3448–3457
High-Throughput Functional
Annotation and Data Mining
of Fungal Genomes to Identify
Therapeutic Targets
55
Gagan Garg and Shoba Ranganathan
Abstract
With the advent of next-generation sequencing approaches and mass
spectrometry techniques, there is a huge explosion in nucleotide and protein
sequence data. Despite this increase in sequence data, several proteins remain
unannotated, such as hypothetical proteins. Annotation and extraction of
secretory proteins from the proteome using labor-intensive wet-lab techniques is prohibitive. Computational tools can be used to provide putative
functionality, prior to experimental validation. This chapter introduces a bioinformatics workflow system using the best currently available free computational tools for the annotation of hypothetical proteins and prediction and
analysis of secreted proteins as therapeutic targets, applied to pathogenic
fungi, Cryptococcus gattii, and Cryptococcus neoformans var. grubii.
Keywords
Annotation • Drug targets • Interproscan • Protein domains • BRITE
• FASTA • KEGG • KAAS • SPAAN
Introduction
G. Garg
Department of Chemistry and Biomolecular Sciences,
Macquarie University, Sydney, NSW 2109, Australia
S. Ranganathan (*)
Department of Chemistry and Biomolecular Sciences,
Macquarie University, Sydney, NSW 2109, Australia
Department of Biochemistry, Yong Loo Lin School
of Medicine, National University of Singapore,
8 Medical Drive, Singapore 117597, Singapore
e-mail: shoba.ranganathan@mq.edu.au
The proteome is the entire set of proteins
expressed by an organism. It is a valuable tool for
studying molecular function, development and
progression of different life stages, and much
more. With many fungal genomes sequenced or
under sequencing led to the identification of
whole genome and protein sequences. Usually
genes are predicted by using gene prediction
tools followed by prediction of coding regions.
These putative proteins are annotated based on
the similarity of other organisms’ proteins in the
same taxonomic class. Many proteins still remain
unannotated using these practices. This chapter
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_55, © Springer Science+Business Media, LLC 2013
559
560
outlines an approach to functionally annotate
putative proteins in terms of pathways, gene
ontology, and protein domains from recently
sequenced fungal genomes, followed by secretory peptides and therapeutic target prediction.
Materials
Data
1. Hypothetical protein or nucleotide fungal
sequences to be annotated are obtained from
NCBI1 or from other fungal genomes sources
in FASTA format. We downloaded a total of
6,210 proteins for Cryptococcus gattii
(Serotype B) [1] and 6,967 proteins for
Cryptococcus neoformans var. grubii
(Serotype A) [2].
2. Mouse and human proteins2 are obtained from
NCBI proteins database.
3. Known drug targets dataset3 are obtained from
DrugBank [3, 4].
G. Garg and S. Ranganathan
6. Interproscan9 [10] for protein domains
mapping.
7. BLASTP10 [11] for sequence similarity search
of the query sequence against different
datasets.
Secretory Proteins Prediction
1. SignalP11 [12] for prediction of classically
secreted proteins (CSP).
2. SecretomeP12 [13] for prediction of nonclassical secreted proteins (NCSP).
3. TargetP13 [14] for prediction of mitochondrial
proteins.
4. TMHMM14 [15] for prediction of transmembrane proteins.
Therapeutic Target Prediction
1. SPAAN [16] to predict the probability of a
protein to act as adhesins. This tool is available under free academic license from the
developers of the tool.
Methods
Software
Protein Level Annotation
1. NCBI’s ORF finder4 or EMBOSS [5] getorf
tool (for local set up) for prediction of open
reading frames from nucleotide sequences.
2. Fungal genomes Blast5 to search for proteins
homologous to known fungal proteins.
3. Blast2go6 [6, 7] for gene ontology annotation
of proteins.
4. KAAS7 [8] for pathway mapping of proteins.
5. iPath28 [9] for graphical representation of pathway associations of proteins in the dataset.
http://www.ncbi.nlm.nih.gov.
http://www.ncbi.nlm.nih.gov/protein.
3
http://www.drugbank.ca/downloads.
4
http://www.ncbi.nlm.nih.gov/projects/gorf/.
5
http://www.ncbi.nlm.nih.gov/sutils/genom_table.cgi?
organism=fungi.
6
http://www.blast2go.org/.
7
www.genome.jp/kaas.
8
http://pathways.embl.de/.
1
The protocol described here is a general approach
to functionally annotate hypothetical proteins
based on similarity searches. However, many
parts of the workflow (shown in Fig. 55.1) are
independent of others, so some parts can be
deleted according to the requirements. We applied
the protocol to download (refer to the Sect. Data)
C. gattii and C. grubii proteins.
Translate Nucleotide Sequences into
Protein Sequences
1. Submit nucleotide sequences (if used) in
FASTA format at NCBI ORF finder web
server for conceptual translation into putative
2
http://www.ebi.ac.uk/Tools/pfa/iprscan/.
ftp://ftp.ncbi.nih.gov/blast/executables/blast+/LATEST/.
11
http://www.cbs.dtu.dk/services/SignalP/.
12
http://www.cbs.dtu.dk/services/SecretomeP/.
13
http://www.cbs.dtu.dk/services/TargetP/.
14
http://www.cbs.dtu.dk/services/TMHMM/.
9
10
55
High-Throughput Functional Annotation and Data Mining of Fungal Genomes…
561
Fig. 55.1 The workflow protocol described here is a general approach to functionally annotate hypothetical proteins based on similarity searches. Many parts of the
workflow are independent of others, so some parts can be
deleted according to the requirements
proteins from predicted open reading frames.
Use standard code (translation table 1) or the
alternative yeast nuclear code (translation
table 12), depending on the fungal organism.
Consider the minimum length of 100 bases for
nucleotide sequences translation to get reliable predictions in the following steps. For
local setup, install EMBOSS getorf package
on your local machine.
After all the nucleotide sequences get translated
into proteins, run them through each phase of the
work flow shown in Fig. 55.1.
Protein Level Annotation
For annotation of proteins, BlastP against known
fungal proteins, Blast2go, KAAS, and Interproscan
are used as shown in Phase 1 of Fig. 55.1.
1. Paste protein sequence on fungal genomes
blast page. Choose query and database as protein and blast program as BlastP. We found
1,278 (20.6 %) C. gattii proteins dissimilar to
Saccharomyces cerevisiae proteins. 508
(7.3 %) C. grubii proteins were found dissimilar to Aspergillus known proteins.
562
2. Load protein sequences into Blast2go software for gene ontology annotation.15 We were
able to annotate 135 (2.17 %) hypothetical
proteins.
3. Submit protein sequences to KAAS web
server. Uncheck the nucleotide box. Choose
the same organism for proteins or the closest
organism from organism list box using bidirectional best hit method. KAAS will map
our KEGG orthology (KO) terms along with
association of proteins with pathways and
BRITE objects. All the files can be downloaded on local machine. We were able to map
2,288 (36.8 %) proteins with KO terms for C.
gattii and 2,399 (34.4 %) for C. grubii.
4. Submit unique KO ids from KAAS mapping
to iPath for graphical representation of pathway associations in proteins sample on global
pathway maps. This tool provides extensive
map customization capabilities. After plotting, the map can be downloaded on local
machine.
5. Submit protein sequences in FASTA format to
Interproscan for protein domains mapping.
This program is computationally very expensive to run locally, especially for large datasets. We were able to annotate 6,001 (96.6 %)
proteins for C. gattii and 6,614 (94.9 %) proteins for C. grubii with protein domains.
Secretory Proteins Prediction
For prediction of ES proteins, a combination of
four tools, SecretomeP, SignalP, TargetP, and
TMHMM is used as shown in Phase 2 of Fig. 55.1.
All the methods described here have been implemented previously in fungal studies [17, 18].
1. Submit protein sequence in FASTA format at
SignalP 3.0 server. Use eukaryotes as an
organism group for fungal proteins. SignalP is
based on neural networks (NN) and Hidden
Markov models (HMM). We recommend the
use of both methods for reliable prediction
A detailed documentation of how to run Blast2go is
available at http://www.blast2go.org/start_blast2go.
15
G. Garg and S. Ranganathan
results with standard output. Truncation field
needs to be set according to the protein
sequence. In the output, choose sequence having D score in SignalP-NN result and signal
peptide probability in SignalP-HMM greater
than 0.5 as classically secreted. These thresholds have been very well tested in other studies [19]. We were able to find 419 (6.7 %)
proteins as CSP for C. gattii and 462 (6.6 %)
for C. grubii using SignalP.
2. Submit protein sequences found nonsecretory
in the previous step to SecretomeP 2.0 server.
This server predicts nonclassical secretory proteins based on large number of amino acid features along with results of other feature
prediction servers such as SignalP to obtain
information on various post-translational and
localization aspects of the protein. SecretomeP
run SignalP as well but SignalP is run separately
in the previous step because of the stringent cut
offs of D score and signal peptide probability
used in the protocol for reliable results. This is
not the case for SignalP running inside
SecretomeP. SecretomeP is based on neural
network and we recommend selecting protein
as nonclassical secreted where NN score is
greater than equal to 0.9. We were able to find
94 (1.5 %) proteins as NCSP for C. gattii and
106 (1.5 %) for C. grubii using SecretomeP.
3. Combine protein sequences predicted as classically and nonclassically secreted and submit
to TargetP 1.1 server. Select Non-plant as
organism group and specificity greater than
0.95 in cut offs section. Consider a protein
sequence as mitochondrial if Loc column is M
in the output. Delete these proteins from the
set of secretory proteins predicted in previous
steps. A total of 19 (0.3 %) for C. gattii and 20
(0.3 %) for C. grubii were predicted as mitochondrial proteins.
4. Submit final proteins dataset after the deletion
of mitochondrial proteins to TMHMM 2.0
server. Consider a protein sequence as nontransmembrane protein if number of predicted
TMHs (transmembrane helices) is 0 or 1 in the
final output. Such proteins are finally considered as secretory proteins. We consider proteins
having one transmembrane helix in our final set
55
High-Throughput Functional Annotation and Data Mining of Fungal Genomes…
of secretory proteins as these can be surface
proteins having therapeutic value as potential
vaccine candidates. A total of 384 (6.2 %) for
C. gattii and 440 (6.3 %) for C. grubii were
finally predicted as ES proteins.
In addition to the computational approach (shown
in Fig. 55.1) for the prediction of ES proteins,
sequence similarity search can be performed
against known fungal ES proteins for the prediction of ES proteins, using BlastP.
Therapeutic Target Prediction
Lot of fungal species are pathogenic to humans.
ES proteins of pathogens play a key role during
pathogenic infections [20]. ES proteins predicted
in Phase 2 can be checked computationally for
therapeutic value.
This phase of the protocol is a tricky one. All
the steps of this phase need to be performed on a
local machine and command line operation is
necessary. No specific web server like other parts
of protocol is available. Different components are
combined together for therapeutic targets prediction as shown in Phase 3 in Fig. 55.1.
1. To be a potential therapeutic target, a protein
should not be present in human. To find out
the secretory proteins similar to human proteins, search secretory proteins for sequence
similarity against human proteins using BlastP.
Sample command line for this operation is as
follows:
blastall -i query -d human.fa –m 8 –e 1e-08 -o
blast.out
Here query is the input file of protein
sequences in fasta format, human.fa is the
database file used for blast search, blast.out is
the blast output file. Use –m 8 option to provide blast result in tabular format, which is
easy to parse. E-value threshold is 1e-08. The
command line shown here can be altered
according to the datasets.16
Detailed description of all command line parameters of
blast is available at http://www.ncbi.nlm.nih.gov/books/
NBK1763/.
16
563
Proteins found dissimilar to human proteins
are searched for therapeutic value in terms of
drug targets and potential vaccine candidates.
We found 245 (3.9 %) secreted proteins for
C. grubii and 261 (3.8 %) for C. gattii, similar
to human proteins.
2. For drug target prediction, human dissimilar
ES proteins are searched for sequence similarity against known drug targets from DrugBank
using BlastP. Use the same command line
mentioned above for this operation by changing the input, database, and output files. We
found six potential drug targets for C. gattii
and four for C. grubii in respective secreted
proteins, mappable to known drug targets.
3. To test human dissimilar ES proteins for a
potential vaccine candidate, a protein should
not be present in mouse along with humans
because most of the vaccine candidates are
tested on mouse before they are tested on
humans, so proteins that are found dissimilar
to humans are tested for similarity against
mouse proteins. Use the same command line
mentioned above for this operation changing
the input, database, and output files. We found
further 86 (1.4 %) for C. gattii and 108 (1.5 %)
for C. grubii secreted proteins similar to
human dissimilar ES proteins.
Adhesins are cell surface proteins that are
present during host pathogen invasion. These
proteins play an important part in pathogenicity. Due to their important role in pathogenic
infection, these proteins are good vaccine
candidates.
4. To predict the probability of a protein to act as
adhesins run SPAAN program according to
program guidelines for proteins found dissimilar to human and mouse proteins. Consider
a protein to be predicted as adhesion if Padvalue in the SPAAN output is greater than 0.7.
This tool has been applied previously to fungal proteins for prediction of adhesins and
adhesin-like molecules [21]. We predict 33
(0.5 %) for C. gattii and 35 (0.5 %) for C. grubii as potential vaccine candidates.
Detailed result files from our fungal protein annotation are available from http://estexplorer.
biolinfo.org/fungal_annotation/
564
Notes
1. All the tools except ORF finder and iPath used
in the protocol are available free to install
locally under academic license. Although
some of the tools available as web servers, it is
recommended to install these tools locally on
a Linux machine by following tool installation
guidelines for big sequence datasets.
2. All the databases used for blast search locally
needs to converted to blastable format
before use by formatdb (provided in blast
executables).
3. All the therapeutic targets predicted using this
protocol are preliminary predictions which
need to be further validated by additional
computation analysis such as structural modeling and by experimental assays.
Acknowledgments We would like to thank Mr. Ben
Herbert, for introducing us to the pathogenic Cryptococcal
fungal genomes. GG acknowledges the award of
Australian Postgraduate Award scholarship from
Macquarie University.
References
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Application of Support Vector
Machines in Fungal Genome
and Proteome Annotation
56
Sonal Modak, Shimantika Sharma, Prashant
Prabhakar, Akshay Yadav, and V.K. Jayaraman
Abstract
Support Vector Machines (SVM) is a statistical machine learning algorithm
that has been used extensively in the past 3 years in computational biology. SVM has been widely used to detect, classify, and predict complex
biological patterns. SVMs have been widely applied to many areas of bioinformatics, including protein function prediction, functional site recognition, transcription initiation site prediction, and gene expression data
classification. This chapter gives a brief overview of SVM algorithm along
with its specific applications in fungal genome and proteome annotations.
Keywords
Support vector machines (SVM) algorithm • Kernel functions • Hyperplane
equation
Introduction
S. Modak
Bioinformatics Centre, University of Pune,
Ganeshkhind, Pune, Maharashtra 411007, India
S. Sharma • P. Prabhakar
Department of Biotechnology, Dr. D.Y. Patil University,
Mumbai- Bangalore Highway, Tathawade, Pune,
Maharashtra 411033, India
A. Yadav • V.K. Jayaraman (*)
Centre for Development of Advanced Computing
(C-DAC), Scientific and Engineering Computing
Group (SECG), University of Pune, Ganeshkhind, Pune,
Maharashtra 411007, India
e-mail: jayaramanv@cdac.in
Some fungi are pathogens and cause diseases in
plants and humans, whereas others are a rich
source of therapeutic metabolites and useful
enzymes, thus serving as basic models for molecular and cellular biology [1]. Genome sequencing
and functional genomics provide an insight into
the biological mechanisms in these fungi. Around
40 fungal genome sequences are now available
and a further 50 genome sequencing projects are
in progress. The advances in genomic technologies, such as microarrays and high-throughput
sequencing, have enabled detailed analysis of
fungal biochemistry. This analysis is automated
using various bioinformatics tools in order to
save time and effort.
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_56, © Springer Science+Business Media, LLC 2013
565
566
Rapid developments in fungal genomics and
proteomics have generated a large amount of biological data. Development and application of
machine learning tools can help to solve challenging problems and pave the way for discovery
of potent drugs. Analyzing fungal data sets
requires understanding of the data by inferring
structure or patterns from the data. Examples of
this type of analysis include fungal protein structure prediction, fungal gene classification, and so
forth.
Artificial Intelligence and machine learningbased methods can be employed for the purpose
of the aforementioned annotations. Different
classifiers based rigorously on principles of
learning theories and machine learning formalisms can be employed for recognizing patterns
in genomic and proteomic data. These classifiers
capture the patterns provided by the domain
information and group the data based on functionalities of the data. Classification is in general
a supervisory learning method. In this method
the algorithm needs domain features along with
their functionalities for training data. As an
example, we can illustrate the classification of
fungal adhesins as follows: We start with a set of
sequences which are experimentally annotated
already as adhesins and a set of non-adhesins.
Domain features can be extracted from these
sequences and the classifier builds a model to
group the data into adhesins and non-adhesins.
This model, known popularly as the supervisory
learning model, can readily be used for annotating new query sequences. Many different
machine learning methods have already been
applied for classification, like k-nearest neighbors [2], hierarchical clustering, self-organizing
maps, [3, 4], and Support Vector Machines
(SVM) or Bayesian networks. For solving
classification problems, machine learning techniques first obtain information from a set of
already labelled instances, called training data,
and then use this information to classify unknown
instances, called test data.
S. Modak et al.
Material
Support Vector Machine
SVM is a classifier that has been formulated from
statistical learning theory and structural risk minimization principle by Vapnik (1995) [5] and is
widely employed in different fields of science
and engineering. It is a parametric statistical linear classifier that performs a nonlinear mapping
of the input space to a new feature space to which
a linear machine can be applied. SVM constructs
a hyperplane separating the positive examples
from negative ones in the new space representation (Fig. 56.1). To avoid overfitting, SVM
chooses the Optimal Separating Hyperplane that
maximizes the margin in feature space [6]. The
margin is defined as the minimal distance between
the hyperplane and the training examples. The
selected data points that support the hyperplane
Fig. 56.1 SVM-based classification using hyperplane
56
Application of Support Vector Machines in Fungal Genome and Proteome Annotation
are called support vectors. A smaller number of
support vectors reflect a better generalization for
linearly separable problems. SVM employs a
maximum margin hyperplane for separating
examples belonging to two different classes. For
nonlinearly separable problems, the data are first
transformed into a higher dimensional feature
space and, subsequently, SVM employs a maximum margin linear hyperplane. Appropriate kernel functions are then employed for carrying out
all the simulations in the input space itself.
During the past 3 years the popularity of
SVMs as classification and prediction tools has
increased drastically [7]. The main reason for the
superior performance of SVMs is due to the fact
that they minimize the structural risk as opposed
to the empirical risk employed by some popular
classifiers. This feature of SVM produces a more
generalized model which performs well, both on
the training data and unseen data.
The tremendous success of SVMs in prediction and classifications has enabled them to be
used in fungal genome and proteome annotations.
Fungi are eukaryotic saprophytic organisms that
produce many commercially important products.
Additionally, many fungi are important clinically
because they cause many diseases in humans,
plants, and other animals. During the postgenomic era, the amount of data available on fungal genomes and proteomes has grown enormously.
Hence, there is an urgent need for development of
new, rapid, high-throughput in silico techniques
for fungal genome and proteome annotations, as
the experimental techniques, though highly reliable, are very expensive and time-consuming.
For the purpose of identification of gene and
protein functions employing SVM we need to
provide domain features. As an example, for
identification of defensin proteins the presence of
certain amino acids or dipeptides in high-propensity levels may be necessary. Hydrophobicity can
be an important domain feature required for prediction of membrane localized proteins. Similarly
structural features, like accessible surface area
and contact order, may correlate with ligand
binding proteins. From the sequences these features can be extracted and subsequently employed
by SVM for building appropriate models.
567
Some of the features that can be used for
genome annotation using SVM are listed below:
• Nucleotide compositions
• Dinucleotide frequencies
• Position-specific scoring matrices (PSSM)
profiles
• K-mer counts
• Motif conservation
• Promoter melting temperature
• DNA bending
• Transcript length
• Binding site degeneracy
• Binding site conservation
Some of the features that can be used for
proteome annotation using SVM are listed
below:
• Amino acid composition
• Frequencies of doublet, triplet, and multiplet
of amino acids
• Charge composition
• Hydrophobicity composition
• Multiplet composition
• Position-specific scoring matrix (PSSM)
• Number of protein–protein interactions
• Gapped amino acid composition
• Signal sequences
• Isoelectric point
• Mean hydropathy
• Codon usage bias
• Remote Homology using PSSM
• Secondary Structure information
• Structural features like surface accessibility,
coordinates of atoms, contact order
Methods
Feature Selection
In order to understand the molecular mechanisms
in fungi, it is essential to identify the relevant biological features/attributes in large and complex
datasets that regulate biological processes underlying the fungal data. Each feature pattern carries
information relevant to describe a specific function in fungal data. However, fungal datasets may
contain redundant features that do not contribute
at all to the classification task. To overcome this
568
S. Modak et al.
Fig. 56.2 Advantages of
feature selection
Fig. 56.3 Filter and
Wrapper: the two
approaches for feature
selection
problem, one way is to select only a small subset
of informative features from the fungal data
(Fig. 56.2). This technique helps in getting rid of
noisy features that do not correlate with the annotation problem at hand and thereby differentiates
classes well. If such noisy features are not eliminated from the dataset, the classification performance of the learning algorithm may decrease
dramatically, which will increase the computation time. Thus, there is a need to incorporate
techniques that search for the best set of features
with maximum classification performance. These
dimensionality reduction techniques are often
referred to as feature selection [8].
Feature selection algorithms mainly fall into
two categories: wrappers and filters (Fig. 56.3).
Wrappers make use of a learning algorithm to
estimate the quality of features. Methods like Ant
Colony Optimization and Genetic Algorithm in
combination with a classifier like SVM fall into
this category. On the other hand, filters evaluate
the quality of features considering the inherent
properties of the individual features without making use of a learning algorithm. Methods based
on statistical tests and mutual information fall
into this category. Since wrappers employ a learning algorithm for evaluating the quality of the
genes, they give more accurate results than
the filter methods. However, they need to train
the learning algorithm, which makes the wrappers
more time-consuming than filters. In addition,
wrappers need to be re-executed when switching
from one learning algorithm to another [9].
Most fungal classification algorithms involve
three main steps: feature extraction, feature selection, and classification (Fig. 56.4). In the feature
56
Application of Support Vector Machines in Fungal Genome and Proteome Annotation
569
Fig. 56.4 Application of
SVM in fungal genome
annotation
extraction step, a set of features is extracted that
are capable of describing a large fungal data
accurately. This step is followed by the feature
selection step, which is a dimensionality reduction step in which only the informative features
are selected from the previously extracted domain
features. Finally, the fungal data are divided into
different groups using a classifier such as SVM.
CASE STUDY: SVM-Based Prediction
Method for Fungal Adhesins
and Adhesin-Like Proteins
In this following case study, the authors have
tried to build a tool using SVM for prediction of
adhesin and adhesin-like proteins from fungal
domain (Fig. 56.5) [10]. Adhesins are cell surface proteins that confer on the microbes the ability to attach to cells, tissues, and/or abiotic
surfaces. Adhesins are the first line of pathogen’s
strategy of host cell invasion and therefore
determine its virulence. Mostly fungal adhesins
have a definite modular structure consisting of
an N-terminal carbohydrate or peptide-binding
domain, central Ser and Thr-rich glycosylated
domains and C-terminal region that mediates
covalent cross-linking to the wall through
modified glycosylphosphatidylinositol (GPI)
anchors. But all fungal adhesins cannot be
identified on this basis. Experimental
identification of adhesins is very expensive and
time-consuming. Two computational algorithms
for the prediction of adhesins are currently available: Software Program for Prediction of Adhesins
and Adhesin-like proteins using Neural Networks
(SPAAN) and Malarial Adhesins and Adhesinlike proteins Predictor (MAAP). Identification of
adhesion molecules would further our understanding of host-tissue adhesion in fungi, thereby
aiding the exploration of novel antifungal drug
targets and vaccine candidates.
570
S. Modak et al.
Fig. 56.5 SVM-based prediction system for Adhesins
Dataset Collection
For the compilation of the dataset, the authors
used different keywords like “adhesin,”
“flocculin,” and “agglutinin,” while limiting the
search to fungal domain to compile a raw pool of
fungal adhesin sequences from sequence
(Genbank and UniProt) databases. For a nonadhesin set they collected the proteins with
known intracellular locations, such as nucleus,
cytoplasm, mitochondria, endoplasmic reticulum, and so forth, and hypothetical proteins and
protein fragments were removed.
Redundancy Removal
The CD-HIT program was used to reduce the
redundancy to 50% in both positive and negative
datasets. Thus, well-curated 75 adhesins and 341
non-adhesins from fungi were collected.
Feature Extraction
The authors used different types of models using
different types of features. Hybrid models were
also built combining different types of features.
The lists of features used are as follows:
• Amino acid composition (AAC)
• Dipeptide composition (DPC)
• Charge composition (CC)
• Hydrophobicity composition (HC)
• Multiplet composition (MPC)
• Position-specific scoring matrix (PSSM)
profile
• Hybrid features combining AAC, CC, HC,
and MPC features (ACHM)
Results
Employing different types of features like AAC,
DPC, and so on as, inputs to the SVM, different
maximum margin hyperplane SVM models were
developed. For maximizing performance they tried
to use different kernels and tuned the kernel parameters for maximizing classification performance.
The SVM models were trained on all the aforementioned features and their accuracy was checked.
The best models were the hybrid model ACHM,
PSSM-a, and PSSM-b, which had accuracies of
86.05%, 86.29%, and 81.97%, respectively.
Selection of Informative Features
The most appropriate domain features need to be
input as features for SVM classification. Irrelevant
features may intrude with the classification process and reduce algorithm performance. There is
a need to incorporate techniques that search for
an “informative” set of features with “increased”
classification performance. These are referred to
as feature selection. We have employed a filterbased feature selection methodology to further
improve adhesin prediction performance.
56
Application of Support Vector Machines in Fungal Genome and Proteome Annotation
571
Table 56.1 Filter-based feature selection methodology to improve adhesin prediction performance
Selected Monopeptide
Selected Dipeptide
Selected Monopeptide and Dipeptide
Selected number
of features
15
192
207
The Waikato Environment for Knowledge
Analysis (WEKA) is a machine learning workbench that provides a general-purpose environment for automatic classification, regression,
clustering, and feature selection for common
data-mining problems in bioinformatics research.
It contains an extensive collection of machine
learning algorithms and data preprocessing methods complemented by graphical user interfaces
for data exploration and the experimental comparison of different machine learning techniques
on the same problem. WEKA can process data
given in the form of a single relational table. Its
main objectives are to (1) assist users in extracting useful information from data and (2) enable
them to easily identify a suitable algorithm from
which to generate an accurate predictive model.
In this work we have employed the information gain heuristics available in WEKA to rank
the features. The top-ranking features that provided the best accuracy were subsequently
employed in the final SVM model. It can be seen
from Table 56.1 that the performance of adhesin
prediction has improved by employing feature
selection.
Review of Some Recent Fungal
Bioinformatics Applications Using SVM
In this section, we outline a few important problems in bioinformatics where SVM and feature
selection have been applied with interesting
results on many real-world fungi case studies.
FaaPred: A SVM-based Prediction Method
for Fungal Adhesins and Adhesion-like
Proteins
This particular application has already been discussed previously. The server has been developed
for SVM-based prediction of adhesins [10].
Total number
of features
20
400
420
Cross-validation
accuracy
91%
88%
89%
Kernel
function
RBF
RBF
RBF
The prediction was based on two of the bestperforming models, as discussed above. The prediction web server is freely accessible at http://
bioinfo.icgeb.res.in/faap.
In silico Prediction of Yeast Deletion
Phenotypes
Gene function can be predicted by studying gene
duplications, gene deletions, and so forth.
A modified simulated annealing algorithm for
feature selection and weighting was developed to
evaluate such phenotypic effects of gene deletions on yeast [11]. The high weight selected features comprised phylogenetic conservation scores
for bacteria, fungi, Ascomycota, plants and mammals, degree of paralogy, and protein–protein
interactions (PPIs) count. SVM along with
weighted k-nearest neighbor were used for
classification. The validation was done by prediction of essential genes that cause morphological
changes in yeast.
Regulatory Analysis and Transcription
Factor Target Prediction in Yeast
A number of regulatory binding sites within fungal genomes have been identified using highthroughput technologies such as array-based
chromatin immunoprecipitation. However, in
lower eukaryotes these sites are not identified
completely. SVM was used to identify these binding sites, and the results were compared to those
obtained using PSSMs for a set of 104
Saccharomyces cerevisiae regulators [12].
Results indicate that when specificity and positive predictive values are the same, SVM-based
target classification is more sensitive (73% vs.
20%). SVM classifier was applied for each transcriptional regulator to all promoters in the yeast
genome to obtain new targets. SVM predicted
possible new roles for transcription factors like
Gnc4 and Rap1. For instance, Rap1 was predicted
572
to be involved in regulation of fermentative
growth. Promoter melting temperature curves
were also examined for the targets YJR060W,
and the targets showed unique physical properties
which distinguish them from other genes. Highquality predictions and biological functions can
be studied, employing the feature reduction, and
clustering strategies accuracy can be increased.
Predictions for 104 transcription factors are available, and remaining factors can also be built.
FGsub: Fusarium Graminearum Protein
Subcellular Localizations Predicted
from Primary Structures
Fusarium graminearum is a fungal pathogen that
causes several destructive crop diseases. The
F. graminearum proteins should be assigned to
different subcellular localizations in order to function properly. Therefore, these subcellular localizations can be used to gain insights into functions
and pathogenic mechanism studies of the respective fungus. A new prediction method, Fgsub, was
developed to predict the F. gramineaurum protein
subcellular localizations from the primary structures [13]. A fungi data set with subcellular localization information is collected from UniProtKB
database and used as training set. The subcellular
locations were classified into ten groups. SVM
was used for training of data and for prediction of
the F. graminearum protein subcellular localizations. Ten-fold cross-validation accuracy showed
efficient prediction results. It was also found that
F. graminearum proteins bear significant sequence
similarity to those present in the training set. For
efficient annotations, BLAST was used, which
increases the prediction coverage. Subcellular
localizations of 12786 F. graminearum proteins
were predicted, thus providing insights into protein functions and pathogenic mechanisms of this
destructive pathogen fungus.
GPI-Anchored-Protein Identification
System to Mine the Protein Databases
of Aspergillus fumigatus, Aspergillus
nidulans, and Aspergillus oryzae
Various computational approaches have been
developed in order to identify GPI-anchored proteins in protein sequence databases. A new
S. Modak et al.
sequence-based approach for identification using
SVM algorithm was developed [14]. The algorithm recognizes COOH-terminal sequences and
uses a classifier based on voting strategy to recognize appropriate NH2-terminal sequences. The
classifier achieved a high accuracy of 96% in
five-fold cross-validation testing. The votingbased classifier gave a higher accuracy of 98.88%
when used on a test dataset of eukaryotic proteins. S. cervisiae protein sequences were used,
which showed the classifier’s ability to classify
new unseen data. On using the predictor on three
aspergilla species, 115 GPI-anchored proteins in
Aspergillu fumigatus, 129 in Aspergillus nidulans, and 136 in Aspergillus oryzae were
identified. Half of these proteins had conserved
domains when the sequence-based conserved
domain search was applied.
Screening Noncoding RNAs in
Transcriptomes from Neglected Species
Using PORTRAIT: Case Study of the
Pathogenic Fungus Paracoccidioides
brasiliensis
Structural genomic information can be obtained
from transcriptome sequences. Such sequences
can also be useful in characterizing neglected
species that do not have the chance of undergoing
whole-genome sequencing. But sequencing of
these organisms is difficult owing to low quality
of reads and incomplete coverage of transcripts.
Another factor is lack of known protein homologs;
also, noncoding RNAs may be caught during
sequencing. These RNAs do not code for protein
and instead perform the unique function of folding
into other structural conformations. The analysis
of such transcriptome sequences is limited.
PORTRAIT is an algorithm that has been developed for analysis of such ncRNAs from poorly
characterized species [15]. These sequences are
translated by software that identifies sequencing
errors and predicts putative proteins along with
their transcripts. These are evaluated for coding
potential by SVM. Two models have been developed. First, if a putative protein is found, a protein-dependent SVM model is used. If one is not
found, a protein-independent SVM model is used.
No homology information is used, as only
56
Application of Support Vector Machines in Fungal Genome and Proteome Annotation
ab-initio features are extracted. PORTRAIT was
used for predicting ncRNAs from the transcriptome of pathogenic fungus Paracoccidioides
brasiliensis and five related other fungi.
Determination of Protein Content
of Auricularia auricula Using
Near-Infrared Spectroscopy
Combined with Linear
and Nonlinear Calibrations
Near-infrared (NIR) spectroscopy has been used
to determine the protein content of Auricularia
auricula, also known as woody ear or tree ear,
using partial least-squares (PLS), multiple regressions (MLR), and least-squares-support vector
machine (LS-SVM) [16]. These were tested and
compared against other different methods such
as Savitzky-Golay (SG) smoothing, standard
normal variate, multiplicative scatter correction
(MSC), first derivative, second derivative, and
direct orthogonal signal correction. A successive
projections algorithm (SPA) was also used to
find effective wavelength selection. Different
combinations of pretreatment and calibration
methods were compared based on the predictions. The optimal full-spectrum PLS model was
obtained by raw spectra, whereas the optimal
SPA-MLR, SPA-PLS, and SPA-LS-SVM model
obtained MSC spectra. The best performance
was achieved by SPA-LS-SVM model. NIR
spectroscopy combined with SPA-LS-SVM is
useful in determining the protein content of
A. auricula.
ESLpred2: Improved Method
for Predicting Subcellular Localization
of Eukaryotic Proteins
ESLpred method uses new features as an input to
SVM [17]. Three kingdom-specific protein
sequence sets, 1198 fungi sequences, were
included in the study for predictions. Evolutionary
information in the form of profile composition
along with whole and N-terminal sequence composition as an input feature vector of 440 dimensions was given. Overall accuracies of greater
than 72% were achieved for fivefold cross-validation. Similarity search-based results, when
used with whole and N-terminal sequence and
573
profile composition information, gave accuracies
of 75.9%, 80.8%, and 76.6%, respectively. 1
Vector-G: Multimodular SVM-Based
Heterotrimeric G Protein Prediction
Heterotrimeric G proteins interact with GPCRs in
response to any stimulus generated by hormones,
neurotransmitters, chemokines, and sensory signals to intracellular signaling cascades. G protein
subunits have been found to play an important
role in different eukaryotic diseases including
inflammation, cardiovascular diseases, and neurological disorders as well as in plant pathogens
response, and differentiation and virulence of
pathogenic fungi. All the methods available for
finding new G proteins are based on homology
search analyses, which are not robust. Vector-G is
an SVM-based pattern recognition algorithm for
finding new G proteins and their homologs [18].
Properties such as physicochemical and dipeptide, tripeptide, and hydrophobicity composition
are used for generating SVM classifiers. This
method gave 96.17%, 95.38%, and 97.6% sensitivity and 99.45%, 100%, and 100% specificity
on tests sets for G protein alpha, beta, and gamma
subunits, respectively. The algorithm correctly
predicts known alpha, beta, and gamma subunits.
New G protein subunits are predicted in 31
genome covering plant, fungi, and animal kingdom. The software is available at the website
http://biomine.cs.uah.edu/bioinformatics/svm_
prog/scripts/GProteins/vectorg.html. The supplementary files are available on http://www.bioinfo.
de/isb/2008/08/0013/supplementary_material/
Physical PPIs Predicted from Microarrays
Microarray data reveal functionally associated
proteins. Predicting physical interactions directly
through microarrays are both important and challenging as most proteins that are associated are
not actually in direct physical contact. Thus, an
SVM-based method was developed to predict the
pairs that are likely to interact [19]. This method
was applied to predict interactions in yeast
The ESLpred2 server is also developed and available
with many models at http://www.imtech.res.in/raghava/
eslpred2/.
1
574
(S. cerevisiae). Literature search revealed several
new predictions that could be experimentally
validated. This new method holds a promise to
improve the annotation of interactions as one
component of multisource integrated systems.
Kernel-Based Machine Learning
Algorithm for the Efficient Prediction
of Type III Polyketide Synthase Family
of Proteins
Type III polyketide synthase is a protein family
that has a significant role in biosynthesis of various polyketides in plants, fungi, and bacteria.
These proteins have positive effects on human
health; thus, developing a tool to identify the
probability of sequence being a type III polyketide
synthase is helpful. PKSIIIpred is a prediction
server for type III PKS where SVM is used for
classification [20]. The tool predicts efficiently
the type III PKS superfamily of proteins with
high sensitivity and specificity.2
Predicting PPIs from Protein Sequences
using Meta Predictor
PPIs prediction based on meta approach has been
developed, which predicts PPIs by using SVM
that combines results by six independent predictors [21]. The method was used of S. cerevisiae
and Helicobacter pylori datasets .The final predicted model trained on the PPIs dataset of S. cerevisiae was used to predict interactions in other
species. The results obtained showed that this
model is also capable in performing cross-species
predictions.3
Conserved Codon Composition
of Ribosomal Protein Coding Genes
in Escherichia coli, Mycobacterium
tuberculosis, and S. cerevisiae: Lessons
from Supervised Machine Learning
in Functional Genomics
Advances in genomics have resulted in creation
of large sequence data. Annotation is currently
relied on sequence comparison and homologs.
The server is available at http://type3pks.in/prediction.
The source code and the datasets are available at http://
home.ustc.edu.cn/~jfxia/Meta_PPI.html.
2
3
S. Modak et al.
Codon composition, a fusion of codon usage and
amino acid composition signals was used to accurately discriminate, in the absence of homology
information, cytoplasmic ribosomal protein genes
from other genes of known function in S. cervisiae, E. coli, and M. tuberculosis using SVM
(light) [22]. Such analysis is helpful in determining features that provide individuality to ribosomal protein genes. Each of the sets of positively
charged, negatively charged, hydrophobic residues, codon bias, contribute to their individual
composition profile. SVM is used to detect, combine, and augment the representation of these signals in order to perform efficient classification.
This method can have several alternatives by
combining codon composition with other gene/
protein classification attributes.
Softwares for SVM
A number of SVM softwares have been developed recently. Table 56.2 presents a list of some
of these softwares.
Summary
Several fungi, form a rich source of therapeutic
metabolites and the others act as pathogens causing diseases. It is thus essential to understand the
biological mechanisms within fungi in order to
gain insights into various biological problems.
The amount of fungal genome data is thus increasing exponentially and in order to handle such a
vast amount of data, machine learning techniques
are applied in the field of fungal bioinformatics.
SVM is a statistical learning tool that employs
simple but very powerful machine learning algorithm for classification and regression. Feature
selection is a method that helps in reducing the
dimensionality of large fungal datasets by selecting only the relevant features. In this chapter, we
have highlighted some of the recent fungal
genome-proteome annotation problems in bioinformatics where SVM and feature selection have
been applied successfully. A number of biological
problems, such as identification of adhesion and
56
Application of Support Vector Machines in Fungal Genome and Proteome Annotation
575
Table 56.2 Software for support vector machines
Software
LIBSVM
SVMlight
SVMstruct
Weka
BioWeka
mySVM
mySVM/db
Gist
SmartLab
MATLAB SVM
Toolbox
Salient features
• Integrated software for support vector
classification, regression, and distribution
estimation.
• Developed at National Taiwan University by
Chang and Lin
• Developed in C++ and Java.
• Supports weighted SVMs for unbalanced data,
multiclass classification, cross-validation, and
automatic model selection.
• Linear, polynomial, radial basis function, and
neural (tanh) kernels available.
• Has a fast optimization algorithm.
• Efficient implementation of the leave–one–out
cross-validation and can be applied to very
large datasets.
• Developed in C++
• Polynomial, radial basis function, and neural
(tanh) kernels available
• Can model complex (multivariate) output data,
such as trees, sequences, or sets.
• Several implementations like SVMmulticlass,
SVMcfg, SVMalign, SVMhmm,
• Popular collection of machine learning
algorithms developed in Java.
• Supports a number of data-mining tasks such
as data preprocessing, clustering, classification,
regression, visualization, and feature selection.
• Also contains an SVM implementation.
• Weka for data analysis tasks in biology,
biochemistry and bioinformatics, and
knowledge discovery.
• Includes integration of the Weka LibSVM
project.
• It is available as C++ source code and
Windows binaries.
• Kernels available include linear, polynomial,
radial basis function, neural (tanh), and anova.
• Efficient extension to mySVM.
• Designed to run directly inside a relational
database using an internal JAVA engine
• C implementation of SVM classification
and kernel Principal Components Analysis.
• Available as an interactive Web server
at http://svm.sdsc.edu.
• Kernels available include linear, polynomial,
and radial.
• Provides several SVM implementations,
including cSVM, mcSVM, rSVM, and
javaSVM1, and javaSVM2.
• Developed by Gunn, implements SVM
classification and regression.
• Various kernels available include linear,
polynomial, Gaussian radial basis function,
exponential radial basis function, neural (tanh),
Fourier series, spline, and B spline.
Availability
http://www.csie.ntu.edu.tw/~cjlin/libsvm/
http://svmlight.joachims.org/
http://svmlight.joachims.org/svm_struct.html
http://www.cs.waikato.ac.nz/ml/weka/
http://sourceforge.net/projects/bioweka/
http://www-ai.cs.uni-dortmund.de/
SOFTWARE/MYSVM/index.html
http://www-ai.cs.uni-dortmund.de/
SOFTWARE/MYSVMDB/index.html
http://www.chibi.ubc.ca/gist/
http://www.smartlab.dibe.unige.it/
http://www.isis.ecs.soton.ac.uk/resources/
svminfo/
(continued)
576
S. Modak et al.
Table 56.2 (continued)
Software
TinySVM
LS-SVMlab
LSVM
ASVM
PSVM
GPDT
Spider
LEARNSC
SVM Toolbox
SVMTorch
Salient features
• C++ implementation of C-classification
and C-regression that uses sparse vector
representation.
• Can handle thousands of training instances
and feature dimensions.
• Developed by Suykens and is a MATLAB
implementation of Least-Squares Support
Vector Machines (LS–SVMs)
• LS–SVMprimal–dual formulations, formulated
for kernel PCA, kernel CCA, and kernel PLS
• Lagrangian Support Vector Machine is
a very fast SVM implementation developed
in MATLAB
• Active Support Vector Machine is a very
fast linear SVM script for MATLAB, by
Mangasarian and Musicant, developed
for large datasets.
• Proximal Support Vector Machine is a
MATLAB script developed by Fung and
Mangasarian.
• Developed by Serafini, et al., is a C++
implementation for large-scale SVM
classification in both scalar and distributed
memory parallel environments
• Object-orientated environment for machine
learning written in MATLAB
• Implements SVM multiclass classification and
regression
• Learning and Soft Computing by Kecman is
available as MATLAB script.
• MATLAB toolbox, contains many classification
algorithms like linear and quadratic
penalization, multiclass classification,
e-regression, n-regression, wavelet kernel,
and SVM feature selection.
• Developed by Collobert and Bengio is a part
of the Torch machine learning library
• Distributed as C++ source code or binaries
for Linux and Solaris
adhesion-like proteins, prediction of transcription
factor target in yeast, prediction of subcellular
localizations from primary structures, G-protein
prediction, prediction of type III polyketide synthase family of proteins, PPIs predictions, and so
on, have been solved using these machine learning approaches. These methods can be further
enhanced by making use of high-performing
computational systems, such as parallel computers, in order to solve complex fungal bioinformatics problems at a much faster rate.
Availability
http://chasen.org/~taku/software/TinySVM/
http://www.esat.kuleuven.ac.be/sista/
lssvmlab/
http://www.cs.wisc.edu/dmi/lsvm/
http://www.cs.wisc.edu/dmi/asvm/
http://www.cs.wisc.edu/dmi/svm/psvm/
http://dm.unife.it/gpdt/
http://www.kyb.tuebingen.mpg.de/bs/
people/spider/
http://www.support-vector.ws/html/
downloads.html
http://asi.insa-rouen.fr/%7Earakotom/
toolbox/index.html
http://bengio.abracadoudou.com/
SVMTorch.html
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Ramana J, Gupta D (2010) FaaPred: a SVM-based
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Holloway DT, Kon M, Delisi C (2007) Machine learning for regulatory analysis and transcription factor
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Sun C, Zhao XM, Tang W, Chen L (2010) FGsub:
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Garg A, Raghava GP (2008) ESLpred2: improved
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Bioinformatics Tools
for the Multilocus Phylogenetic
Analysis of Fungi
57
Devarajan Thangadurai and Jeyabalan Sangeetha
Abstract
Mycologists are generally identifying fungal communities by microscopic
and macroscopic assessment. This conventional approach has several limitations due to the growth and environmental factors. Hence, molecular
techniques and bioinformatics tools are essential in the field identification
and characterization of fungi. Multilocus sequences are widely used in
most of the bioinformatics tools and they can be used to recognize species
boundaries. Nucleic acid and protein sequences-based analysis in fungal
studies are revolutionizing the view on mycology. Numerous bioinformatics tools are available online to guide molecular biologists and biotechnologists. This chapter provides a guide to utilizing the available
bioinformatics tools on the World Wide Web for sequence alignment, editing, and multilocus phylogenetic analysis.
Keywords
Bioinformatics • Tools • Softwares • Databases • Multilocus phylogenetic
analysis • Fungi
Introduction
D. Thangadurai (*)
Department of Botany, Karnataka University, Dharwad,
Karnataka 580003, India
e-mail: drdthangadurai@gmail.com
J. Sangeetha
Department of Zoology, Karnataka University,
580003, Dharwad, Karnataka, India
e-mail: drsangeethajayabalan@gmail.com
Conventional biochemical methods and phenotypic tests for fungal species differentiation are
tedious and time-consuming and may require
specialized tests. Recent developments in molecular biology and bioinformatics allow the consideration of other methods that are more universal
and less time consuming [1]. Currently, scientific
research requires parallel strategy to simultaneously gather, examine, integrate, and store the
large volumes of data. In this scenario, researchers cannot attain their decisive goal without good
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0_57, © Springer Science+Business Media, LLC 2013
579
580
D. Thangadurai and J. Sangeetha
data handling and analyzing skills. Bioinformatics
is a new field of science that examines complex
biological data on the basis of statistics and computer science [2, 3]. Computer programs in biology that help to reveal the principle mechanisms
in biological problems related to the structure
and function of macromolecules, biochemical
pathways, disease processes, genetic analysis and
evolution are discussed [4].
Studies of fungal diversity in natural environment have been detonated in the last 20 years, in
large part due to the advent of the molecular techniques [5]. Regrettably, many of the molecular
methods developed for prokaryotes are not appropriate for fungi. This is due to the fact that the ribosomal small subunit, which is the important key of
prokaryotic diversity analysis, is insufficiently
variable to precisely distinguish fungal taxa [6, 7].
A major challenge in mycology is to make sense of
the enormous quantities of sequence data and
structural data that are generated by genome
sequencing projects, proteomics, and other largescale molecular biology efforts [4]. Bioinformatics
tools and techniques are playing an important
role in the study of fungal systems, especially the
function and evolution of fungal genes and
genomes. These methods are necessary to achieve
a complete, quantitative, and reliable understanding of functions of fungi in molecular level.
The systematic comparison of genomic
sequences and protein sequences from different
fungal species represents a central focus of contemporary fungal genome analysis. Multilevel
sequencing
provides
accurate
species
identifications as predicting evolutionary relationships among species. For the identification of
most species a database of 26S 5’-end base
sequences is adequate. Additional sequences are
needed to predict more secluded relationships.
Sequence alignment and editing can be made
rapidly on computational methods [8, 9]. Various
tools like CLUSTAL W/X, T-Coffee, DbClustal,
Kalign, and MAFFT are available for multiple
sequence alignments (MSAs) (Table 57.1).
BLAST, FASTA, and FRACTURA are available
for analyzing and interpreting algorithm, modeling and computer graphics of the databases on
genomics and proteomics [10].
Table 57.1 List of bioinformatics tools, software, and programs used in multilocus phylogenetic analysis in fungi
Name
Description
A-Bruijn Alignment Multiple alignment of sequences
with repeated and shuffled elements
Advanced
Multiple sequence alignment program
PipMaker
ALIGN Query
Multiple sequence alignment program
COBALT
AMAP
Bali-Phy
BLAST
BOXSHADE
CINEMA
CLUSTAL W
CodonCode aligner
Multiple protein sequence alignment
using conserved domain and local
sequence similarity information
Multiple alignment tool for peptide
sequences
Bayesian alignment and phylogeny
estimation
Compare gene or protein sequences
for similarity between the sequences
Pretty-printing of multiple sequence
alignments
Color interactive editor for multiple
alignments
Multiple sequence alignment program
for DNA and protein
Program for sequence assembly,
editing and mutation detection
Web link
http://nbcr.sdsc.edu/euler/aba_v1.0_dl/
References
[11, 12]
http://pipmaker.bx.psu.edu/pipmaker/
[13]
http://xylian.igh.cnrs.fr/bin/
align-guess.cgi
http://www.ncbi.nlm.nih.gov/
tools/cobalt/
[14]
http://packages.debian.org/sid/
amap-align
http://www.biomath.ucla.edu/
msuchard/bali-phy/
ftp://ftp.ncbi.nlm.nih.gov/blast
[15]
[14]
[14]
[16]
http://www.ch.embnet.org/
software/BOX_doc.html
http://www.bioinf.manchester.ac.uk/
dbbrowser/CINEMA2.1/
http://clustal.org/clustal2
[14]
[17]
http://www.codoncode.com/aligner/
[21]
[18–20]
(continued)
57
Bioinformatics Tools for the Multilocus Phylogenetic Analysis of Fungi
581
Table 57.1 (continued)
Name
ComAlign
Consensus
CVTree
DIALIGN-TX
DSC
eProbalign
ESPript 2.2
FFAS03
FOLDALIGN
GeneOrder 3.0
GOAnno
JDotter
LALIGN
LocARNA
Description
Combining many multiple alignments
in one improved alignment
Calculates the consensus for the
CLUSTAL or MSF multiple alignment
Infer phylogenetic relationships
between microbial organisms by
comparing their proteomes using a
composition vector approach
Multiple sequence alignment program
Divide-and-Conquer Multiple
Sequence Alignment
Generation and manipulation of
multiple sequence alignments using
partition function posterior probabilities
Multiple sequence alignments
in PostScript
Profile-profile alignment
and fold recognition algorithm
Semi-automated multiple global
alignment and structure prediction
Ideal for comparing multiple
GenBank genomes
Multiple alignments
of complete sequences
Multiple alignments
of complete sequences
Finds multiple matching
subsegments in multiple sequences
Multiple alignment of RNA molecules
MOTIF
Parsimony and simulated annealing
in the search for phylogenetic trees
Offers a range of multiple alignment
methods
Multilocus analysis of nucleotide
variation
Recognition of common patterns
and properties in multiple aligned
protein sequences
Motif-based multiple sequence
analysis
Multilocus coalescent simulations
for evolution
Sequence motif search
MULALBLA
Multiple alignment with Blast
Multilocus 1.3b
Facilitate analysis of multilocus
population genetic data for genotypic
diversity indices, linkage disequilibrium
indices and population differentiation
Multiple sequence alignment
LVB
MAFFT
MANVa 0.982b
MASIA
Meme
mlcoalsim 1.42
Multiple Align
Show
Web link
http://www.daimi.au.dk/~ocaprani/
ComAlign/ComAlign.html
http://coot.embl.de/Alignment/
consensus.html
http://cvtree.cbi.pku.edu.cn
References
[14]
[22]
[23]
http://dialign-tx.gobics.de/
http://bibiserv.techfak.
uni-bielefeld.de/dca/
http://probalign.njit.edu/
probalign/login
[14]
[24–28]
http://espript.ibcp.fr/ESPript/ESPript/
[30]
http://ffas.ljcrf.edu/ffas-cgi/cgi/
document.pl?ses=#over
http://foldalign.ku.dk/
[31, 32]
http://binf.gmu.edu:8080/
GeneOrder3.0/
http://bips.u-strasbg.fr/GOAnno/
GOAnnoHelp.html
http://athena.bioc.uvic.ca/pbr/jdotter/
[35, 36]
http://www.ch.embnet.org/
software/LALIGN_form.html
http://rna.informatik.uni-freiburg.
de:8080/LocARNA.jsp
http://biology.st-andrews.ac.uk/
cegg/lvb.aspx
http://mafft.cbrc.jp/alignment/software/
[29]
[33, 34]
[37]
[38]
[39]
[22]
[40]
[41, 42]
http://www.ub.edu/softevol/manva/
[22]
http://born.utmb.edu/masia/
[43]
http://www.sdsc.edu/~tbailey/
MEME-protocol-draft2/protocols.html
http://www.ub.edu/softevol/mlcoalsim/
[22]
http://www.genome.jp/tools/motif/
MOTIF3.html
http://www-archbac.u-psud.fr/
MULALBLA/mulalbla.html
http://www.agapow.net/software/
multilocus/1.3b/view
[22]
http://www.bioinformatics.org/
sms/multi_align.html
[44]
[22]
[22]
[45]
(continued)
582
D. Thangadurai and J. Sangeetha
Table 57.1 (continued)
Name
Mumsa
MUSCLE
MuSiC
NAST
Opal
ParAlign
Pebble 1.0
Prof
SCANMOT
T-Coffee
VerAlign
WebVar
YASS
Description
Multiple sequence alignment
Progressive alignment of nucleic acids
Multiple sequence alignment with
constraints
Multiple sequence alignment server
for comparative analysis of 16S
rRNA genes
Multiple sequence alignment
Rapid and sensitive similarity search
tool
Tool for analysis and simulation of
maximum likelihood and least-squares
methods
Multiple alignment and structure
prediction
Searching similarity using simultaneous
scan of multiple sequence motifs
Align DNA or RNA or protein
sequences
Multiple sequence alignment
comparison
Rapid estimation of relative site
variability from multiple sequence
alignments
Sequence similarity search tool
In general, bioinformatics tools are used to
determine gene functional annotation, gene family evolution, and genome organization. These
bioinformatics tools are developed with the help
of internet tools and World Wide Web. Many universal programs and software are available, and
they are providing with either free access or with
charges to biotechnology databases [10]. Several
programs, databases, and softwares can be used
to perform MSA, multiple sequence editing, and
phylogenetic analysis of individual DNA or protein sequences.
MSA method is a comparative method at the
molecular level and it is a vital component of
most of the bioinformatics techniques. The effectiveness of sequence analysis is based on the
addition of more data to yield stronger analyses
and also, if provided sequences which have
<40 % residue identity the analyses will becomes
unreliable [56]. Many methods are available for
Web link
http://msa.sbc.su.se/cgi-bin/msa.cgi
http://www.drive5.com/muscle/docs.htm
http://genome.life.nctu.edu.tw/MUSIC
References
[46]
[47]
[48]
http://greengenes.lbl.gov/NAST
[49]
http://opal.cs.arizona.edu/
http://www.paralign.org/
[22]
[50–52]
http://www.cebl.auckland.ac.nz/
software2.php
[22]
http://www.aber.ac.uk/~phiwww/prof/
[22]
http://caps.ncbs.res.in/scanmot/
scanmot.html
http://www.tcoffee.org/
[22]
http://www.ibivu.cs.vu.nl/programs/
veralignwww/
http://www.pesolelab.it/Tools/
WebVar.html
[22]
http://bioinfo.lifl.fr/yass/
[55]
[53]
[54]
MSA of three or more biological sequences such
as protein or nucleic acid of similar length. From
the output of MSA, homology can be inferred
and phylogenetic analysis can be carried out to
study the evolutionary relationship between the
sequences. Computational methods and algorithms are used to analyze the sequence alignments. Progressive alignment methods are more
efficient to execute for many sequences and are
commonly available on World Wide Web services [20, 53, 57, 58].
Materials
1.
2.
3.
4.
Computers with a lot of memory.
Macintosh, Windows, and Unix platforms.
Hi-speed internet connectivity.
Bioinformatics tools, softwares, and databases
(freely available or on purchase).
57
Bioinformatics Tools for the Multilocus Phylogenetic Analysis of Fungi
Methods
Understanding the basic principles and methodologies used in bioinformatics is more important
for mycologists to utilize available tools and to
interpret results for the multilocus phylogenetic
analysis. This chapter mainly focuses on several
bioinformatics tools that are commonly used in
the multilocus phylogenetic analysis of fungi
with step-by-step guidance.
CLUSTAL W
Clustal programs text menu are provided with all
of the options to do MSA and to create simple
phylogenetic tree. With these simple menus, this
program is highly convenient for the users and
run on all computers. The output of MSA can be
used to take printout or to manipulate and it can
incorporate secondary structure information into
the process [20, 57, 59, 60]. CLUSTAL X is a
new graphical windows interface for the
CLUSTAL W. CLUSTAL X includes new features like the ability to cut-and-paste sequences,
selection of a subset of the sequences to be
realigned, selection of a subrange of the alignment can be realigned and insert back to the original alignment and coloring option allows to
highlight required features or exceptional residues in the alignment. CLUSTAL W is the most
accurate and faster package [56].
CLUSTAL W can easily be installed by copying executable file to the system directory. Several
parameters (named*.par) and an online help text
file (clustalx.hlp for MS Windows, otherwise
clustalx_help) are also required. These files
should be copied to the directories specified by
the PATH environment variable or user’s current
directory [18, 19].
1. Download and install the CLUSTAL W
program.1
1
Available from http://www.clustal.org/clustal2.
583
2. Open the CLUSTAL W program; in UNIX
type “>CLUSTAL W” at the prompt within
the appropriate directory.
3. In CLUSTAL W menu select option 1.
4. Enter the file name of the sequences; the
screen will be back to the main menu.
5. Align the sequences by selecting option 2
and then choosing option 1 inside the multiple alignment menu.
6. Go to main menu of CLUSTAL W and
choose option 4 for Phylogenetic trees.
7. Choose option 4 inside the phylogenetic tree
menu and choose output file name.
8. Execute the draw tree command.
9. Go to main menu, select option X, and exit
CLUSTAL W.
10. Choose the retree program from Phylip package; in UNIX type “>retree” prompt within
the directory.
11. Type “Y” and as input give the tree file from
CLUSTAL W; use “?” to find all the option.
12. To find the node number of the sequence use
page up and down and choose a node as the
out group.
13. Exit retree after writing the tree with the new
root.
14. To draw the tree using diagram command in
the UNIX type “drawgram” prompt.
15. The program will ask for the input file name;
give the file name “outtree.”
16. Enter the file name of the tree from
CLUSTAL W.
17. Then the program will ask for font file name;
the user should have the font file font 1
among the files.
18. Phylip will take to a series of dragram menus
after giving the path and file name of font 1.
19. To see the postscript of the phylogenetic tree
choose L; choose N to preview the tree.
20. Choose 1 from the main drawgram menu and
choose phylogram.
21. Give option P, select 4, and provide an angle
of 90 ° to get the standard format of phylogenetic trees.
22. The output of phylogenetic tree will be given
in plot file in ps format.
23. View the phylogenetic tree using the command “ghostview.”
584
D. Thangadurai and J. Sangeetha
T-Coffee
Bayesian Estimation of Species Trees
T-Coffee is a MSA program that provides improvement in accuracy and speed as compared to other
alternatives [61]. T-Coffee is used to align nucleic
acid and protein sequences, to compare alignments, reformat, and also allows to combine
results obtained from several alignment methods
and produces new MSAs. To install T-Coffee user
need to have GCC, G77, CPAN, internet connection, and root password to install SOAP [62].
Gathering of the pairwise alignment is the main
step for T-Coffee method. This collection is called
as library. After computation of library, they can
be pooled and used to compute MSA. Mocca is a
special mode of T-Coffee that extracts a series of
repetition from a sequence [61].
1. Download the T-Coffee program.2
2. Type “uncompress distribution.tar.Z” prompt
to install the program.
3. Install CLUSTAL W 3 if not available.
4. Indicate the address and name of the
CLUSTAL W on system.
5. Set the global variable CLUSTAL W_4_
TCOFFEE to “path/name_of_CLUSTAL W”:
Setenv CLUSTAL W_4_TCOFFEE “path/
name_of_CLUSTAL W.”
6. Go to the main directory and type “./install.”
7. Appearance of “Installation of t_coffee
Successful” on the screen indicates successful completion of installation.
8. Add the bin folder to path “set
path = ($path.<address of the t_coffee bin
folders>).”
9. Type the sequence (Swiss-prot, Fasta, or Pir)
prompt “t_coffee sample_seq1.fasta” (see
Note 1).
10. Type the sequence in same file
“mocca<sequence>.”
11. The output of this file will be “<sequence>.
mocca_lib.”
Bayesian Estimation of Species Trees (BEST)
(version 2.3) is a phylogenetic package of programs written by Liang Liu to estimate the posterior distribution of species trees using multilocus
molecular data that accounts for deep coalescence
of alleles [63, 64]. All the BEST parameters are
usually defined in the popular Bayesian phylogenetic package, MrBayes using the preset command
[65, 66]. The program estimates both the posterior
joint distribution of gene trees and the posterior
distribution of the species trees jointly in one
Markov Chain Monte Carlo (MCMC) algorithm.
BEST can also estimate gene trees and the species
trees with divergence times and population sizes.
The posterior distribution can be summarized in a
program like MrBayes or PAUP [67–70].
1. Compile the program by simply typing
“make” in the folder where the source code
is available (see Note 2).
2. Run BEST 2.3 by simply typing “./best”
which will prompt a working environment
where one can type commands to manipulate
and analyze aligned sequences.
3. Alternatively, use the command line “./best
-idata.nex” in which commands in the
MrBayes block will be executed by the
program to analyze pre-prepared input file
“data.nex.”
4. Create the input file for BEST 2.3 in NEXUS
format consisting data block and MrBayes
block.
5. Concatenate the multilocus sequences across
loci in the data block as in the regular
MrBayes.
6. Replace the missing nucleotides/sequences
with question marks.
7. Duplicate the haploid genes if any compatible to the diploid genes (see Note 3).
8. Otherwise, randomly choose one of the two
sequences for the diploid genes compatible
to the haploid genes.
9. Command substitution models and prior distributions at gene tree level.
10. Set prior distribution for the species tree,
population sizes, and variable mutation rates
across genes.
Available at http://www.tcoffee.org/Projects_home_page/
t_coffee_home_page.html.
3
from http://www.ebi.ac.uk/clustal.
2
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Bioinformatics Tools for the Multilocus Phylogenetic Analysis of Fungi
11. Define the location of each gene by the command CHARSET (see Note 4).
12. Divide sequences into genes specified by
CHARSET by activating the “partition”
command (“set partition = gene”).
13. Set the sequence-species relationship by the
command “TAXSET” which tells the program which sequences belong to which
species.
14. Specify the substitution models for genes for
each partition by the command “lset”
15. Set priors for the parameters in the substitution model.
16. Set priors for the species tree, mutation rates
across genes and population sizes in the command “prset.”
17. BEST 2.3 produces .p, .mcmc and .t files for
which the description can be found in the
MrBayes manual with species trees generated from the posterior distribution and saved
in .sptree format.
18. For multiple runs, it also produces a .sptree
file for each run.
19. Summarize the estimated posterior distribution
of the species tree by the command “sumt.”
MrBayes
MrBayes (version 3.1) is a program for the
Bayesian inference of phylogeny that utilizes
MCMC simulation [71] in combination with the
chosen model and data to estimate the posterior
probability distribution of trees, which eliminates
much of the complex summation and integration
and leaves comparatively simple calculations [72].
Typically, the posterior probability of phylogenies
cannot be calculated analytically. However, it can
be approximated by sampling trees from the posterior probability distribution. MCMC can be used
to sample phylogenies according to their posterior
probabilities and the Metropolis-Hastings-Green
algorithm [71, 73] has now been used successfully
to approximate the posterior probabilities of trees
[74, 75]. This plain-vanilla program has a command-line interface to run on a variety of computer
platforms including Macintosh, Windows, and
Unix. Depending on the size of the data matrix, the
585
computer should be reasonably fast and should
have a lot of memory. The program implements a
wide variety of evolutionary models by generating
phylogenetic trees for nucleotide, amino acid,
restriction site (binary), and standard discrete data
[65, 70, 76, 77].
1. Download and install the current version of
MrBayes.4
2. Start by double-clicking the MrBayes application icon or type “./mb” to execute the program (see Note 5).
3. Change the size of MrBayes window to make
it easier to read the output.
4. Type
“execute
fungi.nex”
at
the
MrBayes > prompt to bring the data into the
program (see Note 6).
5. Type “lset nst = 6 rates = invgamma” at the
MrBayes > prompt, to specify the evolutionary model that will be used in the analysis
with gamma-distributed rate variation across
sites and a proportion of invariable sites.
6. Set the priors of topology, branch lengths,
four stationary frequencies of the nucleotides, six different nucleotide substitution
rates, proportion of invariable sites and the
shape parameter of the gamma distribution
of rate variation for the model.
7. Type “showmodel” to check the model before
starting the analysis which will give an overview of the model settings.
8. Review the run settings by typing “help
mcmc” to start the analysis.
9. Type “mcmc” to run the analysis and print
the state of the chains for every 100th generation after the initial log likelihoods.
10. Type “mcmc ngen = 10,000 samplefreq = 10”
at the MrBayes > prompt, which will ensure
to get at least 1,000 samples from the posterior probability distribution.
11. Stop the run by answering “no” when the program asks “Continue the analysis? (yes/no)”
if the standard deviation of split frequencies
is below 0.01 after 100,000 generations.
Available from http://morphbank.ebc.uu.se/mrbayes/.
4
586
12. Type “sump burnin = 250” (value corresponds to 25 % of samples) to summarize the
parameter values as table including mean,
mode, and 95 % credibility interval of each
parameter.
13. Type “sumt burnin = 250” (or value corresponds to 25 % of samples) to summarize the
trees as cladogram with posterior probabilities for each split and a phylogram with mean
branch lengths.
14. Print the trees as a file that can be read by
tree drawing programs such as TreeView,
MacClade, and Mesquite.
Phylogenetic Analysis Using
Parsimony
Phylogenetic Analysis Using Parsimony (PAUP*)
(version 4.0) is one of the most widely used gold
standard optimization methods for phylogenetic
tree reconstruction using algorithms such as
UPGMA and neighbor joining through parsimony, distance-based and maximum likelihood
methods [78]. It is a commercial program available in variety of platforms and licensed by
Sinauer Associates, Sunderland, MA 5 for inferring and interpreting phylogenetic trees in both
interactive and batch mode. The basic procedure
uses the parsimony approach to infer phylogenetic tree and can be used for DNA, RNA, and
protein sequences [79]. The process of maximum-likelihood tree search using PAUP* includes
essential steps as getting a tree, selecting a model
of DNA substitution, and searching for the optimal tree under the selected model. Moreover, this
program can be used to infer phylogenetic trees
using several maximum-likelihood models for
DNA and RNA sequences only, but not provide
maximum-likelihood models for amino acid substitution. Additionally, it also includes many useful tools for visualizing and examining trees. The
initial data file with aligned DNA sequences in
NEXUS, PHYLIP, or PIR format can be recog-
5
http://www.sinauer.com.
D. Thangadurai and J. Sangeetha
nized by PAUP* [80]. PAUP* default setting are
chosen in general as they are compatible with all
the data types which can be read by the program.
In addition to reconstructing phylogenies, the
program is also useful in diagnosing characters,
inferring ancestral states, and testing the robustness of phylogenetic trees using several statistical techniques [81–87].
1. Purchase and download the most current version of PAUP.6
2. Double-click on the application file to start
PAUP* in Macintosh and Windows or type
“./paup,” where as type “paupx.x” in the
command line for Unix-like environments.
3. Set the initial mode to Execute and then
select the batch file modelblock.
4. Select Import Data from the File Menu and
execute NEXUS file with aligned sequences
in the PAUP* interface (File > Execute).
5. Specify the sites to be included in the analysis by excluding noncoding regions (include
coding/only;).
6. Specify the sequences (taxa) to be used.
7. Set an optimality criterion to parsimony for
selecting a tree and define assumptions (set
criterion = parsimony;).
8. Set character weighting (weights 2:2ndpos;)
and character types (ctype 2_1:all;).
9. Check current character settings (cstatus;)
before going on to the search for a tree.
10. Define search strategy as exact and heuristic
(hsearch addseq = random;).
11. PAUP* will start processing the command
modelblock first to estimate a neighborjoining tree.
12. PAUP* will then calculate the parameter
estimates corresponding to the best-fit model
based on likelihood scores for 56 substitution models which will take few minutes to
even days depending on the complexity of
the data.
13. PAUP* will create two files during the run as
sample.scores and sample.log, which is
intended to check everything has proceeded
normally.
Available from http://paup.csit.fsu.edu/index.html.
6
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Bioinformatics Tools for the Multilocus Phylogenetic Analysis of Fungi
14. Define the outgroup sequence and display
the tree found by the search (show trees all;)
with the branching order of the sequences.
15. Save or print a high-resolution tree (savetrees
file = parsTree.tre brlens = yes;) and exit the
program (quit;).
16. Interpret the results to find out the best-fit
model of nucleotide substitution.
MacClade
MacClade (version 4.08) is a compact, easyto-use phylogenetic program written by David
Maddison and Wayne Maddison for the purpose
of analyzing character evolution, entering and
editing of phylogenetic data for MSAs and for
producing tree diagrams and charts. It has been
widely used to map matrotrophy indices onto the
mitochondrial DNA phylogeny and for assembling data matrices for downstream phylogenetic
analyses. It allows to move branches of the tree
around, build own trees, explore evolutionary
patterns, and trace the evolution of characters on
different trees [88]. The program provides an
interactive environment to manipulate hypothesized phylogenetic trees and to visualize character evolution upon them. In order to manipulate
the tree, many tools are provided to move
branches, reroot clades, create polytomies, and
search automatically for more parsimonious trees
[89]. The summaries of changes in all characters
are depicted on the tree in graphics and charts
with statistics such as number of trees of each
length, number of characters on the tree with different consistency indices and so on. There are
also charts exclusively designed for DNA/RNA
sequence data showing the number of changes on
the tree, codon positions, and the relative frequencies of various transitions and transversions
[90–93]. The MacClade’s data editor has numerous features including rows, columns, and blocks
of data for manipulating and recoding systematic
and comparative data in tune with the abundance
of molecular data. Several display features and
tools were also available in the editor specifically
for graphical manipulation and alignment of
molecular sequences [94–101].
587
1. Download MacClade.7
2. Open the MacClade 4.08 folder and then
double-click on the sample file.
3. Import the file by opening it in MacClade.
4. Data matrix appears something like an Excel
file with taxa in the left column, characters
along the top of each column (indicated by a
number) and character states (T, A, C, or G)
in each cell.
5. Check the entire data set using the scroll bar
at the bottom.
6. Click the Display menu and select Go To
Tree Window to see the tree.
7. Choose the file that represents the most parsimonious tree by selecting “Open tree file”
in the dialog box.
8. Click the “∑” menu to select “Tree Changes”
and repeat with Consistency Index (see Note 7).
9. Click the “Trace” menu and select “Trace
Character” which will highlight each branch
on which the different character states have
evolved in color.
10. Drag the branches around the screen to create new tree topologies.
11. Click through the characters until getting an
informative character with two colors on the
tree.
12. Examine the total number of changes that
occur along each branch by going to the
Trace menu and choosing “Trace all changes”
which will be illustrated by colors on different branches.
13. Alternatively, view the number of changes
directly by changing the “trace all changes
option” under the Trace menu.
14. Click “graphics options” and change the setting to “label by amount of change.”
15. Print out the tree with the amounts of change
labeled.
16. Identify the two “longest” and two “shortest”
branches in the printout.
17. Pull down the Trees menu and select Save
Trees at the bottom to save the tree.
18. Choose Quit from the File menu to quit
MacClade.
7
Freely available from http://macclade.org/download.html.
588
TREE-PUZZLE
TREE-PUZZLE (version 5.2) is a fast tree search
algorithm that provides methods for reconstruction, comparison and testing of trees and models
on DNA as well as protein sequences using quartets and parallel computing [102, 103]. The program computes pairwise maximum likelihood
distances as well as branch lengths with and without the molecular-clock assumption for user
specified trees [104, 105]. It also offers likelihood mapping to visualize the phylogenetic content of a sequence alignment by investigating the
support of internal branches without computing
an overall tree and conducts chi-square test for
homogeneity of base composition, likelihood
ratio to test clock hypothesis, one and two-sided
Kishino-Hasegawa test, Shimodaira-Hasegawa
test, and Expected Likelihood Weights on the
data set [106–109]. In addition, the program
computes the number and the percentage of completely unresolved maximum likelihood quartets
which is an indicator of the suitability of the data
for phylogenetic analysis [110]. In order to avoid
overflow of internal integer variables, the program has a built-in limit to allow data sets only
up to 257 sequences and TREE-PUZZLE terminates the program execution if problems are
encountered during the data analysis [111–113].
1. Download and install the recent version of
TREE-PUZZLE.8
2. Type “b” to switch between tree reconstruction by maximum likelihood and likelihood
mapping.
3. Open the “data” directory to load sequence
input file in CLUSTAL W or PHYLIP output
format or tree input file in DRAWTREE or
DRAWGRAM output format.
4. Type “y” at the input prompt to start the
analysis.
5. TREE-PUZZLE computes pairwise maximum
likelihood distances for all the sequences in
the data file automatically.
D. Thangadurai and J. Sangeetha
6. TREE-PUZZLE displays quartet puzzling tree
with its support values and maximum likelihood branch lengths in “INFILENAME.
puzzle”/“outfile.”
7. View the tree both with its branch lengths and
with the support values for the internal
branches using TreeView and TreeTool.
8. Print the tree topology along with branch
lengths.
9. Type “q” to quit the analysis.
TreeView
TreeView (version 1.6) is a simple program useful for displaying and printing phylogenies. This
software runs on almost all identical interfaces,
reads many different tree file formats including
NEXUS, PHYLIP, Hennig86, NONA, MEGA
and CLUSTAL W and supports native graphics
file format for copying pictures into other applications and for saving graphics files. The current
version reads trees with up to 1,000 taxa and provides tree editor tools for moving branches,
rerooting, polytomy formation, and rearranging
the appearance of the tree. TreeView also displays a scale bar in the bottom left corner of the
tree window if the tree being viewed as branch
lengths, unrooted or as a phylogram [114].
1. Obtain the current version of TREEVIEW.9
2. Double-click on the data file icon to launch
TREEVIEW.
3. Right click on TREEVIEW document to display context menu containing both an Open
and an Edit command.
4. Load the file into TREEVIEW by choosing
“Open.”
5. TREEVIEW will display a single tree in
the tree window with tree’s name on the
status bar.
6. Enable “Previous” and “Next” buttons to
browse among the trees in the file, if it contains more than one tree.
Available from http://taxonomy.zoology.gla.ac.uk/rod/
treeview.html.
9
8
Available from http://www.tree-puzzle.de/.
57
Bioinformatics Tools for the Multilocus Phylogenetic Analysis of Fungi
7. Choose “Edit” to open the file in the program
Notepad to quickly see the tree file in a text
editor.
8. Click tree editor to display the tools to move
branches, reroot, form polytomy, and rearrange the appearance of the tree.
9. Click on the icon to select a tool which will
change to the appropriate shape.
10. Click the cursor on a branch and drag it to the
new position to manipulate the tree.
11. Specify the font used as Plain, Bold, Italic,
Size, Font Type when drawing the tree.
12. Choose “Undo” from the “Edit” menu to
undo a change and recover the old tree, if
essential (see Note 8).
13. Save trees to different file formats.
14. Click Print preview to view how the tree will
appear on the printed page.
15. Set the page orientation and choose a printer
to print the tree.
16. Click Quit to exit TREEVIEW.
Notes
1. User will get two output files as a multiple
alignment (sample_seq1.aln) and a dendrogram (sample_seq1.dnd).
2. Make sure that Architecture in the file Makefile
is correctly set to the platform available.
3. If the dataset contains both diploid genes
(nucleotide DNA) and haploid genes
(mtDNA).
4. For example, CHARSET gene1 = 1–400 which
indicates the first 400 nucleotides belong to
the gene, gene1.
5. Type “help” or “help <command>” for information on commands that are available.
6. The data file (fungi.nex) must be in the same
directory in the MrBayes program and must
have aligned nucleotide or amino acid
sequences, morphological (“standard”) data,
restriction site (binary) data or any mix of
these data types in Nexus file format.
7. Consistency Index is a ratio of the number of
characters in the data set to the length of the
current tree displayed and measures how good
the tree is at evolving those characters.
589
8. Undo is only available for move branch,
collapse branch, collapse clade and reroot;
cosmetic changes such as rotate branches cannot be undone.
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Index
A
ADCC. See Antibody-dependent cell cytotoxicity
(ADCC)
Aflatoxigenic molds
aflatoxin-secreting fungi, 496
chemicals and glasswares, 496
consensus sequence, 496
equipment, 496
multiple sequence alignment and primer
design, 497
nucleic acid-based amplification process, 495
one substance one assay concept, 495
PCR amplification, 497–498
PCR reagents, 496
Aflatoxins, 75
AFM. See Atomic force microscopy (AFM)
agsA gene induction
cell wall and membrane susceptibility testing, 229
inhibitory testing, 227–228
materials, 226–227
Sytox Green assay, 228–229
Airborne fungal spore
Air-O-Cell cassettes, 125–126
allergenco sampler, 126
burkard sampler, 126–127
identification of, 123
25-mm mixed cellulose ester (MCE) filter
cassette, 127
reference data collection, 123, 125
spore traps identification
pass method, 123, 124
random and zigzag field methods, 123, 125
sample trace reading methods, 123, 124
Air sampling
airborne microbial food contamination, 340
fungal strains, 340
health effects, 339
materials, 340–341
microorganisms, 340
microscopic organisms, 339
mycological samples
DNA extraction, 342
identification of, 341–342
isolation and enumeration, 341
RT-PCR quantitation, 342
passive and active, 340
portable SAS-super-180 sampler, 341
RCS, 341
All-Russian collection of microorganisms
(VKM)
control of viability, 62
cryopreservation protocols
cryovials preparation, 61
cultures preparation, 62
fast cooling rates regimen, 62
filling of vials, 62
filamentous fungi
cryopreservation of, 18–60
freeze-drying of, 61
in sterile soil, 61
freeze-drying protocols
cultures preparation, 63
culture viability, 63
filling of ampoules, 63
lyoprotectant agent, 63
primary drying, 63
secondary drying, 63
in sterile soil, 63–64
storage, 63
vacuum control, 63
freezing protocols, 62
thawing, 62
Alternaria Brassicicola
Alternaria alternata, 397
black spot disease, 395
fungal culture, 398
host-pathogen interface, 395
Maganporthe grisea, 397
materials, 396–397
melanin, 396
MPSS, 395
necrotrophic fungus, 395
protoplast production, 396
purify tagged proteins, 396, 397
SAGE and CAGE, 395
targeted gene knockout methods, 395–396
transformation, 398–400
Amerospores, 334
Antibody-dependent cell cytotoxicity
(ADCC), 408
V.K. Gupta et al. (eds.), Laboratory Protocols in Fungal Biology: Current Methods in Fungal Biology,
Fungal Biology, DOI 10.1007/978-1-4614-2356-0, © Springer Science+Business Media New York 2013
593
594
Antifungal susceptibility testing, 220–223.
See also Candida biofilms formation
ArrayTubeT, 536
Aspergillus flavus, 75, 326–327, 365, 496
Aspergillus fumigatus
amplification and quantification, DNA, 317–319
antifungal drugs, 316
BAL, 315–316
DNA Preparation, 316–317
fungal burden and load, 316
immunocompromised patients, 315, 316
intensive cytotoxic chemotherapy, 315
LightCycler PCR Assay, 319–320
LightCyclerT technology, 316, 317
materials, 316
neutropenic patients, 320
Aspergillus niger, agsA gene induction
cell wall and membrane susceptibility testing, 229
inhibitory testing, 227–228
materials, 226–227
Sytox Green assay, 228–229
Atomic force microscopy (AFM)
hyphal viscoelasticity determination, 157–158
imaging spores and fixed hyphae, 155, 156
live hyphae, 155–157
polysaccharides, 191
surface adhesion determination, 158
B
BAL. See Bronchoalveolar lavage (BAL)
Basic local alignment search tool (BLAST), 234
Bayesian Estimation of Species Trees (BEST), 584–585
BEST. See Bayesian Estimation of Species Trees (BEST)
Biochemical methods
automated
abbott yeast identification system, 251
API 20C yeast identification system, 250
biomerieux vitek yeast biochemical card,
250–251
manual
b(beta)-glucosidase activity, 249
carbohydrate fermentation, 247–248
carbon sources utilization, 246–247
casein hydrolysis, 248
cellulose hydrolysis, 248
cyclohexamide resistance, 248
fatty acid esterase activity, 248–249
gelatin hydrolysis, 249
lipase activity, 249
nitrogen sources utilization, 247
urease activity, 249
Bioinformatic protocols. See Secreted proteins
Biosafety levels (BSL)
accidents, 6
BSL-1, 2
BSL-2, 2–3
BSL-3, 3
emergency preparedness and response, 6
experimentation, 5–6
Index
information and training, 6–7
materials, 3
risk assessment for, 3, 4
routine precautions, 4–5
Black spot disease, 395
BLAST Search, 556
BRITE objects, 562
Bronchoalveolar lavage (BAL), 315–316
BSL. See Biosafety levels (BSL)
C
CAGE. See Cap analysis of gene expression (CAGE)
Candida biofilms formation
materials, 218–219
on 96 well microtiter plates
antifungal susceptibility testing, 220–221
schematic diagram of, 219, 220
XTT-reduction assay usage, 221
Cap analysis of gene expression (CAGE), 395
Capillary electrophoresis (CE), 352
CARD-FISH. See Catalyzed reporter deposition-FISH
(CARD-FISH)
Catalyzed reporter deposition-FISH (CARD-FISH)
classical FISH probing, 2
materials, 233
CE. See Capillary electrophoresis (CE)
CFU. See Colony forming units (CFU)
Chytrids
calcofluor stock solution, 171
cells concentrations, 171
materials, 171
staining and visualization, 171
CLUSTAl W program, 583
Colletotrichum spp., 404
Colletrotrichum graminicola, 383
Colony forming units (CFU)
fungal culturing approach
assessing and counting, 205
dichloran and rose bengal, 204
medium pH, 204
membrane filtration (MF) technique, 203–204
pour plate technique, 203
sabourand dextrose agar (SDA), 204
spread plate technique, 203
materials, 206
media, 205–206
techniques
counting, 207
incubation conditions, 207
MF technique, 207
pour plate technique, 207
spread plate technique, 206–207
Comet assay, 501
Community Annotation, 556
Compound microscopes
alignment and setting-up, 120–121
eyepieces, 121
ocular micrometer, 120
Conidia, 304
Index
Coomassie dye-binding assay
colorimetric/fluorescent labels, 309
endoprotease assay, 311, 312
endoproteolytic activity, fungus, 309
exopeptidase activity, 309
fluorescently labeled proteins, 310
fungal culture and enzyme sample preparation,
310–311
inhibition assay, 312
materials, 310
protease cleavage calculation, 311–312
protein substrate standard curve, 311
proteolytic cleavage, 309
proteolytic enzymes isolated, 309
Talaromyces emersonii, 310
traditional techniques, 309
Crustose lichens, 94, 100–109
Cryofixation and freeze-substitution technique, 142–143
Cryofracture
ethanol method, 137–138
glycerol method, 136, 137
Cryopreservation
culture collections, 9
culture preparation and freezing-thawing
protocols, 13
freeze-drying, 10
freezing protocols, 11
genetic and physiological features preservation, 11
growth estimation and morphological analysis, 13–14
Hoffmann’s technique vs. agar blocks, 10
lyophilization, 10
materials for, 12
perlite protocol (PP), 12
protocols
cryovials preparation, 61
cultures preparation, 62
fast cooling rates regimen, 62
filling of vials, 62
strains (cultures), 13
survival of, 11
thawing, 11
viability test, 13
Cryptococcus neoformans, 383
D
Deletions via Gateway (DelsGate)
ccdB gene, 377
Gateway cloning technology, 377
materials
bacterial strains transformation, 378
BP clonase reaction, 378
culture media, 377–378
deletion construct verification, 378–380
fungal transformation (see Fungal transformation)
PCR amplification, 378
methods
A. tumefaciens transformation, 389–391
BP clonase reaction, 387
Colletrotrichum graminicola, 383
595
Cryptococcus neoformans, 383
deletion constructs verification,
379, 383–384, 388
E. coli transformation, 382
Fusarium verticillioides, 383
gene flanks amplification, 379, 385–386
generate deletion mutants, 382–384
PCR amplification, 386–387
protoplast-mediated fungal transformation,
389–390
Ustilago maydis, 376, 377
Denaturing gradient gel electrophoresis (DGGE)
equipment and materials, 292
fingerprinting
gel casting, 293, 295
gel staining, drying, and scanning, 295–296
pre-run, 295
sample loading and electrophoresis, 295
initial material, 292
semi-nested PCR
first amplification step, 292–293
second amplification step, 293
Deoxynivalenol, 76–77
Desiccator method, 141, 142
DGGE. See Denaturing gradient gel electrophoresis
(DGGE)
DNA isolation method
Fusarium spp., 404–405
materials, 404
molecular manipulation techniques, 403
DNA microarray-based detection
ArrayTubeT, 536
DNA extraction, 540
fungal diagnostics and microarray, 536
fungal panels, 536
fungal PCR Preparation, 539
fungal species, 539
HRP, 536
materials, 538–539
MycXtra kit, 541–542
NorDiag Arrow, 540–541
NucliSENS easyMAG, 541
PCR-based assays, 535–536
Prove-it Sepsis assay, 537–540
Drug-induced permeabilization
amphotericin B binds, 520
cell lytic enzyme or zymolyase, 519
ERG6 mutations, 520
sesqueterpene dialdehyde polygodial, 520
yeast cells, 519
yeast permeabilization, 520–521
dsDNA quantification
Comet assay, 501
DNA damage, 501
ethidium bromide staining, 501
fragmented and intact, 503–504
fragmented DNA stock solution, 502
fungal isolation, 503
isolated DNA sample, 504
isolated fungal concentration, 503
596
dsDNA quantification (cont.)
materials, 502
necrotic and apoptotic, 502
PEG-NaCl stock solution, 502–503
PicoGreen stock solution, 502
standard curve, 503
tris-EDTA (TE) buffer, 502
E
Epifluorescence microscopy. See also Chytrids
calcofluor stock solution, 171
cells concentrations, 171
materials, 171
staining and visualization, 171
EPS. See Exopolysaccharide (EPS)
ESLpred method, 573
Exopolysaccharide (EPS)
alcohol precipitation, 427
materials, 428–429
membrane proteins extraction, 431–432
nitrile, 433
nucleic acid and protein extraction, 428
protein isolation method, 431
RNA, DNA, and proteins, 427–428
Sclerotium rolfsii
CsCl-Pads, 430–431
cultivation, 428
genomic DNA, 429–430
transcriptomic and proteomic analyses, 428
soluble and insoluble proteins extraction, 432
F
FASTA download, 556
Filamentous fungi
Agrobacterium-mediated transformation, 345
cryopreservation, 18–60
electroporation, 345
freeze-drying of, 61
genetic transformation, 345
materials, 345–346
particle bombardment, 345
PEG-mediated protoplast transformation, 345
Penicillium chrysogenum, 345
protoplastation, 346
regeneration, 347–349
in sterile soil, 61
transformation, 346–347
FLA. See Fragment length analysis (FLA)
Foliose lichens, 94–97
Force spectroscopy (FS)
cleaning coverslips, 153
fixed and dehydrated fungal hyphae
preparation
formaldehyde fixation, 154
OsO4 fixation, 154–155
viable fungal hyphae, 155
fungal spores preparation, 153
live and fixed hyphae, 156–157
materials, 152–153
Index
silanization, 153
spore sample preparation, 153
Fourier-transform infrared (FTIR) microscopy
infected tissue, 162
in liquid media, 162
materials, 162
slides preparation, 163
on solid media, 162
spectra analysis, 164, 165
spectra measurement, 163, 164
statistical analysis
cluster analysis, 164, 166
principal component analysis, 166
FragAnchor, 552
Fragment length analysis (FLA), 352
Freeze-drying protocols
cultures preparation, 63
culture viability, 63
filling of ampoules, 63
lyoprotectant agent, 63
primary drying, 63
secondary drying, 63
in sterile soil, 63–64
storage, 63
vacuum control, 63
Fruticose lichens, 94, 98–99
FS. See Force spectroscopy (FS)
Fumonisins, 76
Fungal cell wall
b(1,3)-glucan synthase (GS)
CHAPS, 180
membrane extracts preparation, 179
microtiter-based fluorescence assay, 180–181
permeabilized whole cells, 181
protein assay, 179–180
Zymolyase-100T degradation, 180
chitin synthase (CS)
CS assay, 181–182
membrane extracts preparation, 181
microtiter-based fluorescence assay, 182
permeabilized whole cell, 182
components
chitin, 177–178
glucan, 177, 178
glycoproteins, 178–179
polysaccharides
alkali-fractionation, 184–185
AMF, 191
b(1,6) glucan determination, 187–188
cell wall labeling, 182–183
chemical fractionation, 187
colorimetric determination, chitin, 187
cryoscanning and cryosectioning electron
microscopy, 191
electron microscopy (EM), 190
enzymatic degradation, 189
enzymatic fractionation, 185–187
fluorescence microscopy, 190
hypersensitivity/resistance, biosynthesis
inhibitors, 190
proteins determination, 188–189
Index
radioactive labeling and fractionation, 182, 183
transmission electron microscopy, 190
transmission immunoelectron microscopy
(IEM), 191
structure and composition of, 176
Fungal genomes
genome and protein sequences, 559
protein level annotation, 560–562
secretory peptides, 560
secretory proteins prediction, 560–563
therapeutic target prediction, 560, 561, 563
translate nucleotide sequences, protein sequences,
560–561
Fungal proteomics. See Proteomic protocols
Fungal Secretome Knowledge-Base (FunSecKB),
546, 554
Fungal transformation
materials
Agrobacterium tumefaciens-mediated, 379,
381–382
confirm gene deletion analysis, 382
DelsGate deletion constructs, 380
protoplast-mediated, 380–381, 389–390
methods
Agrobacterium tumefaciens-mediated, 390–392
confirm gene deletion analysis, 392
DelsGate deletion construct preparation, 388–389
protoplast-mediated, 389–390
Fungi in water. See Colony forming units
FunSecKB. See Fungal Secretome Knowledge-Base
(FunSecKB)
Fusarium, 197, 198
Fusarium graminearum, 572
Fusarium spp., 404–405
G
Gateway cloning technology, 377
Gene silencing
data analysis
data normalization, 285–286
PCR efficiency, 285
relative expression values, 286
validate reference genes, 286
materials
DNase treatment, 281–282
qPCR quantification and analysis, 282
RNA concentration and quality, 281
RNA isolation, 281
reverse transcription
cDNA synthesis, 283–284
DNase treatment, 283
quantitative polymerase chain reaction, 284–285
RNA concentration and quality, 282–283
RNA isolation, 282
454 Genome Sequencer FLX, 358
Genome shuffling protocol
materials
random mutagenesis, 448
suitable selection regimes, 448, 449
yeast mating, 448
597
methods
random mutagenesis procedure, 450–452
yeast mating optimization, 452–453
microbial strain improvement technology, 448
recursive protoplast fusion/cross-mating, 448
GFP. See Green fluorescent protein (GFP)
Gliocladium roseum, 404
Good manufacturing practice (GMP) bioprocess
scale, 407
Green fluorescent protein (GFP), 468
H
Hardwood spent sulphite liquor (HW SSL). See Genome
shuffling protocol
Heterotrimeric G proteins, 573
Heterotrophic nanofiagellates (HNFs), 422
High-performance liquid chromatography (HPLC)
aflatoxins detection, 86
deoxynivalenol detection, 87
fumonisins detection, 87–88
ochratoxins detection, 86–87
Histopathological technique
dehydration, 198
infiltration and embedding, 199
killing and fixation, 198
materials, 198
microscopy and imaging, 199–200
sectioning, 199
staining and mounting, 199
HNFs. See Heterotrophic nanofiagellates (HNFs)
Horseradish peroxidase (HRP), 536
HPLC. See High-performance liquid chromatography
(HPLC)
HRP. See Horseradish peroxidase (HRP)
I
IMAC. See Immobilized metal ion affinity
chromatography (IMAC)
Immobilized metal ion affinity chromatography
(IMAC)
biomass, 415–416
copper priming and equilibrating, 415
deionized water, 415
downstream processing, 416
eluting target protein, matrix, 416
expanded and radial flow bed adsorption IMAC,
411–413
fermentation, 411
NaOH, 414
nonspecifically bound contaminants, 416, 417
plasmid construction, 411
Proxcys AXCIS MD 62 MKIII manual, 413
regeneration, 416
seed lot production, 411, 412
Watson Marlow pump, 414, 415
Internal transcribed spacer (ITS)
amplified products detection, 325–326
internal transcribed spacer region, 324–325
molecular methods, 323
598
Internal transcribed spacer (ITS) (cont.)
PCR-based DNA sequencing targeting, 328–329
PCR-based restriction fragment length
polymorphism, 329–331
semi-nested polymerase chain reaction
amplified products detection, 325–326
Aspergillus flavus, 327
DNA extraction, 327
DNA sequencing, 327–328
fungal genome detection, 326
materials, 324
multicopy genes, 324
nucleotide polymorphisms, 326–327
RFLP, 327
targeting ITS region, 326
Isoelectrofocusing, 305
Isolation
antibiotics inclusion, 68
C:N ratio, 68
colony identification, 71
culture identification
cold temperature, 71
direct culture methods, 71
freeze drying/lyophilization, 72
fungi storage, 71
low-temperature storage, 72
under oil, 71
storage in water, 71
cultured sample data, 70–71
culturing, 70
low pH media, 68
materials for, 69
nonselective media, 67
sample collection, 69
plates preparation, 70
sample dilution, 69–70
saprotrophic fungi, 68
selective media, 68
spores, 68
ITS. See Internal transcribed spacer (ITS)
K
KAAS mapping, 562
KEGG orthology (KO), 562
Kernel-based machine learning algorithm, 574
L
Lasiodiplodia theobromae, 404
Lichenized fungi
identification keys
crustose lichens, 94, 100–109
foliose lichens, 94–97
fruticose lichens, 94, 98–99
squamulose lichens, 94, 109–110
materials, 92–93
mycobionts isolation, thallus, 93
spore discharge method, 93
Index
LightCyclerT technology, 316, 317
Lignocellulolytic enzymes. See Talaromyces
emersonii
Lignocellulosic biomass
biofuels, 475
biological pretreatments, 476
biorefinery conversion technology, 476
biorefinery strategy, 475
cellulose and hemicellulose, 476
commercial enzymes, 477
enzymatic saccharification, 476, 478
enzyme cocktails, 478
enzyme dosage optimization, 478
enzyme pretreated biomass, 479
feedstock collection and characterization, 477
fermentation/thermochemical processing, 476
lignin-derived compounds analysis, 479
materials, 477
microbial fermentation and enzymatic
hydrolysis, 476
petroleum-derived energy sources, 475
pH optimization, 478
plant biomass, 475
pretreatment technologies, 476
reducing sugars, hydrolysates, 479
temperature optimization, 478
total sugars quantification, hydrolysates, 479
Linear temperature programmed retention indices
(LTPRIs)
alkane standards calibration, 461
automated peak detection, deconvolution, 461–462
deconvolution program AMDIS, 462
GC-MS measurements, alkane standards, 459–461
reference database, 461
Liquid auxanographic method, 246
LTPRIs. See Linear temperature programmed retention
indices (LTPRIs)
Luria-Bertani medium preparation, 436
Lyophilization, 10
M
MacClade (version 4.08), 587
Maganporthe grisea, 397
Massively parallel signature sequencing (MPSS),
395–396
Melanin, 396
Microbial volatile organic compounds (MVOCs)
C8-compounds, 455
filamentous fungi produce, 455
fungal cultures, 458, 459
Fusarium graminearum, 457, 458
GC-MS, 456
HS-SPME-GC-MS analysis, filamentous fungi,
458–460
LTPRI values (see Linear temperature programmed
retention indices (LTPRIs))
materials, 456–457
metabolites, 455
Index
PDA plates and slants preparation, headspace
vials, 457–458
Trichoderma atroviride, 457, 458
Microsatellite markers
equipment and consumables, 358
eukaryotes, 357
genome mapping and population genetics, 357
454 Genome Sequencer FLX, 358
genomic DNA sequences, 358
media and reagents, 358–359
microsatellite discovery and primer design, 360
NGS technology, 358
non-model organisms, 357
PCR amplification, 360
polymorphism analysis, 361
Sanger sequencing technology, 357–358
Shotgun pyrosequencing, 359–360
traditional methods, 357
Microtiter plates. See Filamentous fungi
Molecular fingerprinting
CE-SSCP, 352
data analysis, 354
DNA extraction, 353
fungal biomass, 351
ITS1 fragments, PCR, 353–354
materials, 352
PCR amplicons, 352
phylogenetic structure, 351
RFLP, 352
snapshot of fungal diversity., 352
soil fungal communities, 351
soil sampling, 353
MPSS. See Massively parallel signature sequencing
(MPSS)
MrBayes program, 585–586
MudPIT. See Multidimensional Protein Identification
Technology (MudPIT)
Multidimensional Protein Identification Technology
(MudPIT), 302
Multilocus phylogenetic analysis
BEST program, 584–585
bioinformatics tools and techniques, 580–582
CLUSTAl W program, 583
conventional biochemical methods, 579
fungal diversity, 580
MacClade, 587
materials, 582
MrBayes program, 585–586
MSA method, 582
PAUP*, 586–587
T-Coffee program, 584
TREE-PUZZLE program, 588
TreeView program, 588–589
MVOCs. See Microbial volatile organic compounds
(MVOCs)
Mycelium, 303–304
Mycotoxigenic fungi
array construction, 533
control probes preparation, 533
culture media, 530
599
DNA extraction buffer, 530
food contaminants, 529
fungal DNA extraction, 531
fungal strains, 530
hybridization, array slides, 533–534
ITS, 530
labelled target DNA preparation, 533
microarray technology, 530
molecular biology techniques, 529
oligonucleotide probe, 533
PCR amplification, 531
PCR detection assays, 529–530
probe sequences, 531, 532
scanning and data processing, 534
Mycotoxins
aflatoxins, 75
chromatography-based method
aflatoxin detection, 83
DON detection, 84–85
extraction and clean-up, 83
fumonisins detection, 85, 86
HPLC (see High-performance liquid
chromatography)
ochratoxin detection, 84
sample extraction and clean-up, 81–82
thin-layer chromatography, 82–83
TLC plate, 83–84
deoxynivalenol, 76–77
DNA extraction
contaminated food samples, 78–79
pure cultures, 78
DNA fingerprinting methods, 78
fumonisins, 76
ochratoxin, 75
polymerase chain reaction
aflatoxigenic fungal species, detection of, 79, 80
aflatoxin producer, 79
assay set-up, 77
equipments for, 77
fumonisins producing fusarium species, 81, 82
ochratoxigenic fungi, detection of, 79–80
reagents, 77
target genes, amplification, 78
toxigenic fungi, detection of, 79, 80
tricothecene-producing fusarium species, 80–81
MycXtra kit, 541–542
N
Neotyphodium, 467
Next-generation sequencing (NGS) technology, 358
NorDiag Arrow, 540–541
NucliSENS easyMAG, 541
O
Ochratoxin, 75
Oligonucleotide microarray.
See Mycotoxigenic fungi
One-dimensional electrophoresis, 305
600
One-step construction of agrobacterium-recombinationready plasmids (OSCAR)
Gateway cloning technology, 377
materials
bacterial strains transformation, 378
BP clonase reaction, 378
culture media, 377–378
deletion construct verification, 378–380
fungal transformation (see Fungal transformation)
PCR amplification, 378
methods
attB sites, 383
A. tumefaciens transformation, 389–391
BP clonase reaction, 387
confirm gene deletion analysis, 391–392
deletion constructs verification,
379, 383–384, 388
E. coli transformation, 382, 387
gene flanks amplification, 379, 385–386
generate deletion mutants, 382–385
PCR amplification, 386–387
pOSCAR and pA-Hyg-OSCAR vectors, 384
protoplast-mediated fungal transformation,
389–390
T-DNA borders, 384
MultiSite Gateway system, 377
Verticillium dahliae, 376, 377
Optical microscope
airborne fungal spore (see Airborne fungal spore)
compound microscopes, 118–119
illumination techniques
bright field microscopy, 119–120
differential interference contrast, 119, 120
phase contrast, 119, 120
light microscope, 121
photomicrographic system and usage, 121–122
sample mounting
squash/tease mounts, 116–117
staining, 117
tapelift mounts, 115–116
wet mounts, 116
sectioning
freehand, 114–115
freezing microtome, 115
specimen preparation, 114
stereo microscopes, 117–118
Optimal Separating Hyperplane, 566
OSCAR. See One-step construction of agrobacteriumrecombination-ready plasmids (OSCAR)
Osmium tetroxide vapor technique, 140–141
P
Padlock probes
fungal species, 363
Luminex suspension array, 364
materials
dilution series of standards, 364–367
DNA isolation, 364
ligation and RCA, 365–368
Luminex, 368–369
Index
PCR, 368
ultrapure water, 364
method
Candida glabrata sequence, 371, 372
DNA isolation, 369–370
ligation and RCA, 370
Luminex 200 instrument measurements, 371
MFI, 371
PCR, 370–371
oligonucleotide designed, 363
pathogenic fungi
conventional oligonucleotide probe, 507
culture-based morphological features, 507
design of, 509
eukaryotic diversity, 505
fungal pathogens, 507
fungal pathogens detection, 506, 508
ligation detection assay, 512–514
materials, 507, 509
OpenArrayr instrument, 510–512
plant and tree species, 506
RCA, 509–510
routine DNA sequencing, 507
SNPs, 507
target-complementary sequence regions, 507
suspension array technology, 364
SybrGreenT real-time PCR, 364
target sequence, 364, 365
PEG-NaCl stock solution, 502–503
Penicillium chrysogenum, 345
Pentose-fermenting yeast Scheffersomyces stipitis.
See Genome shuffling protocol
Perlite protocol (PP), 12
Personal Environmental Monitor (PEM), 335
Personal protective equipment (PPE), 4
Phobius, 547, 549–550
Phylogenetic Analysis Using Parsimony (PAUP*),
586–587
Phyto-fungal pathogens. See Fourier-transform infrared
(FTIR) microscopy
Pichia pastoris
ADCC, 408
chemicals, 408
EBA-IMAC, 408
equipment, 408–410
Escherichia coli platform, 407–408
glycoforms, 408
GMP bioprocess scale, 407
IMAC
biomass, 415–416
copper priming and equilibrating, 415
deionized water, 415
downstream processing, 416
eluting target protein, matrix, 416
expanded and radial flow bed adsorption IMAC,
411–413
fermentation, 411
NaOH, 414
nonspecifically bound contaminants, 416, 417
plasmid construction, 411
Proxcys AXCIS MD 62 MKIII manual, 413
Index
regeneration, 416
seed lot production, 411, 412
Watson Marlow pump, 414, 415
materials, 408
posttranslational modifications, 407
yeast production, 407
PicoGreen stock solution, 502
Plant pathogenic fungi, 300
Plant samplings. See Histopathological technique
Polyethylene glycol (PEG)-mediated protoplast
transformation, 345
Portable SAS-super-180 sampler, 341
Pour plate auxanographic method, 246–247
PROSITE, 552
Proteomic protocols
Aspergillus ssp., 301
electrophoresis, 301
extracellular compounds, 301
gel-based analysis, 302
mass spectrometry (MS), 302
materials, 302–303
MudPIT, 302
plant pathogenic fungi, 300
protein concentration, 301
protein precipitation, 301
protein separation, 305
recalcitrant biological material, 300
sample collection, 303
secretomics, 300
staining techniques, 302
TCA/acetone-phenol/methanol method
conidia, 304
mycelium, 303–304
secreted proteins, 304
two-dimensional electrophoresis, 301–302
Tyromyces palustris, 301
Protoplast fusion techniques
commercial lysing enzymes, 483
conventional mating systems, 483
double-stranded RNA mycoviruses, 484
fungal genetic transformation systems, 484
lysed protoplasts, 484
materials, 485
microbial cells, 483
mycelia markedly influences, 484
PEG, 484
Trichothecium roseum, 485–486
Prove-it Sepsis assay, 537–540
Psidium guajava, 197
R
RCA. See Rolling circle amplification (RCA)
RCS. See Reuter centrifugal air sampler (RCS)
Real-time quantitative polymerase chain reaction
(qPCR) assay
accuracy, precision, and method detection
limits, 337
allergens and pathogens, 333
fungal DNA extraction, 335–336
fungal spores quantitation, 336
601
growth-based methods, 333–334
materials, 334
PCR inhibition assay, 336–337
PEM, 335
qPCR-based methods, 333–334
8-stage nonviable Andersen sampler, 335
standard fungal DNA samples, 334–335
Restriction fragment length polymorphism (RFLP),
327, 352
Reuter centrifugal air sampler (RCS), 341
RFLP. See Restriction fragment length polymorphism
(RFLP)
Rhizopus microspores, 536
RNA isolation, S. cerevisiae
batch cultivation, 266, 267
cell size, 267
cell sorting, 272–273
cells staining, 272
chemostat cultivation, 268
cultivation conditions, 272
flow cytometry, 266
materials, 271–272
2-NBDglucose, 267–268
quantity and integrity estimation, 273–274
RNeasy mini kit, 273
substrate concentrations, 268
transient state cultivated live cells, 268, 269
work flow of, 269, 270
Rolling circle amplification (RCA), 509–510
S
Sabouraud agar medium
acidic medium, 212
Aspergillus colonies, 215
history of, 211–212
materials, 212–213
neutral pH, 212
peptones, 212
preparation methods
antimicrobials, 213, 214
Emmons modification preparation, 213–214
ingredients for, 213
inoculation and incubation, 214
sugar maltose, 212
Trichophyton spp, 214, 215
Saccharomyces cerevisiae, RNA isolation
batch cultivation, 266, 267
cell size, 267
cell sorting, 272–273
cells staining, 272
chemostat cultivation, 268
cultivation conditions, 272
flow cytometry, 266
materials, 271–272
2-NBDglucose, 267–268
quantity and integrity estimation, 273–274
RNeasy mini kit, 273
substrate concentrations, 268
transient state cultivated live cells, 268, 269
work flow of, 269, 270
602
SAGE. See Serial analysis of gene expression (SAGE)
Sanger sequencing technology, 357–358
Scanning electron microscopy (SEM)
cellophane membrane, sample preparation, 148
cryochamber usage, 144
cryofixation and freeze-substitution technique,
142–143
cryofracture
ethanol method, 137–138
glycerol method, 136, 137
desiccator method, 141, 142
fixation with tannic acid, 142
internal cellular, fungal fruiting body, 136
leaf epidermis removal, 138–139
materials, 134
microculture usage, 148
mycelia preparation, filter, 148
nematophagous fungi, 147
nuclei and chromosomes, 147–148
osmium tetroxide vapor technique, 140–141
polysaccharides, 191
proteases usage, 139
routine protocol
plant pathogenic fungi., 135, 136
rust fungi, 135
seed-borne fungi identification (see Seed-borne
fungi)
thick section preparation, 138
ultrasonication, 141–143
yeast
poly-L-lysine coverslips, 140
polystyrene surface usage, 139–140
Sclerotium rolfsii, 428–431
Secreted proteins
BLAST Search, 556
cell signaling, 545
Community Annotation, 556
complete secretome/curated proteins, 556
FASTA download, 556
FragAnchor, 552
fungal-specific secretome databases, 546
FunSecKB, 546, 554
Golgi complex, 546
materials, 546
metabolism and regulation in growth, 545
NCBI RefSeq database, 546, 554
Phobius, 547, 549–550
PROSITE, 552
protocol evaluation, 553–554
SecretomeP, 552–553
SignalP 3.0, 547, 548
TargetP, 550
TMHMM 2.0, 550–551
UniProt database, 546
Webservers, 553
WoLF PSORT, 550
Secretome. See Secreted proteins
Index
Seed-borne fungi
blotter test incubation method
individual examination, 146
water restriction, 146
material, 145–146
Serial analysis of gene expression (SAGE), 395
Sesqueterpene dialdehyde polygodial, 520
Shotgun pyrosequencing, 359–360
Single sequence repeats (SSRs). See Microsatellite
markers
Single-stranded conformation polymorphism (SSCP), 352
Slow-growing fungi
DAPI staining, 469–470
fungal strains and culture conditions, 468, 469
GFP, 468
hygromycin B selection, 468–469
materials, 468
microscopy, 470–472
protoplasts, transformation, and regeneration
preparation, 469–470
sporulation and hyphal tipping, 470
toxic locoweeds, 467
transformation procedure, 470
transformed gene/marker, immunoblot detection,
470–472
Solid phase microextraction (SPME), 456
SPME. See Solid phase microextraction (SPME)
Spore discharge method, 93
Squamulose lichens, 94, 109–110
SSCP. See Single-stranded conformation polymorphism
(SSCP)
Staining techniques
acridine orange staining, 243
calcoflour white staining, 243–244
double-oxidation thiosemicarbazide, Schmorl, 245
Fontana-Masson staining, 245–246
giemsa staining, 242
gram staining, 242
gridley staining, 243
Grocott-Gomori methenamine silver staining,
244–245
materials, 239–240
Mayer’s mucicarmine staining, 244
periodic acid-Schiff staining, 244
toluidine blue O staining, 246
Weigert’s iron hematoxylin staining, 243
wet mount techniques, 241
Wright staining, 242–243
Stereo microscopes, 117–118
Superoxide radical production
materials, 259–260
20 mM HE stock solution
alkaline acetone extraction, 263
fluorescence extinction coefficient, 261–262
fluorometric quantification, 264
microcolumn cation exchange
chromatography, 263
Index
preparation of, 260
purification, 263–264
quantification, fluorescence, 260
sample protein determination, 262–263
sample treatment, 262
synthesis of, 260–261
solutions and standard curves, 260
Support vector machines (SVM)
artificial intelligence and machine learning-based
methods, 566
Aspergillu fumigatus, 572
Aspergillus nidulans, 572
Auricularia auricula, 573
bioinformatics applications, 571
Codon composition, 574
dataset collection, 570
ESLpred method, 573
FGsub, 572
fungal adhesins and adhesin-like proteins,
568, 569, 571
fungal classification algorithms, 568, 569
fungal data, 568
genome annotation, 567
genomic technologies, 565
heterotrimeric G proteins, 573
hybrid models, 570
hydrophobicity, 567
microarray data, 573–574
molecular mechanisms, 567
Optimal Separating Hyperplane, 566
Paracoccidioides brasiliensis, 572–573
PPIs prediction, 574
proteome annotation, 567
regulatory analysis, 571–5
software for, 575–576
statistical learning theory, 566
structural risk minimization principle, 566
therapeutic metabolites, 565
transcription factor target prediction, 571–572
type III polyketide synthases, 574
WEKA, 571
wrappers and filters, 568
yeast deletion phenotypes, 571
SVM. See Support vector machines (SVM)
T
Talaromyces emersonii
enzyme activities determination, 492
extracellular polysaccharide, 489
fungal cultivation, 148-L fermenter, 491–492
fungal cultivation monitoring, 493
harvesting and concentrating, enzyme cocktails, 492
inoculum development, 10-L fermenter, 491
materials, 489
plant polymer hydrolysis, 489
pre-inoculum preparation, 491
thermozyme cocktails, 489
thermozymes production, 489
total protein and sugar analysis, 492
603
Targeted gene knockout methods, 395–396
Taxol
cell proliferation, 524
chemotherapic drug, 524
Coelomycetes fungi, 525
endophytic taxol-producing fungus, 524
HPLC, 527
infrared spectroscopic analysis, 526
isolation and identification, endophytic fungi,
524–525
large-scale industrial fermentation, 524
materials, 524
plant pathogenic fungi, 525
selected fungal isolates, 526
single pathogenic fungi, 526
Taxus brevifolia, 524
thin-layer chromatography, 526
ultraviolet spectroscopic analysis, 526
Taxus brevifolia, 524
TCA. See Trichloroa-cetic acid (TCA)
T-Coffee program, 584
Thawing, 11, 62
TMHMM 2.0 program, 550–551
TREE-PUZZLE program, 588
TreeView program, 588–589
Trichloroa-cetic acid (TCA), 301
Trichoderma spp., 404
U
Ultrasonication, 141–143
Uncultured zoosporic fungi. See Catalyzed reporter
deposition-FISH
Undifilum oxytropis. See Slow-growing fungi
UniProt database, 546
Uromyces appendiculatus, 302
Ustilago maydis, 376, 377
W
Waikato Environment for Knowledge Analysis
(WEKA), 571
WEKA. See Waikato Environment for Knowledge
Analysis (WEKA)
Wet mount techniques
India ink, 241
lactophenol cotton blue, 241
potassium hydroxide, 241
WoLF PSORT program, 550
X
Xylanase
cloning and expression, 436
genetic and protein engineering, 436
materials
agarose gel electrophoresis, 437
alkaline screening assay, 439
ampicillin preparation, 436
cloned gene(s), interest and strains, 436
604
Xylanase (cont.)
competent cells preparation, 437
DNA sequence analysis, 440
growth and extraction, 439
ligation, 437
Luria-Bertani medium preparation, 436
mutant library and enzyme extraction growth, 439
plasmid DNA isolation, 437
random mutagenesis, 437, 438
RBB-Xylan-LB Plates preparations, 439
restriction endonuclease digestion, 437
stability, 439–440
staggered extension process, 440
thermostability screening assay, 439
transformation, 439
methods
agarose gel electrophoresis, 441
alkaline screening assay, 443–444
ampicillin preparation, 440
cloning, 440
competent cells preparation, 442
DNA concentration measurement, 441
DNA sequence analysis, 445
growth and extraction, 444
LB medium preparation, 440
mutant library and enzyme extraction growth, 443
Index
plasmid isolation, 440
random mutagenesis, 438, 441
RBB-xylan-LB plates preparation, 442
restriction analysis, 441–442
stability, 444
staggered extension process, 444–445
T4 DNA ligase, 442
thermostability screening assay, 443
transformation, 442–443
protracted spontaneous mutations, 435
Thermomyces lanuginosus, 436
Z
Zoosporic fungi
choanoflagellate-like ancestors, 422
chytrids, 422
DNA extraction and purification, 423–424
environmental samples, 425
HNF, 422
lyticase, 425
materials, 422
microbial eukaryotes, 422
molecular approaches, 422
phenol-chloroform purification procedure, 425
real-time qPCR assays, 424