Metabolism and Infection in the
Stagonospora nodorum-Wheat
Pathosystem
Ormonde Dominick Creagh Waters
Bachelor of Science (Hons) (Murdoch)
Bachelor of Economics (Hons) (UWA)
This thesis is presented for the degree of Doctor of Philosophy
2008
I declare that this thesis is my own account of my own work and contains as its main
content work which has not been previously submitted for any degree at any tertiary
institution.
………………………………………………………
Ormonde Dominick Creagh Waters.
ii
Ode to an Ectopic Fungal Mutant (Pmk1-61)
Thy hyphae fair didst bloom upon my plate
Of medium minimal, yet enough to grow.
And
d with
i h selective
l
i
ffungicides
i id to ensure
Lest non-transformants would contaminate.
In Stygian darkness, but near-UV also
I nourished
i h d you and
d waited
it d you tto spoor.
A picture portrait I did make of you,
Your handsome colours did my eye delight
And I did hope that you might be the one!
An homologous recombinant mutant – Oh so true
On you an Honours chapter I would write
And you a thesis cover would become.
become
Alas! By PCR you proved ectopic
And now you moulder in a bin necrotic.
Ormonde Waters 2007
iii
ABSTRACT
Stagonospora nodorum is a necrotrophic fungal pathogen, and the causal
agent of stagonospora nodorum blotch of wheat. Despite the economic importance of
this disease, the molecular basis of the pathosystem is poorly understood. The aim of
this study was to investigate the interaction between metabolism and infection in this
pathosystem, with particular reference to the metabolism of mannitol.
In common with many fungi, the main metabolite produced by S. nodorum is
the acyclic hexitol mannitol. Among the previously suggested roles for this compound
is a role in pathogenicity. The metabolism of mannitol has been hypothesised as
occurring in a cycle involving the enzymes mannitol 2-dehydrogenase (Mdh1) and
mannitol 1-phosphate 5-dehydrogenase (Mpd1). A strain was created harbouring
disruption constructs for both of these genes. The double mutant was unable to
synthesise or catabolise mannitol, and was unable to sporulate. Addition of exogenous
mannitol completely restored in vitro sporulation, and partially restored in planta
sporulation. This demonstrated an essential role for mannitol in asexual sporulation.
This is the first demonstrated role for this compound.
A 13C NMR study of the wild type strain, the mdh1 and mpd1 single mutants,
and mpd1mdh1 double mutant was undertaken to investigate carbon utilisation and
cycling. Disruption of Mpd1 significantly altered the metabolite profile with the mpd1
mutants producing trehalose and glycerol in place of mannitol. Labelling patterns in
the double mutant showed that scrambling of label can be explained by the
iv
triosephosphate isomerase triangle and pentose phosphate pathway. This suggests the
contribution of mannitol to label scrambling has been overstated in previous studies.
The evidence did not support the metabolism of mannitol in S. nodorum as
occurring in a cycle, but rather as two separate pathways.
A GC-MS analysis of diseased and non-diseased tissue from infected leaves,
compared to non-infected and mock-inoculated leaves, could not detect any
metabolites associated with a systemic host reaction to pathogen attack.
v
ACKNOWLEDGEMENTS
When the opportunity arose to undertake PhD studies with Prof. Richard
Oliver and Dr. Peter Solomon, it required no lengthy consideration. Having completed
my Honours degree with these same supervisors, and being familiar with the ACNFP,
I knew that I was in good hands and that in addition to undertaking some seriously
good science, there would be no shortage of fun along the way. In this I have not been
disappointed and I am very grateful to my supervisors for being willing to take on a
rather more mature (in years) student than average. I suspect that the social and
cultural baggage I have accumulated along the way has not always been conducive to
the best scientific method and approach, and they have borne my idiosyncrasies well.
There has been a lot of laughter along the way and that is never a bad thing! My
candidature has been rather complicated by the arrival of a third child, the lengthy
hospitalisation my mother, and two root canals. In between these there have been
some exciting moments in the lab, not least of which was the creation of the elusive
double mutant mpd1mdh1 strain. I am grateful to Yair Shachar-Hill of Michigan State
University for introducing me to the world of NMR spectroscopy and to Rob
Trengove and the Murdoch Separation Science Laboratory for assistance with aspects
of the GC-MS part of the study. A big thank you is due to all members of the ACNFP
for their camaraderie and support. In particular our wonderful RAs, especially Kasia
Rybak, and my fellow stagonaut PhD students (some now post-docs) Kar-Chun Tan,
Rohan Lowe, Simon Ip Cho, James Hane, Joel Gummer and Christian Krill. I have
been very fortunate to have been funded by the Grains Research and Development
Corporation and their support is very sincerely appreciated.
vi
I would like to dedicate this PhD to my family who have offered ongoing love
and support. In particular my parents Ormonde and Mina, and my parents-in-law
Neville (deceased) and Daphne Harris, and my three great wonders – Clare, Oscar and
Fintan (who have been promised a camping trip once I submit – apologies for the
delay). Most importantly I dedicate this to Suzanne who knows me better than myself,
puts up with a lot and gives so much, and is the sun, moon, stars and music of my
life…agus do Íosa mhín, mo Thiarna, do chum ghlóire Dé agus d’onóir Éireann ‘s An
Astráil.
vii
ABBREVIATIONS
1-P
1-phosphate
6-P
6-phosphate
ACNFP
Australian Centre for Necrotrophic Fungal Pathogens
b
base(s)
BCA
bicinchoninic acid
BLAST
Basic Local Alignment Search Tool
bp
nucleotide base pair(s)
BSA
bovine serum albumin
cDNA
complementary DNA
CFE
cell-free extract
cm
centimetre(s)
cv.
cultivar
CzV8CS
Czapek Dox V8 juice complete supplement
DLA
detached leaf assay
DNA
deoxyribonucleic acid
dNTP
deoxyribonucleotide triphosphate(s)
dpi
days post inoculation/days post infection
EC
Enzyme Commission
EDTA
ethylenediaminetetra-acetic acid, disodium salt pH 8.0
EST
expressed sequence tag
f. sp.
forma(e) species
FT
Fourier transform
g
gram(s)
viii
g
gravity
GC-MS
Gas Chromatography-Mass Spectrometry
gDNA
genomic DNA
GFP
Green Fluorescent Protein
GPS™-M
Genome Priming System - Mutagenesis
h
hour(s)
HSD
honestly significant difference
HST
host-specific toxin
kb
kilobase(s)
kPa
kiloPascal(s)
kV
kilovolt(s)
L
litre(s)
μg
microgram(s)
μL
microlitre(s)
μM
microMolar
μm
micron(s)
M
Molar
MAP kinase
mitogen-activated protein kinase
Mb
Megabase(s)
Mdh1/Mdh1
mannitol 2-dehydrogenase (gene/protein)
mg
milligram(s)
min
minute(s)
mL
millilitre(s)
MM
minimal medium
MM-C
minimal medium minus carbon
ix
mM
milliMolar
mm
millimetre(s)
mol
mole(s)
MPa
Mega Pascal(s)
Mpd1/Mpd1
mannitol-1-phosphate 5-dehydrogenase (gene/protein)
NA
natural abundance
NAD+
nicotinamide adenine dinucleotide (oxidised)
NADH
nicotinamide adenine dinucleotide (reduced)
NADP+
nicotinamide adenine dinucleotide phosphate (oxidised)
NADPH
nicotinamide adenine dinucleotide phosphate (reduced)
ng
nanogram(s)
nm
nanometre(s)
NMR
nuclear magnetic resonance
PCA
Principal Components Analysis
PCR
polymerase chain reaction
PEG
polyethylene glycol
PEP
phosphoenolpyruvate
pers. comm.
personal communication
pH
potential of hydrogen
Pi
inorganic phosphate
pl.
plural
ppm
parts per million
QTL
quantitative trait locus/loci
qPCR
quantitative polymerase chain reaction
rcf
relative centrifugal force
x
RNA
ribonucleic acid
RNAase
ribonuclease
ROS
reactive oxygen species
rpm
revolutions per minute
SDBS
Spectral Database for Organic Compounds
SDS
sodium dodecylsulphate
SE
standard error
sec
second(s)
sing.
singular
SNB
stagonospora nodorum blotch
sp.
species (sing.)
spp.
species (pl.)
subsp.
subspecies
syn.
synonym
TCA
tricarboxylic acid
TMS
trimethylsilyl
Tween 20
polyoxyethylenesorbitan monolaurate
U
unit(s)
UV
ultraviolet
V
volt(s)
v/v
volume per volume
WT
wild type
w/v
weight per volume
xi
TABLE OF CONTENTS
CHAPTER 1: METABOLISM AND INFECTION ................................................. 1
1.1 THE PATHOSYSTEM CONCEPT...................................................................................................... 2
1.2 THE STAGONOSPORA NODORUM-WHEAT PATHOSYSTEM............................................................. 4
1.2.1 The Host (Triticum aestivum L.) .......................................................................................... 4
1.2.2 The Pathogen (Stagonospora nodorum) .............................................................................. 4
1.2.2.1 Discovery and nomenclature of the organism ................................................................................ 4
1.2.2.2 Taxonomic placement ..................................................................................................................... 7
1.2.2.3 Host range ...................................................................................................................................... 7
1.2.2.4 Economic importance ..................................................................................................................... 8
1.2.2.5 Nomenclature of the disease........................................................................................................... 8
1.3 THE INFECTION PROCESS ............................................................................................................. 9
1.3.1 Life Cycle of Stagonospora nodorum ................................................................................... 9
1.3.2 Modes of Host Inoculation ................................................................................................. 12
1.3.3 Disease Symptoms ............................................................................................................... 12
1.4 METHODS OF DISEASE CONTROL ............................................................................................... 13
1.4.1 Chemical Control ................................................................................................................ 13
1.4.2 Host Resistance/Tolerance.................................................................................................. 14
1.4.3 Cultural Practices ............................................................................................................... 15
1.4.4 Biological Antagonists ........................................................................................................ 16
1.4.5 Genetic Manipulation of S. nodorum ................................................................................. 16
1.5 METABOLISM AND INFECTION.................................................................................................... 18
1.5.1 Definition of Metabolism .................................................................................................... 18
1.5.2 The Impact of Disrupted Metabolism on Infection ............................................................ 18
1.5.2.1 Germination and penetration ....................................................................................................... 19
1.5.2.2 Proliferation ................................................................................................................................. 20
1.5.2.3 Sporulation ................................................................................................................................... 21
1.6 MANNITOL METABOLISM AND INFECTION ................................................................................ 22
1.6.1 Postulated Roles of Mannitol.............................................................................................. 22
1.6.2 Enzymatic Metabolism of Mannitol in Fungi .................................................................... 23
1.6.2.1 D-mannitol:NADP+ 2-oxidoreductase (EC 1.1.1.138) ................................................................. 30
1.6.2.2 D-mannitol:NAD+ 2-oxidoreductase (EC 1.1.1.67) ...................................................................... 30
1.6.2.3 D-mannitol-1-phosphate:NAD+ 5-oxidoreductase (EC 1.1.1.17) ................................................. 31
1.6.2.4 D-mannitol-1-phosphate phosphohydrolase (EC 3.1.3.22) .......................................................... 32
1.6.2.5 D-mannitol kinase (EC 2.7.1.57 (created 1972, deleted 1984)).................................................... 32
1.6.2.6 D-mannitol acetyl phosphate phosphotransferase (no EC number) ............................................. 33
1.6.2.7 D-mannitol phosphoenolpyruvate phosphotransferase (no EC number) ...................................... 33
1.6.2.8 Hexokinases ................................................................................................................................. 34
1.6.2.9 D-fructose-6-phosphate phosphatase (no EC number) ................................................................. 34
1.6.3 The Postulated Mannitol Cycle........................................................................................... 35
1.7 SUMMARY AND AIMS................................................................................................................... 39
CHAPTER 2 – GENERAL MATERIALS AND METHODS ............................... 40
2.1 FUNGAL AND BACTERIAL STRAINS............................................................................................. 41
2.2 WHEAT VARIETY ........................................................................................................................ 41
2.3 GENERAL MEDIA ........................................................................................................................ 42
2.4 GROWTH OF TRITICUM AESTIVUM CV. AMERY........................................................................... 42
2.5 GROWTH OF STAGONOSPORA NODORUM ..................................................................................... 46
2.5.1 Routine Maintenance and Culture ..................................................................................... 46
2.5.2 Harvesting of Pycnidiospores ............................................................................................. 46
2.6 GROWTH OF ESCHERICHIA COLI ................................................................................................. 47
2.7 NUCLEIC ACID EXTRACTION AND MANIPULATION ................................................................... 47
2.7.1 Homogenisation of Fungal Mycelium/Pycnidiospores ...................................................... 47
2.7.2 Genomic DNA Extraction from Lysed Fungal Mycelium/Pycnidiospores ....................... 48
2.7.3 Plasmid DNA Extraction .................................................................................................... 48
xii
2.7.4 Gel Electrophoresis of DNA ............................................................................................... 49
2.7.5 Determination of DNA Concentration ............................................................................... 50
2.7.6 Restriction Endonuclease Digestion of DNA ..................................................................... 50
2.7.7 Purification of Linearised Plasmid DNA ........................................................................... 51
2.7.8 DNA Amplification by Polymerase Chain Reaction .......................................................... 51
2.8 GAS CHROMATOGRAPHY – MASS SPECTROMETRY .................................................................. 52
2.8.1 Extraction of Polar Metabolites.......................................................................................... 52
2.8.2 Derivatisation of Polar Metabolite Extracts ....................................................................... 53
2.8.3 Gas Chromatography – Mass Spectrometry ....................................................................... 54
2.8.4 Data Normalisation ............................................................................................................. 55
2.9 SOFTWARE ................................................................................................................................... 55
2.10 STATISTICAL ANALYSIS ............................................................................................................ 56
CHAPTER 3 - CONSTRUCTION AND CHARACTERISATION OF A
STRAIN OF STAGONOSPORA NODORUM HARBOURING DISRUPTED
GENES FOR MANNITOL 2-DEHYDROGENASE (MDH1) AND MANNITOL
1-PHOSPHATE 5-DEHYDROGENASE (MPD1). ................................................. 57
3.1 INTRODUCTION...................................................................................................................... 58
3.1.1 Nomenclature, Class and Structure of D-Mannitol ........................................................... 58
3.1.2 Taxonomic Distribution ...................................................................................................... 58
3.1.3 Mannitol Metabolic Pathways in Stagonospora nodorum ................................................ 60
3.2 MATERIALS AND METHODS ............................................................................................... 65
3.2.1 Fungal Transformation ...................................................................................................... 65
3.2.1.1 Preparation of Protoplasts ........................................................................................................... 65
3.2.1.2 Transformation of Protoplasts ..................................................................................................... 66
3.2.1.3 Screening of Transformants ......................................................................................................... 68
3.2.1.4 Sub-Culturing of Transformant Colonies ..................................................................................... 69
3.2.2 Southern Hybridisation....................................................................................................... 69
3.2.2.1 PCR Amplification of DNA Probes .............................................................................................. 69
3.2.2.2 DIG-Labelling of DNA Probes ..................................................................................................... 70
3.2.2.3 Genomic DNA Digestion and Electrophoresis ............................................................................. 71
3.2.2.4 Southern Blot................................................................................................................................ 71
3.2.2.5 Hybridisation and Immunological Detection ............................................................................... 72
3.2.3 In vitro Growth Assays ........................................................................................................ 73
3.2.3.1 Growth on Solid Media ................................................................................................................ 73
3.2.3.2 Ability to Grow on Selected Carbon Sources ............................................................................... 74
3.2.3.3 Germination Assay ....................................................................................................................... 74
3.2.4 Enzyme Assays .................................................................................................................... 75
3.2.4.1 Preparation of Mycelium from Liquid Culture ............................................................................. 75
3.2.4.2 Determination of Protein Concentration ...................................................................................... 76
3.2.4.3 Measurement of Relative Enzyme Activity.................................................................................... 76
3.2.4.3.1 NADP+-dependent glucose 6-phosphate oxidation (glucose 6-phosphate dehydrogenase) . 77
3.2.4.3.2 NADPH-dependent fructose reduction (mannitol dehydrogenase) ...................................... 77
3.2.4.3.3 NADP+-dependent mannitol oxidation (mannitol dehydrogenase) ...................................... 78
3.2.4.3.4 NADH-dependent fructose reduction (NAD-mannitol dehydrogenase) ............................... 78
3.2.4.3.5 NAD+-dependent mannitol oxidation (NAD-mannitol dehydrogenase) .............................. 78
3.2.4.3.6 NAD+-dependent sorbitol oxidation (sorbitol dehydrogenase) ............................................ 79
3.2.4.3.7 NADH-dependent fructose 6-phosphate reduction (mannitol 1-phosphate dehydrogenase) 79
3.2.4.3.8 NAD+-dependent mannitol 1-phosphate oxidation (mannitol 1-phosphate dehydrogenase) 80
3.2.4.4 Calculation of Specific Enzyme Activity ....................................................................................... 80
3.2.5 Stress Tolerance Assays ...................................................................................................... 80
3.2.5.1 Osmotic Stress Assay .................................................................................................................... 80
3.2.5.2 Oxidative Stress Assay ................................................................................................................. 81
3.2.6 Pathogenicity Assays ........................................................................................................... 81
3.2.6.1 Detached Leaf Assay .................................................................................................................... 81
3.2.6.2 Whole Plant Spray........................................................................................................................ 82
3.2.6.3 Latent Period Assay ..................................................................................................................... 84
3.2.6.4 Microscopic Examination of Host Penetration ............................................................................ 84
3.2.7 Mannitol Supplementation Assays ..................................................................................... 86
3.2.7.1 In vitro Response to Mannitol Supplementation ........................................................................... 86
xiii
3.2.7.1.1 In vitro sporulation response to altered mannitol concentration ......................................... 86
3.2.7.1.2 Assay of mannitol content of spores ..................................................................................... 87
3.2.7.2 In planta Response to Mannitol Supplementation......................................................................... 87
3.3 RESULTS ................................................................................................................................... 88
3.3.1 Isolation of the mpd1mdh1 Double Mutant Strain ............................................................ 88
3.3.1.1 Transformation of Protoplasts ..................................................................................................... 88
3.3.1.2 PCR Screening ............................................................................................................................. 88
3.3.1.3 Southern Hybridisation ................................................................................................................ 91
3.3.2 In vitro Phenotype ............................................................................................................... 91
3.3.2.1 Minimal Media Agar .................................................................................................................... 95
3.3.2.2 CZV8CS Agar ............................................................................................................................... 97
3.3.2.3 V8-PDA ........................................................................................................................................ 98
3.3.2.4 Mean Daily Growth Rates on Solid Medium ................................................................................ 99
3.3.2.5 Ability to Grow on Selected Carbon Sources ............................................................................. 101
3.3.2.6 Germination Assay ..................................................................................................................... 101
3.3.3 Enzyme Assays. ................................................................................................................. 104
3.3.4 Stress Tolerance Assays .................................................................................................... 104
3.3.4.1 Osmotic Stress Assay .................................................................................................................. 104
3.3.5 Pathogenicity Assays ......................................................................................................... 107
3.3.5.1 Detached Leaf Assay .................................................................................................................. 107
3.3.5.2 Whole Plant Spray...................................................................................................................... 111
3.3.5.3 Latent Period Assay ................................................................................................................... 111
3.3.5.4 Microscopic Examination of Host Penetration .......................................................................... 113
3.3.6 Mannitol Supplementation Assays ................................................................................... 113
3.3.6.1 In vitro Response to Mannitol Supplementation ......................................................................... 113
3.3.6.1.1 In vitro sporulation response to altered mannitol concentration ....................................... 113
3.3.6.1.2 Assay of mannitol content of spores ................................................................................... 117
3.3.6.2 In Planta Response to Mannitol Supplementation ...................................................................... 117
3.4 DISCUSSION ........................................................................................................................... 120
3.4.1 Isolation of the mpd1mdh1 Double Mutant Strain .......................................................... 120
3.4.2 Enzyme Assays .................................................................................................................. 120
3.4.3 Mannitol Synthesis can Occur by Two Pathways ............................................................ 121
3.4.4 Mannitol Catabolism is Facilitated Primarily via Mannitol 1-Phosphate ...................... 122
3.4.5 Mannitol is Required for Asexual Sporulation ................................................................ 123
3.5 CONCLUSION ........................................................................................................................ 125
3.5.1 Mannitol is Required for Pathogenicity ........................................................................... 125
3.5.2 Enzymatic Cycling of Mannitol is Physiologically Unimportant .................................... 125
CHAPTER 4: METABOLOMICS ANALYSIS OF HEALTHY AND DISEASED
LEAVES.................................................................................................................... 128
4.1 INTRODUCTION.................................................................................................................... 129
4.1.1 The Metabolome and Antimicrobial Metabolites ............................................................. 129
4.1.2 Overview of Technique ..................................................................................................... 130
4.1.2.1 Gas Chromatography-Mass Spectrometry ................................................................................. 130
4.1.2.2 Principal Components Analysis ................................................................................................. 131
4.1.3 Aims of the Study .............................................................................................................. 132
4.2 MATERIALS AND METHODS ............................................................................................. 133
4.2.1 Sample Collection ............................................................................................................. 133
4.2.2 Sample Preparation for GC-MS ....................................................................................... 134
4.2.3 Data Analysis..................................................................................................................... 134
4.3 RESULTS ................................................................................................................................. 135
4.3.1 GC-MS Peaks .................................................................................................................... 135
4.3.2 Principal Components Analysis ........................................................................................ 135
4.3.3 Statistical Analysis of Metabolites Identified by PCA ...................................................... 144
4.3.3.1 Metabolites Present Only in Diseased Samples ......................................................................... 144
4.3.3.2 Metabolites Increasing with Time of Infection in Diseased Samples.......................................... 146
4.3.3.3 Metabolites Significantly Higher in Healthy Tissue than Diseased Tissue................................. 146
4.3.3.4 Metabolites Significantly Lower in Late Stage Diseased Tissue ................................................ 146
4.4 DISCUSSION ........................................................................................................................... 151
4.4.1 Compounds Associated with Infected Tissue Only .......................................................... 151
4.4.1.1 Mannitol ..................................................................................................................................... 151
xiv
4.4.1.2 Trehalose .................................................................................................................................... 152
4.4.1.3 L-Arabitol................................................................................................................................... 153
4.4.2 Plant Specific Compounds ................................................................................................ 154
4.4.3 Miscellaneous Metabolites ................................................................................................ 155
4.4.4 No Evidence Found For an Induced Defence Response in the S. nodorum-Wheat
Pathosystem ................................................................................................................................ 157
4.5 CONCLUSION ........................................................................................................................ 158
CHAPTER 5: 13C-NMR INVESTIGATION OF MANNITOL METABOLISM
IN STAGONOSPORA NODORUM......................................................................... 159
5.1 INTRODUCTION.................................................................................................................... 160
5.1.1 Overview of Technique ..................................................................................................... 160
5.1.2 Advantages and Disadvantages of NMR .......................................................................... 161
5.1.3 13C-NMR Studies in Filamentous Fungi .......................................................................... 163
5.1.4 Aims of the Study .............................................................................................................. 164
5.2 MATERIALS AND METHODS ............................................................................................. 165
5.2.1 Preparation of Standards .................................................................................................. 165
5.2.2 Flask Culture of Fungal Strains ...................................................................................... 165
5.2.2.1 Natural Abundance Cultures ...................................................................................................... 165
5.2.2.2 [1-13C]-Glucose-Labelled Cultures ............................................................................................ 166
5.2.2.3 [1-13C]-Mannitol-Labelled Cultures .......................................................................................... 166
5.2.2.3.1 Assay of mannitol uptake ................................................................................................... 167
5.2.2.3.2 Preparation of [1-13C]-mannitol-labelled cultures ............................................................ 168
5.2.2.4 [1-13C]-Glucose Feed-Chase Cultures ....................................................................................... 168
5.2.3 Methanol/Water Extraction of Polar Metabolites ............................................................ 169
5.2.4 NMR Tube Preparation .................................................................................................... 169
5.2.5 Sample Preparation for NMR Analysis ............................................................................ 170
5.2.6 NMR Spectra Acquisition ................................................................................................. 170
5.2.7 NMR Spectra Analysis ...................................................................................................... 171
5.2.7.1 Software ..................................................................................................................................... 171
5.2.7.2 Compound Identity and Label Quantification ............................................................................ 171
5.2.7.2.1 Internal referencing of chemical shifts ............................................................................... 171
5.2.7.2.2 MaxVar(RCS)..................................................................................................................... 172
5.2.7.2.3 Max(RPI) ........................................................................................................................... 173
5.2.7.2.4 Missing peaks ..................................................................................................................... 174
5.2.7.2.5 Comparison of relative abundances between spectra ........................................................ 175
5.2.7.2.6 Quantification of 13C-labelling .......................................................................................... 176
5.3 RESULTS ................................................................................................................................. 176
5.3.1 Standards ........................................................................................................................... 176
5.3.2 Identified Compounds ....................................................................................................... 182
5.3.2.1 13C Natural Abundance Spectra ................................................................................................. 182
5.3.2.1.1 Replicates Inoculated from CZV8CS Agar Cultures .......................................................... 182
5.3.2.1.2 Replicates Inoculated from Minimal Medium Agar Cultures............................................. 186
5.3.2.2 [1-13C]-Glucose-Labelled Spectra ............................................................................................. 186
5.3.2.2.1 [1-13C]-Glucose ................................................................................................................. 189
5.3.2.2.2 Mannitol ............................................................................................................................. 189
5.3.2.2.3 Trehalose ........................................................................................................................... 189
5.3.2.2.4 Glycerol ............................................................................................................................. 190
5.3.2.2.5 Alanine ............................................................................................................................... 190
5.3.2.2.6 Glutamine........................................................................................................................... 191
5.3.2.2.7 Glutamate........................................................................................................................... 191
5.3.2.2.8 Arabitol .............................................................................................................................. 191
5.3.2.2.9 Arginine ............................................................................................................................. 192
5.3.2.3 [1-13C]-Mannitol-Labelled Spectra ............................................................................................ 192
5.3.2.3.1 Assay of mannitol uptake ................................................................................................... 192
5.3.2.3.2 Gross features of spectra ................................................................................................... 194
5.3.2.3.3 Mannitol ............................................................................................................................. 194
5.3.2.3.5 Glucose .............................................................................................................................. 196
5.3.2.3.6 Glycerol ............................................................................................................................. 196
5.3.2.3.7 Arabitol .............................................................................................................................. 196
5.3.2.3.8 Amino acids........................................................................................................................ 197
5.3.2.4 [1-13C]-Glucose Feed-Chase Spectra ........................................................................................ 198
xv
5.3.2.4.1 Carbohydrates ................................................................................................................... 198
5.3.2.4.2 Amino acids........................................................................................................................ 202
5.3.3 Miscellaneous Peaks ......................................................................................................... 203
5.4 DISCUSSION ........................................................................................................................... 207
5.4.1 Disruption of Mpd1 Alters the Metabolite Profile............................................................ 207
5.4.1.1 Mannitol ..................................................................................................................................... 208
5.4.1.2 Trehalose and Glycerol .............................................................................................................. 210
5.4.1.3 Glucose ...................................................................................................................................... 213
5.4.1.4 Arabitol and Amino Acids .......................................................................................................... 215
5.4.2 No Third Pathway of Mannitol Metabolism Detected in S. nodorum ............................. 216
5.4.3 Scrambling of Label is not Proof of a Mannitol Cycle .................................................... 217
5.4.3.1 The Mdh1 Pathway does not Contribute to Label Scrambling ................................................... 218
5.4.3.2 The Aldose/Triosephosphate Isomerase Triangle....................................................................... 219
5.4.3.3 The Pentose Phosphate Pathway (Forward Flux) ...................................................................... 221
5.4.3.4 The Pentose Phosphate Pathway (Reverse Flux) ....................................................................... 224
5.4.4 Mannitol Metabolism does not Contribute to NADPH Regeneration ............................. 226
5.4.5 Experimental Considerations ........................................................................................... 227
5.4.5.1 Co-located Peaks in Biological Samples Obscure Labelling ..................................................... 227
5.4.5.2 Low Sample Weights Affect Detection of Low Abundance Metabolites...................................... 228
5.4.5.3 Spectrometer Artefacts/Variation ............................................................................................... 228
5.4.5.4 Quantification of 13C Labelling .................................................................................................. 229
5.4.5.5 Internal Referencing of Chemical Shifts ..................................................................................... 229
5.4.5.6 Limitations of Published Chemical Shifts ................................................................................... 231
5.4.5.7 Necessity for a Local Library of Compound Standards.............................................................. 232
5.4.5.8 Assumption of Labelling of Mannitol on One Terminal Carbon ................................................ 233
5.5 CONCLUSION ........................................................................................................................ 233
CHAPTER 6: GENERAL CONCLUSIONS ........................................................ 237
6.1 OVERVIEW ................................................................................................................................. 238
6.2 KEY FINDINGS ........................................................................................................................... 238
6.3 FUTURE DIRECTIONS ................................................................................................................ 240
CHAPTER 7: REFERENCES ................................................................................ 242
CHAPTER 8: APPENDICES ................................................................................. 297
xvi
LIST OF TABLES
Table
1.1
Caption
Page
Area set aside for production of all crops, for wheat alone, and wheat
5
as a percentage of the total area cropped in Australia from 2002/03 to
2005/06.
1.2
Area, Production and Gross Value of cereal crops grown in Australia
5
from 2002/03 to 2005/06.
1.3
Distribution of genes reported as being involved in the metabolism of
24
mannitol in fungal species.
2.1
Media used in this study.
43
3.1
Relative growth of Stagonospora nodorum strains SN15, mdh1-71,
102
mpd1-1 and mpd1mdh1-107 on selected media in liquid culture
3.2
Specific enzyme activities for selected Stagonospora nodorum strains.
105
All activities are shown as U/mg protein unless otherwise indicated.
4.1
Library of retention times (RT) and identities for metabolites detected
136
by GC-MS from healthy and diseased tissue of wheat leaves infected
with Stagonospora nodorum and harvested at 0.5, 1, 3, 5 and 8 days
post infection. Metabolites from negative controls including mockinoculated and uninoculated leaves are included.
5.1
13
C natural abundance NMR spectra
177
C-NMR chemical shifts (ppm) for the peaks of D-mannitol from
179
Standard compounds for which
were acquired.
5.2
13
Standard Compound compilations and from reported experimental
observations. The calculated differences in the relative positions of the
C2,5 and C3,4 – and C2,5 and C1,6, and the net difference in
published chemical shifts for each peak, are shown.
5.3
Peak clusters from
13
C-NMR spectra of strains of Stagonospora
204
nodorum for peaks comprising >1% of total intensity. The range for
each cluster, the number of spectra comprising each cluster, the strains
and treatments (including the number of replicates), and the best
match for the cluster from the ACNFP Compound Standard Library
are shown.
Continued on the following page
xvii
LIST OF TABLES (contd)
Table
5.4
Caption
Page
Distribution of unidentified peaks from 13C-NMR spectra of strains of
205
Stagonospora nodorum into clusters.
8.1
Plant species other than Triticum aestivum L. subsp. aestivum
298
(excluding hybrids)
8.2
Names (in English) which have been used to describe the disease
304
caused by Stagonospora nodorum on wheat.
8.3
ACNFP library of
13
C chemical shifts, carbon assignments, peak
306
intensities, and calculated ideal natural abundance relative peak
intensities (RPI) for compound standards.
xviii
LIST OF FIGURES
Figure
1.1
Caption
Page
Distribution and yield of wheat production in Australia 2000-01.
6
Source: ABS (2006).
1.2
The life cycle of Phaeosphaeria nodorum [anamorph Stagonospora
10
nodorum]. Source: Eyal et al. (1987).
1.3
(A) The mannitol enzymatic cycle as proposed by Hult and
36
Gatenbeck (1978). Figure as given in Hult et al. (1980).
(B) The modified mannitol cycle proposed by Jennings and Burke
(1990).
3.1
The structure of D-mannitol (Fischer projection)
59
3.2
Diagram outlining the construction of the knockout vector pGPSH-
62
Mpd8.
3.3
Diagram outlining the construction of the knockout vector pGPSP-
63
Mdh1.
3.4
Score chart for assigning disease scores to wheat cv. Amery
83
seedlings infected with strains of Stagonospora nodorum.
3.5
Criteria for assigning developmental stages in Stagonospora
85
nodorum pycnidia on leaves of wheat cv. Amery.
3.6
Duplex PCR amplification of gDNA from SN15 and mutant strains
89
transformed with pGPSP-Mdh1 or having this construct as their
background. PCR amplification was conducted using actinF/R
primers (~300 bp) and mdhkoF/R primers (~461 bp), with an
annealing temperature of 57 °C.
3.7
Duplex PCR amplification of gDNA from SN15 and mutant strains
90
transformed with pGPSH-Mpd8. PCR amplification was conducted
using actinF/R primers (~300 bp) and mpdkoF/R primers (~500 bp).
3.8
PCR amplification of gDNA from SN15 for use as a probe for
Southern analysis.
92
PCR amplification was conducted using
mdhSOUTHF/R primers (~393 bp) or mpdSOUTHF/R (~311 bp).
Continued on the following page
xix
LIST OF FIGURES (contd)
Figure
3.9
Caption
Page
A: Southern analysis of ApaI-digested gDNA transformed with the
93
pGPSH-Mpd8 disruption construct, using probes homologous to
Mpd1.
B: Southern analysis of HindIII-digested gDNA transformed with the
pGPSP-Mdh1 disruption construct, using probes homologous to
Mdh1.
3.10
Phenotypic characterisation of strains of Stagonospora nodorum
94
grown on three different media.
3.11
Mean daily growth rate (cm/day) (±SE) of strains of Stagonospora
100
nodorum on solid media.
3.12
Mean percentage of germinated spores (±SE) for selected strains of
103
Stagonospora nodorum at 24 hpi on 1% agarose.
3.13
Assays of the ability of strains of Stagonospora nodorum to grow
106
under conditions of osmotic stress (A) and oxidative stress (B).
3.14
Mean lesion size (±SE) on detached wheat Amery leaves inoculated
108
with SN15 (♦), mdh1-71 (■), mpd1-1 (▲), mpd1mdh1-102 (●),
mpd1mdh1-107 (О) Mpd1mdh1-101 (X), Tween control (*) and
uninoculated control ().
3.15
Detached leaf assay at 12 days post infection with strains of
109
Stagonospora nodorum as noted above.
3.16
Lesion formation on a detached leaf assay at 12 days post-inoculation
110
with selected strains of Stagonospora nodorum on wheat as noted
above. Note the absence of pycnidia in the mpd1 mutants.
3.17
Mean disease scores (±SE) for wild type and selected mutant strains
112
of Stagonospora nodorum from a whole plant spray pathogenicity
assay.
3.18
Trypan blue-stained lesions from detached leaves infected with
114
Stagonospora nodorum strains SN15 and mpd1mdh1-107. Arrows
indicate penetration attempts.
Continued on the following page
xx
LIST OF FIGURES (contd)
Figure
3.19
Caption
Page
A: The effect of mannitol supplementation upon sporulation of the
115
strains SN15, mdh1-71, mpd1mdh1-107 and Mpd1mdh1-101. Mean
spores/mL (±SE) for strains grown on minimal media agar
supplemented with 0, 1, 3, 10 and 30 mM mannitol are shown. N=3.
B: Pycnidia production by mpd1mdh1-107 in response to changes in
mannitol concentration in supplemented minimal media agar.
3.20
Comparison of mean spores/mL (±SE) for strains of Stagonospora
116
nodorum as shown. Blue columns result from growth on minimal
media agar. Purple columns result from growth on minimal media
agar supplemented with 3 mM mannitol. The inoculum for the
double mutant strain mpd1mdh1-107 came from minimal medium
agar plates on which the strain had been serially sub-cultured for 1, 2
and 3 generations as indicated by the suffix.
3.21
GC-MS chromatograms demonstrating the amount of mannitol
118
present in spores of SN15, mpd1-1 and mpd1mdh1-107 harvested
from minimal medium agar plates.
3.22
Chemical complementation of the in planta sporulation defect of the
119
Stagonospora nodorum double mutant strain mpd1mdh1-107.
Lesions were inoculated with 5 μL 3 mM mannitol on a daily basis
from 3 days post infection.
3.23
The two pathways for mannitol metabolism in Stagonospora
129
nodorum showing the enzymes involved in each step including a
putative mannitol phosphorylation step catalysed by unknown
enzyme(s).
4.1
Principal components analysis (PCA) score plot (A) and loading plot
141
(B) for PC1 versus PC2 from a PCA of polar metabolites processed
by GC-MS.
Continued on the following page
xxi
LIST OF FIGURES (contd)
Figure
4.2
Caption
Page
The top 20 variables (metabolites) contributing to the variation
143
accounted for by PC1 (A) and PC2 (B) in a PCA of healthy and
Stagonospora nodorum-infected wheat leaf tissue.
4.3
Mean normalised abundance (±SE) for metabolites present only in
145
diseased tissue.
4.4
Mean normalised abundance (±SE) for metabolites significantly
147
higher in later stage infected tissue.
4.5
Mean normalised abundance (±SE) for metabolites significantly
148
higher in healthy tissue.
4.6
Mean normalised abundance (±SE) for metabolites significantly
149
lower in late stage diseased tissue.
4.7
Mean normalised abundance (±SE) for metabolites significantly
150
lower in late stage diseased tissue.
5.1
13
C NMR spectra for D-mannitol illustrating source-dependent
180
differences in relative height of peaks.
5.2
13
C-NMR spectra showing co-location of the chemical shifts of the
181
C1 resonance peak of L-arabitol (red) and a spinning sideband of the
C1,6 resonance peak of [1-13C]-D-mannitol (black).
5.3
Natural abundance 13C NMR spectrum of SN15 showing the regions
183
from 75-100 ppm, 60-76 ppm and 15-60 ppm.
5.4
Natural abundance
13
C NMR spectrum of mpd1mdh1-107 showing
184
the regions from 75-100 ppm, 60-76 ppm and 15-60 ppm.
5.5
Mean relative abundance (±SE) of (A) major (>10%), and (B) minor
185
(<10%) soluble metabolites in extracts of strains of Stagonospora
nodorum cultured for 3 days in flasks with 40 mM glucose, as
determined by 13C NMR analysis.
5.6
Natural abundance 13C NMR spectrum of mpd1mdh1-107 (inoculum
187
sourced from minimal medium agar plates) showing the regions from
(A) 75-100 ppm, (B) 60-76 ppm and (C) 15-60 ppm
Continued on the following page
xxii
LIST OF FIGURES (contd)
Figure
5.7
Caption
13
Page
C-NMR spectra of SN15 (A) and mpd1mdh1-107 (B) showing the
188
region from 15-100 ppm for cultures grown for 3 days on [1-13C]glucose.
5.8
A: Standard curve relating concentration of mannitol to net change in
193
absorbance at 340 nm due to the mannitol oxidation activity of
mannitol dehydrogenase in a cell-free extract of Stagonospora
nodorum strain SN15. N≥ 3.
B: The concentration of mannitol in various samples and controls as
determined by observed mannitol oxidation activity in conjunction
with the mannitol standard curve above.
5.9
13
C-NMR spectra of Stagonospora nodorum strains SN15, mdh1-71,
195
mpd1-1 and mpd1mdh1-107 grown for two days on 40 mM glucose
followed by 24 h on 20 mM [1-13C]-mannitol. Each spectrum is
representative of three independent experiments.
5.10
13
C-NMR spectra of SN15 cultures from a feed-chase experiment for
199
the range 15-100 ppm. Each spectrum is representative of at least
three independent experiments.
5.11
Changes in mean (±SE) fold labelling above
13
C natural abundance
200
for selected compounds over the course of a feed-chase experiment.
5.12
Aldolase/triosephosphate isomerase triangle mechanism for 13C label
220
scrambling from [1-13C]-glucose to [1-13C]/[6-13C] trehalose.
5.13
Pentose phosphate pathway mechanism for
13
13
13
C label scrambling
223
13
from [1- C]-mannitol to [1- C]/[6- C] trehalose.
5.14
Pentose phosphate pathway mechanism for
13
C label scrambling
225
from [1-13C]/[6-13C]-glucose 6-phosphate to [1-13C]/[5-13C] Larabitol.
5.15
Summary of the pathways of primary metabolism demonstrated to be
234
active in Stagonospora nodorum based on the detection of metabolic
intermediates in 13C-NMR spectra.
xxiii
PAPERS PUBLISHED FROM THIS STUDY
Solomon, P.S., Waters, O.D.C. and Oliver, R.P. (2007) Decoding the mannitol
enigma in filamentous fungi. Trends in Microbiology 15 (6): 257-262.
Solomon, P.S., *Waters, O.D., Jorgens, C.I., Lowe, R.G., Rechberger, J., Trengove,
R.D. and Oliver, R.P. (2006) Mannitol is required for asexual sporulation in
the wheat pathogen Stagonospora nodorum (glume blotch). Biochemical
Journal 399 (2): 231-9.
Solomon, P.S., Lowe, R.G.T., Tan, K.-C., Waters, O.D.C. and Oliver, R.P. (2006)
Stagonospora nodorum: cause of stagonospora nodorum blotch of wheat.
Molecular Plant Pathology 7 (3): 147-56.
*As equal first author. My contribution to this paper included the creation and
characterisation of the mdh1 mutant and the mpd1mdh1 double mutant and the
discovery that the double mutant was unable to undergo asexual sporulation without
the addition of exogenous mannitol.
A paper reporting the results of the
13
C-NMR study conducted here is in
preparation.
xxiv
CHAPTER 1: METABOLISM AND
INFECTION
1.1 The Pathosystem Concept
A pathosystem is defined by the phenomenon of parasitism i.e. where one
organism, the parasite or pathogen, derives some or all of its energy requirements
from a second organism, a living host (Robinson, 1976). Every living organism is
potentially a host, but can only actually be defined as such when another organism
becomes parasitic upon it. A pathosystem can therefore be defined as the biological
relationship which exists between a single pathogen and a single host.
In the case of plant pathosystems, the pathogens may be fungi, mycoplasmas,
bacteria, nematodes, oomycetes, viruses, or viroids. Occasionally it may be another
plant (March and Watson, 2007), but the definition does not extend to herbivory or to
the more mutualistic relationships between plants and mycorrhizal fungi, or insect and
mammalian pollination vectors.
The genomes of the participating organisms are the result of ongoing selection
pressures. They will continue to be shaped by such pressures, although the nature and
intensity of the pressures may alter over time. One particular selection pressure which
is applicable to a pathosystem is the asymmetric “evolutionary arms race” whereby
the host species are under selection pressure to favour the progeny of those
individuals with better mechanisms of resistance to the pathogen, while the pathogen
species will be under selection pressure to favour the progeny of those individuals
with better mechanisms of overcoming host resistance (Dawkins and Krebs, 1979).
These selection pressures can also be influenced by the nature of the host-pathogen
2
relationship which has been traditionally categorised as biotrophic or necrotrophic
(reviewed in Oliver and Ipcho, 2004).
Biotrophs are typically obligate pathogens with a narrow host range, causing
little damage to their host and feeding off living cells via haustoria, engaging in
classical gene-for-gene interactions, and with the hypersensitive response being a
feature of incompatible interactions (Both and Spanu, 2004). Necrotrophs are
characterised by being non-obligate with a broad host range, production of cell-wall
degrading enzymes and toxins, feeding off dead or dying host cells, and with host
resistance being polygenic (Oliver and Ipcho, 2004). Secreted toxins include hostspecific toxins (HSTs) which are important determinants of host range and may be
proteinaceous or low molecular weight compounds (Scheffer, 1983; Sarpeleh et al.,
2007). These include ToxA, initially characterised in the Pyrenophora tritici-repentiswheat pathosystem (Tuori et al., 1995; Ciuffetti et al., 1997), and subsequently shown
to have most likely originated in Stagonospora nodorum, providing the most
convincing evidence to date for eukaryotic interspecific virulence gene transfer
(Friesen et al., 2006). A special sub-category of necrotrophs is the hemibiotrophs,
which commence with an asymptomatic infection and after a latent period switch to a
host-cell destructive necrotrophic mode (Oliver and Ipcho, 2004).
The pathosystem which was the focus of this study is that occurring between
the host plant wheat (Triticum aestivum subsp. aestivum) and the necrotrophic fungal
pathogen Stagonospora nodorum.
3
1.2 The Stagonospora nodorum-Wheat Pathosystem
1.2.1 The Host (Triticum aestivum L.)
Wheat (Triticum aestivum L.) is one of the major crops produced worldwide in
terms of the amount of arable land reserved for its production, the volume of
production, and the value of the crop, with a forecast record production of 658 million
tonnes predicted for 2008/09 (FAO, 2008). In Australia, from 2003/04 to 2005/06
over 50% of all land farmed for crops was accounted for by wheat production (Table
1.1). Wheat accounted for over 60% of total cereal production (Table 1.2), and in
2006/07 wheat had a gross value of AUD 5.1 billion (ABS, 2007; ABS, 2008). The
cultivation of wheat in Australia occurs in a wide sub-coastal band known as the
wheat-belt, which extends around most of the southern half of the continent (Figure
1.1). The two main types of wheat cultivated are bread wheat (Triticum aestivum
subsp. aestivum) and durum or macaroni wheat (Triticum turgidum subsp. durum),
with the latter comprising 3% of the total crop (ABS, 2006).
1.2.2 The Pathogen (Stagonospora nodorum)
1.2.2.1 Discovery and nomenclature of the organism
Stagonospora nodorum (Berk.) E. Castell. and Germano (1977) [teleomorph:
Phaeosphaeria (syn. Leptosphaeria) nodorum (E. Müll.) Hedj. (1968)] was first noted
on wheat by Berkeley in 1845 (Weber, 1922). The anamorph has a number of
synonyms including Depazea nodorum Berk., Hendersonia nodorum (Berk.) Petr.,
4
Table 1.1: Area set aside for production of all crops, for wheat alone, and
wheat as a percentage of the total area cropped in Australia from 2002/03 to
2005/06 (Australian Bureau of Statistics, 2007, 2008).
Year
02/03
03/04
04/05
05/06
Total Area Under
Crops (‘000 ha)
23,575
26,080
26,742
24,255
Total Area of Wheat
(‘000 ha)
11,170
13,067
13,399
12,703
Wheat as a Percentage of
Total Area Cropped
47.4%
50.1%
50.1%
52.4%
Table 1.2: Area, Production and Gross Value of cereal crops grown in Australia from 2002/03 to 2005/06 (Australian Bureau of Statistics, 2007, 2008).
Area (‘000 ha)
Production (‘000 t)
Gross Value ($m)
2002/03 2003/04 2004/05 2005/06 2002/03 2003/04 2004/05 2005/06 2002/03 2003/04 2004/05 2005/06
Barley
3,864
4,477
4,646
4,481
3,865
10,382
7,740
9,641
984
1,750
1,233
na
Grain Sorghum
667
734
755
792
1,465
2,009
2,011
1,999
300
319
270
na
*
*
Maize
50
70
72
69
310
395
420
370
72
88
81
na
Oats
911
1,089
894
945
957
2,018
1,283
1,723
210
279
172
na
Rice
46
66
51
100
438
553
339
982
153
180
101
na
Wheat
11,170
13,067
13,399
12,703 10,132
26,132
21,905
25,704
2,692
5,636
4,317
5.1 bill.
Lupins
1,025
851
845
853
726
1,180
937
1,357
212
278
193
na
Total
17,733
20,354
20,662
19,943 17,893
42,274
34,635
41,776
4,623
8,442
6,367
Wheat (% of Total)
62.99
64.20
64.85
63.70
56.63
61.82
63.25
61.53
58.23
66.76
67.80
*
Estimate has a relative standard error of 10% to less than 25% and should be treated with caution.
na =not available
Figure 1.1: Distribution and yield of wheat production in Australia 2000-01.
Source: ABS (2006).
6
Macrophoma hennebergii (Kühn) Berl. & Vogl., Phoma hennebergii (Kühn) Lopr.,
Septoria glumarum Pass., Septoria nodorum (Berk.) Berk. apud Berk. & Br., and
Stagonospora hennebergii (Kühn) Petr. & Syd (Sutton and Waterston, 1966). This
pathogen was recently reviewed by Solomon et al. (2006c).
1.2.2.2 Taxonomic placement
The taxonomic placement of S. nodorum is as follows: kingdom Fungi;
phylum Ascomycota; subphylum Euascomycota; class Dothideomycetes; order
Pleosporales; family Phaeosphaeriaceae; genus Phaeosphaeria; species nodorum. Its
genome was recently sequenced and the current assembly consists of a nuclear
genome of 37,164,227 bp and a mitochondrial genome of 49,761 bp, with a minimum
of 10,762 nuclear genes (Hane et al., 2007). Electrophoretic karyotyping indicated
that the nuclear genome is comprised of 14-19 chromosomes (Cooley and Caten,
1991).
1.2.2.3 Host range
In common with necrotrophic pathogens, S. nodorum has a broad host range
despite a recent claim that it was pathogenic solely on wheat (Prell and Day, 2001). It
is a pathogen of bread wheat (Triticum aestivum L. subsp. aestivum) and related
cereals and wild grasses, with infections reported from over 70 species and subspecies
in 20 genera (Table 8.1). The fungus was isolated from a further 11
species/subspecies, including an additional 4 genera, although it was not clear from
the reports whether growth was pathogenic or epiphytic (Table 8.1).
7
1.2.2.4 Economic importance
The importance of S. nodorum has varied historically and geographically, and
since the 1970s it has been overtaken in Europe as the major fungal pathogen of
wheat by Septoria tritici (Bearchell et al., 2005). It is currently considered to be one
of the major diseases of wheat in North America (Singh et al., 2007) and Australia
(Solomon et al., 2006c), and is still regarded as being of worldwide significance
(Kluge et al., 2006). Crop yields can be significantly reduced by epidemic outbreaks
with reported heavy losses ranging from 15% in South Africa (Le Roux, 1984) to
46% in Poland (Pielka, 1957). Australia-wide estimated yield losses have been 1831% (Bhathal et al., 2003), with higher localised losses of 50% (Loughman et al.,
2001) to 70% in reported in Western Australia (Brown and Rosielle, 1980).
1.2.2.5 Nomenclature of the disease
The disease was initially referred to as “septoria” from the then assigned genus
of the causal organism (Grove, 1916; Cromwell, 1920; Sutton, 1920). It was
subsequently named glume blotch, since the observation of its ability to cause disease
symptoms in the glume was used as a major means of distinguishing it from another
common wheat pathogen, Septoria tritici (Rosen, 1921; Weber, 1922a). However, due
to the fact that: (i) this symptom is not always observed; (ii) the taxonomic placement
of the organism has undergone several reviews and subsequent name changes; and
(iii) some studies have focussed on different components of the disease; there are over
8
30 names which have been used to describe the disease, with nearly half of these
having been employed in publications in the last five years (Table 8.2).
A potential for confusion has been created by terms which describe a disease
“complex” caused by two or more independent pathogens (one of which is S.
nodorum), and by the usage of terms such “Stagonospora leaf blotch disease” to
describe the diseases caused by various Phaeospharia ssp. (including P. nodorum) in
a variety of cereals (Ueng et al., 2003; Wang et al., 2007). Since the late 1990s there
has been a growing acceptance of the term stagonospora nodorum blotch (SNB), and
it is this term which will be used in this study.
1.3 The Infection Process
1.3.1 Life Cycle of Stagonospora nodorum
The life cycle of Stagonospora nodorum is depicted in Figure 1.2. Mycelium
arising from a germinating spore which has landed on the plant surface or from a
seed-borne colony, can penetrate the plant by three methods. Firstly it can exploit
natural openings such as stomata or non-natural openings caused by physical damage
to the host. Secondly the mycelium can differentiate penetration structures called
hyphopodia which can directly penetrate the cell wall. Thirdly, the hyphal tip is also
able to directly penetrate the surface of the leaf by breaching the periclinal or
anticlinal epidermal cell wall (Solomon et al., 2006f). An apparent host defence
response to direct penetration attempts is the deposition of callose at the penetration
site, which has been suggested as the target of a β1,3-glucanase produced by the
9
[sic]
Figure 1.2: The life cycle of Phaeosphaeria nodorum [anamorph Stagonospora
nodorum]. Source: Eyal et al. (1987). N.B. “disposal” should read “dispersal”.
10
pathogen (Lehtinen, 1993). The reported rate of success of direct penetration attempts
has varied from 1-5% (Bird and Ride, 1981) to 57% (Solomon et al., 2006f) which
most likely reflects differences in the host and pathogen strains used in the different
studies and/or the infection conditions.
The period between the initiation of infection and sporulation is referred to as
the latent period (Jeger et al., 1984). During this time the mycelium ramifies through
the host tissue and initiates a process of host cell destruction resulting in the collapse
of the epidermal and mesophyll cells by 6 dpi (Solomon et al., 2006f). Areas of
necrotic destruction are macroscopically visible 3-5 days after penetration, appearing
as brown oval-shaped lesions. As these increase in size the lesion typically has a light
brown to almost translucent, totally necrotic centre with a darker perimeter
comprising the active area of ongoing cell destruction. From 5-7 days after initial
infection, pycnidia form within the necrotic centre of the lesion. These initially appear
beneath the surface of the leaf and subsequently swell to protrude through the leaf
surface. The mature pycnidium develops an ostiole which eventually ruptures under
the pressure of the pycnidium contents, releasing a pink mucilaginous cirrhus
containing pycnidiospores, which are then ready for splash dispersal by rain
(Douaiher et al., 2004). The whole process of infection to production of new
inoculum takes about 8 days to complete and the potential for serious damage to a
wheat crop comes from the polycyclic nature of the disease. With ideal weather
conditions, regular bouts of infection can exponentially increase pathogen numbers
within a crop.
11
1.3.2 Modes of Host Inoculation
There are three main sources of primary inoculum for SNB infection. The
teleomorph, Phaeosphaeria nodorum, produces wind-dispersed ascospores which
spread genetic variability within the population and can transport the disease over
longer distances (Keller et al., 1997a; Keller et al., 1997b).
Secondly, the anamorph produces asexual pycnidiospores which can survive
outside the host growing season in necrotic tissues, volunteers and alternative hosts,
and initiate a polycyclic, splash-dispersed infection in the following crop (Eyal et al.,
1987).
Thirdly, the fungus can infect the seed, where it can survive as a mycelial
colony, causing disease in the sprouted seedling and leading to subsequent
pycnidiospore-mediated dispersal to neighbouring plants (Baker, 1970). The fungus
has been isolated from harvested seed after 11 years in storage (Cunfer, 1991).
Infection rates of 54%-59% of wheat seed have been reported (Cunfer, 1978;
Turkington et al., 2002) and without seed treatment this can be a major source of
primary inoculum.
1.3.3 Disease Symptoms
The disease is identified in the field by the appearance of chlorotic spots on
the leaf although this symptom is similar to that exhibited by infection with Septoria
tritici. As the disease progresses, the pathogens can be distinguished by the darker
12
colour of the S. tritici pycnidia which give a speckled appearance to the lesion
(Weber, 1922a). Stagonospora nodorum is also distinguished from S. tritici by its
ability to infect all above-ground parts of the plant, with S. tritici being regarded as
limited to infecting leaves (Eyal et al., 1987). However, it has been demonstrated that
under controlled conditions, S. tritici is also able to cause disease in all above-ground
parts of wheat (Jones and Odebunmi, 1971).
1.4 Methods of Disease Control
1.4.1 Chemical Control
The correct application of fungicidal treatments, both in terms of timing and
dosage, is a most effective means of controlling outbreaks of disease. Treatment of
wheat seed with systemic fungicide can suppress S. nodorum for some weeks after
germination as the fungicide is translocated from the seed to the leaves (Cunfer,
1993). Treatment with ergosterol inhibitors such triademinol and difenoconazole
(Bockus and Shroyer, 1998a) and strobilurin inhibitors of mitochondrial respiration
such as azoxystrobin and pyraclostrobin (Jørgensen et al., 1999), or combinations of
these, have been shown to significantly suppress disease progress and significantly
increase yield (Gaurilčikienė and Ronis, 2006). It has also been reported that these
classes of fungicides can induce defence-related genes in wheat (Pasquer et al., 2005).
Treatment of seed rather than foliage also has the advantages of more efficient and
economic application, and the smaller amount of chemical required reduces any final
concentration of resides in the harvest (Bockus and Shroyer, 1998a).
13
The use of chemical control has a number of limitations:
1. it is a significant expense both to purchase and apply and is not universally
economic, especially where the yield is less than 3 tonnes/hectare (Verreet et
al., 2000).
2. there are questions regarding the environmental effects of residues
3. it introduces selection pressure, with the capacity for pathogens to adapt to
azoxystrobin demonstrated (Morzfeld et al., 2004).
1.4.2 Host Resistance/Tolerance
Host plants usually have some genetic resources which confer resistance or
tolerance to a pathogen, and this is considered by some to be the most effective and
economic means of achieving durable resistance (Xu et al., 2004b; Singh et al., 2007).
Resistance refers to host-pathogen interactions which limit the ability of the pathogen
to cause infection, whereas tolerance refers to the ability of the host to maintain its
yield and quality in the face of severe infection (Schafer, 1971). Exploiting host
resistance is advantageous where it is possible, as there is little cost in terms of
application and there is no environmental or dietary implications in terms of chemical
residues (Bockus and Shroyer, 1998a). It has been noted, however, that there is a
potential yield penalty resulting from use of resistant lines (Oliver et al., 2008b).
Strains of wheat which have exhibited some degree of resistance to S.
nodorum have been observed or produced by breeding in the past, and recommended
for use in areas where the pathogen was prevalent (Brown and McNish, 1974),
although the genetic basis of this resistance and the chromosomal location of the
14
genes involved was not known at the time. It has since been shown that resistance to
S. nodorum in bread wheat is inherited as a quantitative trait and efforts are focused
on the genetic mapping of quantitative trait loci (QTLs) to enable improved breeding
for resistance through “Marker Assisted Selection” and gene pyramiding (Servin et
al., 2004; Xu et al., 2004b; Uphaus et al., 2007).
1.4.3 Cultural Practices
Prior to the identification of resistant/tolerant strains of wheat and availability
of effective and economic fungicides, the disease was controlled through cultural
practices designed to reduce the amount of inoculum and minimise the impact of
disease on the mature crop. These measures include the avoidance of early maturing
varieties and later sowing of crops (Sutton, 1920; Brown and McNish, 1974).
Sometimes, however, cultural practises can have antagonistic effects. The growing
practise of reduced till/no till agriculture, for the purposes of reducing soil erosion,
increasing soil moisture, and conserving energy, has the potential to lead to higher
pathogen carryover to the following crop, if crop rotation is not coupled with the
practise (Bockus and Shroyer, 1998). A crop rotation of two years between wheat
crops in Canada is still considered necessary in reducing disease severity (Duczek et
al., 1999), although when environmental conditions are unfavourable to the pathogen,
a rotation of one year may be sufficient (Pedersen and Hughes, 1992).
15
1.4.4 Biological Antagonists
Co-inoculation of S. nodorum with field-equivalent densities of the
saprophytes Aureobasidium pullulans, Sporobolomyces roseus, and Cryptococcus
laurentii var. flavescens, is reported to reduce superficial mycelial growth and
infection of wheat leaves by S. nodorum 50% or more (Fokkema and van der Meulen,
1976).
1.4.5 Genetic Manipulation of S. nodorum
The elucidation of the molecular basis of pathogenicity is considered to be one
of the keys to identifying novel antifungal compounds (Divon and Fluhr, 2007).
Stagonospora nodorum has a haploid genome which has proven amenable to targeted
gene disruption, with transformation protocols well established (Solomon et al.,
2006c). The release of the S. nodorum genome sequence will progress this
understanding through comparative genomics. A recent dramatic example of the
potential of this approach was the discovery of an operon in Escherichia coli
consisting of seven previously uncharacterised genes putatively involved in the
degradation of nucleic acid precurors (Piškur et al., 2007). In the initial analysis of the
S. nodorum genome, there were a large number of genes predicted to encode secreted
proteins (Hane et al., 2007). The importance of this tool for the isolation and
characterisation of these genes cannot be overestimated. To date, a small number of
mutants have been produced in S. nodorum which have been affected in their ability
to initiate and progress SNB in wheat. These include genes involved in metabolism
(discussed further below) and signalling pathways.
16
A gene encoding the Gα sub-unit (Gna1) of the heterotrimeric G protein
involved in signal transduction was disrupted in S. nodorum (Solomon et al., 2004b).
While mutants were able to differentiate hyphopodia, they had decreased protease
activity and were found to be deficient in their ability to penetrate the cuticle.
Although the mutants could still exploit natural openings such as stomata, they were
compromised in their ability to cause infection, and were unable to sporulate. A gene
(Mak2) in the MAP kinase signalling cascade was disrupted in S. nodorum (Solomon
et al., 2005b) with mutants exhibiting a severely altered phenotype. These were
unable to produce hyphopodia and while hyphae were seen to enter via stomata, the
infection did not progress and the strains were essentially non-pathogenic on wheat.
The strain was also unable to sporulate either in planta or in vitro. These two mutants
in different signalling pathways underscore the importance of signalling to the
infection process.
17
1.5 Metabolism and Infection
1.5.1 Definition of Metabolism
Metabolism refers to the operation and integrative functioning of the complex
of metabolic pathways which are required by an organism in order to complete its life
cycle (Vining, 1990). As such it encompasses the anabolic and catabolic manipulation
of molecules, together with the regulation of those processes. There is a distinction
between primary and secondary metabolism. Primary metabolism defines those
pathways which are essential to the basic growth, development and reproduction of an
organism (Mann, 1978). These pathways and metabolites are common to all
organisms (Vining, 1992), although some major differences occur between
prokaryotes and eukaryotes. Secondary metabolism defines the pathways involved in
the production of a vast array of metabolites that are operationally characterised by
being non-essential for growth, specific to particular organisms, and have a wide
range of structures and activities (Idnurm and Howlett, 2001). This differentiation is
artificial and does not imply that pathways of one class are unrelated to, or do not
interact with, pathways of the other class. There are also some which can appear in
both classes.
1.5.2 The Impact of Disrupted Metabolism on Infection
There are a number of S. nodorum mutants which have been produced in
which genes involved in metabolism have been disrupted. These include some which
18
have been affected in their virulence or pathogenicity during the three main stages of
infection.
1.5.2.1 Germination and penetration
The early stages of infection require the pathogen spore to germinate,
recognise the host and initiate metabolism of cell wall degrading enzymes and
proteases. The glyoxylate cycle gene malate synthase (Mls1) was deleted and mutant
spores found to be incapable of germination without the addition of exogenous
glucose or sucrose (Solomon et al., 2004a). This implied that germination and
infection are dependent upon lipid catabolism in order for the glyoxylate cycle and
gluconeogenesis to mobilise energy stores for germination.
The secretion of cell wall degrading enzymes is characteristic of necrotrophic
fungi as a means of penetrating host cells and reducing their contents for nutrient
uptake (Oliver and Ipcho, 2004). The cuticle of plants consists of a complex of
compounds designed to protect it from abiotic stress and biotic attack. Stagonospora
nodorum produces a wide variety of these including xylanases, polygalacturonases,
glucanases, xylosidases, glucosidases and galactosidase, most of them with multiple
isoenzymes (Lehtinen, 1993), as well as a trypsin-like protease (SNP1) (Carlile et al.,
2000). Enzyme family genes are poor candidates for disruption, since the disruption
of one member of the family leaves the others unaffected. Disruption of the Snp1 gene
abolished trypsin activity, but the mutants were unaffected in pathogenicity since an
unknown alkaline protease compensated for the disrupted gene (Bindschedler et al.,
2003).
19
1.5.2.2 Proliferation
Following the initial stage of infection, the pathogen must obtain nutrients
from the host in order to enable it to proliferate, to continue to produce degradative
molecules, and counteract any further host defence responses. A 3-isopropylmalate
dehydrogenase mutant of S. nodorum which was auxotrophic for leucine and lost
pathogenicity to wheat, added to evidence that basic biosynthetic pathways can
contribute to pathogenicity (Cooley et al., 1999). The di/tripeptide transporter gene
(Ptr2), which is upregulated during early infection and is solely responsible for uptake
of degraded peptides, was abolished but without any effect on pathogenicity
(Solomon et al., 2003). Targeted deletion of the gene encoding 5-aminolevulinate
synthase (Als1) produced a mutant which was auxotrophic for δ-aminolaevulinic acid,
a precursor in the synthesis of haem (Solomon et al., 2006a). Mutant strains had
severely stunted germ tubes and became unviable even upon wounded leaves,
indicating a dependence on exogenous stores and synthesis of the compound.
Disruption of the ornithine decarboxylase (Odc1) gene which converts ornithine to the
polyamine putrescine, required for cell division and resistance to oxidative stress,
resulted in mutants with reduced virulence (Bailey et al., 2000). The emerging picture
from these studies is that whilst the fungus apparently has access to large reserves of
host resources at this time, it is still dependent upon its own ability to synthesise many
essential primary metabolites from simple precursors.
Secondary metabolites such as host-specific toxins are important determinants
of host range in pathogens. The ToxA gene of S. nodorum confers pathogen virulence
to wheat varieties containing the Tsn1 gene and culture filtrates from strains in which
20
ToxA had been ablated were non-toxic (Friesen et al., 2006). Further studies have
shown the presence of multiple S. nodorum toxins each of which are required to
interact with specific wheat gene products in an inverse gene-for-gene fashion in
order to cause disease (Friesen et al., 2007). This is an exciting new area of research
with immediate applicability in the field.
1.5.2.3 Sporulation
Sporulation completes the life cycle of the phytopathogenic fungus and as
noted above, signalling pathways play an important role in this process. An
investigation of the calcium/calmodulin-dependent protein kinases in S. nodorum
demonstrated that while CpkB was redundant for pathogenicity, disruption of the
CpkA gene resulted in an inability to complete differentiation of pycnidia, while
disruption of the CpkC gene resulted in delayed lesion development and sporulation
(Solomon et al., 2006d).
The disaccharide trehalose was found to be upregulated during sporulation and
the gene encoding trehalose 6-phosphate synthase (Tps1) was inactivated by targeted
gene deletion (Lowe, 2006). Mutants were almost completely deficient in trehalose
accumulation, and while lesion development was only slightly affected, sporulation
was reduced to 30% of wild type levels.
Stagonospora nodorum mutants in which the mannitol 1-phosphate 5dehydrogenase (Mpd1) gene was disrupted were phenotypically similar to the wild
type except that they were unable to sporulate in planta (Solomon et al., 2005a). A
21
second gene involved in mannitol metabolism, Mdh1, which encodes mannitol 2dehydrogenase, was subsequently inactivated in S. nodorum (Waters, 2004), and
found to be phenotypically identical to the wild type. The observed behaviours of
these two mutants implied a role in pathogenicity for the compound itself, and also
contradicted the accepted theory of how mannitol metabolism occurs as outlined
below.
1.6 Mannitol Metabolism and Infection
1.6.1 Postulated Roles of Mannitol
The acyclic 6-carbon polyol D-mannitol is the most abundant soluble
metabolite found in many filamentous fungi (Lewis and Smith, 1967). There have
been a number of postulated roles for mannitol, not necessarily mutually exclusive,
including:
•
Carbohydrate storage (Birkinshaw et al., 1931; Corina and Munday, 1971)
•
Translocation of carbon (Trip et al., 1964; Lewis and Smith, 1967; Koide et
al., 2000)
•
Compatible solute/protein protection (Stoop and Mooibroek, 1998; Ortbauer
and Popp, 2008)
•
Storage of reducing power (Lewis and Smith, 1967; Ruijter et al., 2003)
•
Co-enzyme regulation/NADPH regeneration (Hult and Gatenbeck, 1978;
Diano et al., 2006)
•
Energy dissipation by futile cycling (Jennings and Burke, 1990)
22
•
Morphogenesis/conidiation/spore discharge (Corina and Munday, 1971;
Webster et al., 1995; Trail et al., 2005)
•
Environmental stress, including quenching of reactive oxygen species
produced as a defence response by the host (Chaturvedi et al., 1996; Stoop and
Mooibroek, 1998; Ruijter et al., 2003; Bois et al., 2006)
•
Host carbon sequestration (Joosten et al., 1990; Noeldner et al., 1994)
These last three imply a specific role for mannitol in the process of infection.
While there is circumstantial evidence which supports some of the above roles, almost
none have been experimentally proven. Mannitol is not the sole candidate compound
for some of these roles - other polyols have better experimental support for some roles
in different fungal species (see Solomon et al. (2007) for a summary of these
arguments) - and the absence of mannitol in some species (Lewis and Smith, 1967),
particularly laboratory strains of Saccharomyces cerevisiae, indicates that alternative
compounds must be able to compensate.
1.6.2 Enzymatic Metabolism of Mannitol in Fungi
There are a number of enzymes which have been demonstrated or purported to
be involved in the metabolism of mannitol in fungi. A summary of the organisms in
which these enzymes have been reported is given in Table 1.3 and the enzymes are
outlined below.
23
Table 1.3: Distribution of genes reported as being involved in the metabolism of mannitol in fungal species (see below for abbreviations and notes).
Species
Phylum2 Hex
F6PP
Alternaria alternata
A
Y
Aspergillus candidus
Aspergillus clavatus
Aspergillus fumigatus
Aspergillus nidulans
A
A
A
A
Aspergillus niger
Aspergillus oryzae
M1Pdh
M1PP MK
MPPT
MAPT
References
-
NADP- NAD- Mdh
Mdh
Mdh
Y
N
-
Y
Y
-
-
-
Y
Y
-
Y
Y
N
-
-
Y
Y
Y
Y
Y
Y
N
-
-
-
A
Y
-
Y
Y
-
Y
Y
-
-
-
A
-
-
Y
Y
-
Y
Y
-
-
-
Aspergillus parasiticus A
-
-
Y
-
-
Y
-
-
-
-
Aspergillus sp.
(UC4177)1
Botrytis cinerea
Candida magnoliae
Candida utilis
A
Y
-
N
N
-
Y
Y
N
N
Y
A
A
A
Y
Y
-
Y
Y
Y
N
N
-
Y
N
Y
Y
-
-
-
(Hult and Gatenbeck, 1978; Hult
and Gatenbeck, 1979; Hult et
al., 1980; Schneider et al., 2006;
Vélëz et al., 2007)
(Strandberg, 1969)
(Corina and Munday, 1971)
(Boonsaeng et al., 1976)
(Hankinson, 1974; Bailey and
Arst, 1975; Hankinson and
Cove, 1975; Singh et al., 1988)
(Yamada et al., 1959;
Boonsaeng et al., 1976; Hult et
al., 1980; Kiser and Niehaus,
1981; Foreman and Niehaus,
1985; Ruijter et al., 2003)
(Yamada et al., 1959; Horikoshi
et al., 1965; Boonsaeng et al.,
1976; Ruijter et al., 2004)
(Niehaus and Dilts, 1982;
Buchanan and Lewis, 1984b;
Buchanan and Lewis, 1984a;
Foreman and Niehaus, 1985;
Niehaus and Jiang, 1989)
(Lee, 1967a; Lee, 1967b; Lee,
1970)
(Hult et al., 1980)
(Lee et al., 2003)
(Hult et al., 1980)
Continued on following page
Table 1.3: Distribution of genes reported as being involved in the metabolism of mannitol in fungal species (cont.) - (see below for abbreviations and notes).
Species
Phylum2 Hex
Cenococcum
graniforme
Ceratocystis
multiannulata
Chaetomium globosum
Chaetomium
thermophile var.
dissitum1
Cladosporium
cladosporioides
Cladosporium fulvum
Cladosporium
herbarum
Coccidioides immitis
Dendryphiella salina
Diplodia viticola
Geotrichum candidum
Gibberella zeae
A
Y3
A
F6PP
M1Pdh
M1PP MK
MPPT
MAPT
References
-
NADP- NAD- Mdh
Mdh
Mdh
Y
Y
-
Y9
-
-
-
-
(Martin et al., 1985)
Y
-
Y
N
-
N
Y
-
-
-
(Hult et al., 1980)
A
A
Y3
-
-
N
-
Y
-
-
N9
Y
-
N
-
N
-
N
-
(Adomako et al., 1972)
(Boonsaeng et al., 1976)
A
Y
-
Y
Y
-
Y
Y
-
-
-
(Hult et al., 1980)
A
A
-
-
Y
Y
-
-
-
-
-
-
-
(Noeldner et al., 1994)
(Simon-Nobbe et al., 2006)
A
A
A
A
A
Y
-
Y
Y
Y
Y
Y
N
N
N
-
Y
N9
N
N
Y
N
-
-
-
Hypogymnia physodes
Magnaporthe syn.
Pyricularia oryzae
Malbranchea
pulchella var. sulfurea
Microsporum gypseum
Neurospora crassa
A
A
Y
-
Y/N
Y/N
N
Y
Y
Y
-
-
-
A
-
-
-
-
-
Y
-
-
-
-
(Lones and Peacock, 1964)
(Holligan and Jennings, 1972)
(Strobel and Kosuge, 1965)
(Chang and Li, 1964)
(Hult et al., 1980; Trail and Xu,
2002; Trail et al., 2002)
(Jensen et al., 1991)
(Yamada et al., 1959; Yamada
et al., 1961; Hult et al., 1980)
(Boonsaeng et al., 1976)
A
A
Y3,4
Y
-
Y
Y
Y
-
Y
N10
Y
Y
Y
-
Y
-
-
Neurospora sitophila
A
-
-
-
-
-
N
Y
-
-
-
(Leighton et al., 1970)
(Yamada et al., 1959; Hult et
al., 1980)
(Yamada et al., 1959)
Continued on following page
Table 1.3: Distribution of genes reported as being involved in the metabolism of mannitol in fungal species (cont.) - (see below for abbreviations and notes).
Species
Phylum2 Hex
F6PP
Paracoccidioides
brasiliensis
Penicillium
chrysogenum syn.
notatum
Penicillium cyclopium
Penicillium duponti1
A
-
A
M1Pdh
M1PP MK
MPPT
MAPT
References
-
NADP- NAD- Mdh
Mdh
Mdh
-
Y
-
-
-
-
(Castro et al., 2002)
-
-
-
Y
-
Y
Y
-
-
-
A
A
-
-
-
-
-
Y
Y
-
-
-
-
A
Y
-
Y
N
-
Y
Y
-
-
-
(Ballio et al., 1964; Boonsaeng
et al., 1976; Boonsaeng et al.,
1977; Boutelje et al., 1983)
(Boonsaeng et al., 1976)
(Boonsaeng et al., 1976;
Boonsaeng et al., 1977)
(Hult et al., 1980)
A
A
A
Y
-
-
Y
-
Y
-
-
Y
Y
Y
Y
-
-
-
-
(Hult et al., 1980)
(Boonsaeng et al., 1976)
(Jensen et al., 1991)
A
-
-
-
-
-
Y
Y
-
-
-
Saccharomyces
cerevisiae
A
-
-
Y
Y
-
-
-
-
-
-
Sclerotinia
sclerotiorum
Sphaerosporella
brunnea
Stagonospora
nodorum
Trichothecium roseum
A
-
-
Y
-
-
Y
Y
-
-
-
(Wright and Le Tourneau, 1966;
Aitken et al., 1969a; Aitken et
al., 1969b)
(Kulbe et al., 1986; Quain and
Boulton, 1987; Perfect et al.,
1996)
(Wang and Le Tourneau, 1972)
A
Y
-
Y
Y
-
Y
Y
-
-
-
(Ramstedt et al., 1987)
A
-
-
Y
-
-
Y
-
N11
-
-
A
Y
-
Y
Y
-
Y
Y
-
-
-
(Solomon et al., 2005; Solomon
et al., 2006b)
(Hult et al., 1980)
Penicillium glabrum
syn. frequentans
Penicillium islandicum
Penicillium urticae
Pseudevernia
furfuracea
Pyrenochaeta
terrestris
Continued on following page
Table 1.3: Distribution of genes reported as being involved in the metabolism of mannitol in fungal species (cont.) - (see below for abbreviations and notes).
Species
Phylum2 Hex
F6PP
Thermomyces
lanuginosus syn.
Humicola lanuginosa
Tuber borchii
A
Y
A
Agaricus bisporus
M1Pdh
M1PP MK
MPPT
MAPT
References
-
NADP- NAD- Mdh
Mdh
Mdh
Y
Y
-
Y
Y
-
-
-
(Boonsaeng et al., 1976; Hult et
al., 1980)
Y5
-
Y
-
-
Y
Y
-
-
-
B
Y6
Y
Y
N
-
N
N
-
-
-
Agaricus campestris
B
-
-
Y
Y
-
-
-
-
-
-
Amanita muscaria
Armillariella mellea
Chondrostereum
purpureum
Coprinus cinereus
Cryptococcus
neoformans
Flammulina velutipes
Fomes pinicola
Heterobasidion
annosum
B
B
B
Y
Y
Y
-
Y
N
N
Y
N
Y
-
N
N
N
Y
N
N
-
-
-
(Ceccaroli et al., 1999;
Ceccaroli et al., 2003; Ceccaroli
et al., 2007)
(Hammond, 1977; Ruffner et
al., 1978; Hult et al., 1980;
Hammond, 1981; Morton et al.,
1985a; Morton et al., 1985b;
Stoop and Mooibroek, 1998;
Wannet et al., 1999; Hörer et
al., 2001; Sassoon et al., 2001;
Sassoon and Mooibroek, 2001)
(Edmundowicz and Wriston,
1963)
(Ramstedt et al., 1987)
(Ramstedt et al., 1987)
(Ramstedt et al., 1987)
B
B
-
-
Y
Y
Y
-
Y
-
-
-
-
B
B
B
Y
Y
-
N
N
N
Y
Y
Y
-
N
N
N
N
N
-
-
-
(Nyunoya et al., 1984)
(Perfect et al., 1996; Suvarna et
al., 2000; Loftus et al., 2005)
(Kitamoto et al., 2000)
(Hult et al., 1980)
(Ramstedt et al., 1987)
Continued on following page
Table 1.3: Distribution of genes reported as being involved in the metabolism of mannitol in fungal species (cont.) - (see below for abbreviations and notes).
Species
Phylum2 Hex
F6PP
Laccaria laccata
B
Y
Lentinus edodes
Marasmius
scorodonius
Melampsora lini
Mycena metata
Phanerochaete
chrysosporium
Piloderma croceum
B
B
M1Pdh
M1PP MK
MPPT
MAPT
References
-
NADP- NAD- Mdh
Mdh
Mdh
Y
N
-
N
Y
-
-
-
Y7
Y
Y
-
Y
N
Y
-
N
N
Y
Y
-
-
-
(Ramstedt et al., 1987; Deveau
et al., 2008)
(Kulkarni, 1990)
(Ramstedt et al., 1987)
B
B
B
Y
-
-
Y
N
-
Y
Y
-
-
N
Y
Y
-
-
-
-
(Clancy and Coffey, 1980)
(Ramstedt et al., 1987)
(Matsuzaki et al., 2008)
B
Y
-
Y
Y
-
N
Y
-
-
-
Pisolithus tinctorius
Pleurotus ostreatus
B
B
Y7
Y
Y
-
-
Y
Y
Y
-
-
-
Polyporus versicolor
Puccinia graminis
Schizophyllum
commune
B
B
B
Y
-
-
N
-
N
Y
Y
-
N
-
N
-
-
-
-
Suillus bovinus
Suillus variegatus
Uromyces phaseoli1
Uromyces viciae-fabae
syn. fabae
Absidia glauca
B
B
B
B
Y
Y
-
-
Y
Y
Y
Y
N
N
Y
-
-
N
N
-
N
Y
-
-
-
-
(Ramstedt et al., 1986; Ramstedt
et al., 1987)
(Kong et al., 2000)
(Chakraborty et al., 2003;
Chakraborty et al., 2004)
(Hult et al., 1980)
(Maclean, 1971)
(Niederpruem et al., 1965;
Isenberg and Niederpruem,
1967)
(Ramstedt et al., 1987)
(Ramstedt et al., 1987)
(Wynn, 1966)
(Voegele et al., 2005)
IS
-
-
-
Y
-
Y
-
Y
-
-
Amylomyces syn.
Mucor rouxii
IS
-
-
-
-
-
Y
-
-
-
-
(Ueng et al., 1976; Ueng and
McGuinness, 1977)
(Boonsaeng et al., 1976)
Continued on following page
Table 1.3: Distribution of genes reported as being involved in the metabolism of mannitol in fungal species (cont.) - (see below for abbreviations and notes).
Species
Phylum2 Hex
F6PP
NADP- NAD- Mdh
Mdh
Mdh
Y
-
M1Pdh
M1PP MK
MPPT
MAPT
References
IS
(Birken and Pisano, 1976)
Cephalosporium
chrysogenum?
Y
Y
Y
N
N
(Hult et al., 1980)
Mucor circinelloides f. IS
lusitanicus
Mucor genevensis1
IS
N
(Boonsaeng et al., 1976)
IS
Y
N
N
N
N
(Hult et al., 1980)
Phycomyces
blakesleeanus
Rhizomucor syn.
IS
N
(Boonsaeng et al., 1976)
Mucor miehei
Rhizomucor syn.
IS
N
(Boonsaeng et al., 1976)
Mucor pusillus
IS
Y
N
N
N
N
(Hult et al., 1980)
Rhizopus arrhizus
Abbreviations: Hex – hexokinase; F6PP – fructose 6-phosphate phosphatase; NADP-Mdh – NADP+-dependent mannitol dehydrogenase (EC 1.1.1.138); NAD-Mdh – NAD+dependent mannitol dehydrogenase (EC 1.1.1.67); Mdh – mannitol dehydrogenase (co-factor not specified); M1Pdh – mannitol 1-phosphate dehydrogenase (EC 1.1.1.17);
M1PP – mannitol-1-phosphate phosphatase (EC 3.1.3.22); MK – mannitol kinase; MPPT – mannitol phosphoenolpyuvate phosphotransferase; MAPT – mannitol acyl
phosphotransferase; A – ascomycota; B – basidiomycota; IS – fungi incertae sedis; Y = activity detected; N = activity not detected; Y/N = activity present in some studies but
not in others; - = activity not assayed.
1
Organisms not found in Taxbrowser database (NCBI, 2008). Names have been given as reported.
2
Some organisms were originally reported from obsolete phyla e.g. phycomyces. The currently accepted phylum as assigned in Taxbrowser (NCBI, 2008) is used here. In the
case of organisms noted in the previous note – these were assigned to phyla based on their genus.
3
Reported a separate fructokinase activity in addition to hexokinase
4
Glucokinase activity was reported to be greater than fructokinase activity
5
Three types of hexokinase activity reported, expressed at different stages of development
6
Three studies report hexokinase activity, one study reports fructokinase activity
7
Only reported fructokinase activity
8
One study reports hexokinase, the other specifies fructokinase
9
Both co-factors assayed
10
A gene identified as M1PDH was reported in the genome sequence of Neurospora crassa (Galagan et al., 2003)
11
pers. comm., Dr. P. Solomon
1.6.2.1 D-mannitol:NADP+ 2-oxidoreductase (EC 1.1.1.138)
Synonyms: mannitol 2-dehydrogenase (NADP); mannitol 2-dehydrogenase
(NADP+); NADP-mannitol dehydrogenase; mannitol dehydrogenase (the last of these
will be used hereafter).
Reaction:
D-fructose + NADPH + H+
D-mannitol + NAPD+
The first identification of this enzyme in fungi resulted from an investigation
into mannitol metabolism in the basidiomycete Agaricus campestris (Edmundowicz
and Wriston, 1963). It was first purified and characterised in Aspergillus parasiticus
(Niehaus and Dilts, 1982). In Aspergillus nidulans, this enzyme was shown to be
localised exclusively in the cytosol (Singh et al., 1988). The enzyme was first
inactivated by gene disruption in Stagonospora nodorum (Waters, 2004) and mutants
found to be phenotypically identical to the wild type.
1.6.2.2 D-mannitol:NAD+ 2-oxidoreductase (EC 1.1.1.67)
Synonyms: mannitol 2-dehydrogenase; D-mannitol dehydrogenase; mannitol
dehydrogenase; NAD-mannitol dehydrogenase (the last of these will be used
hereafter)
Reaction:
D-fructose + NADH + H+
D-mannitol + NAD+
30
In cases where NAD-mannitol dehydrogenase has been detected in addition to
the NADP-linked mannitol dehydrogenase, its activity has been lower (5-10% in A.
campestris and 25% in Aspergillus oryzae) than that of the NADP-linked enzyme
(Edmundowicz and Wriston, 1963; Horikoshi et al., 1965).
1.6.2.3 D-mannitol-1-phosphate:NAD+ 5-oxidoreductase (EC 1.1.1.17)
Synonyms: hexose reductase; mannitol-1-phosphate 5-dehydrogenase; Dmannitol-1-phosphate dehydrogenase; fructose 6-phosphate reductase; mannitol 1phosphate dehydrogenase (the last of these will be used hereafter)
Reaction:
D-fructose 6-phosphate + NADH + H+
D-mannitol 1-phosphate + NAD+
Activity corresponding to this enzyme was first reported in fungi in
Magnaporthe (syn. Pyricularia) oryzae, A. oryzae and A. niger (Yamada et al., 1959).
It was purified and characterised in terms of its kinetic parameters in A. niger and
found to be highly specific for its substrates and co-factors (Kiser and Niehaus, 1981).
Two isoenzymes were detected in A. nidulans, one of which was localised in the
cytosol, and the other apparently present on the outer face of the inner mitochondrial
membrane (Singh et al., 1988). The MpdA gene encoding this enzyme in the saprobe
A. niger was inactivated, with a reduction of mannitol concentration in conidiospores
to 30% of wild type levels (Ruijter et al., 2003). Conidiospores were found to be
extremely sensitive to heat stress and oxidative stress, demonstrating a role for
31
mannitol in this non-pathogen species. The mpd1 mutants of S. nodorum had mannitol
reduced to 20% of the wild type, and lost the ability to sporulate in planta, the first
demonstrated role for mannitol metabolism in pathogenicity (Solomon et al., 2005a).
1.6.2.4 D-mannitol-1-phosphate phosphohydrolase (EC 3.1.3.22)
Synonyms: mannitol-1-phosphatase; mannitol-1-phosphate phosphatase (the
last of these will be used hereafter).
Reaction:
D-mannitol 1-phosphate + H20
D-mannitol + Pi
Activity corresponding to this enzyme was first reported in fungi in M. oryzae
and activity also noted in the crude extracts of A. oryzae, A. niger, Neurospora crassa
and N. sitophila (Yamada et al., 1959). It has been shown to be localised exclusively
in the cytosol in A. nidulans (Singh et al., 1988).
1.6.2.5 D-mannitol kinase (EC 2.7.1.57 (created 1972, deleted 1984))
Reaction (theoretical):
D-mannitol + ATP Æ D-mannitol 1-phosphate + ADP
While evidence of activity corresponding to this enzyme was reported for
Absidia glauca (Ueng et al., 1976) and Microspora gypseum (Leighton et al., 1970),
in the majority of studies where such an enzyme was investigated in fungi, there was
32
no activity reported (Lones and Peacock, 1964; Lee, 1967b; Strandberg, 1969;
Adomako et al., 1972). Mannitol kinase activity was reported in a number of bacteria
(Klungsøyr, 1966; Mehta et al., 1977), however, no gene has been forwarded as a
candidate for transcription of this enzyme, and experimental evidence for its existence
is poor. It is possible that there are one or more non-specific kinases which could
contribute independently or in concert to such an activity.
1.6.2.6 D-mannitol acetyl phosphate phosphotransferase (no EC number)
Activity corresponding to this enzyme was reported in an Aspergillus strain by
which mannitol was phosphorylated to mannitol 1-phosphate (Lee, 1967b). Acetyl
phosphate and carbamyl phosphate were both able to serve as phosphate donors in the
reaction. No activity for this enzyme was found in Chaetomium globosum (Adomako
et al., 1972).
1.6.2.7 D-mannitol phosphoenolpyruvate phosphotransferase (no EC number)
This enzyme is considered to be part of the bacterial system of mannitol
catabolism (Ramstedt et al., 1986). There is a single report of activity corresponding
to this enzyme system in fungi in M. gypseum (Leighton et al., 1970). No activity for
this enzyme was found in C. globosum (Adomako et al., 1972) or Aspergillus (Lee,
1967b).
33
1.6.2.8 Hexokinases
This group of enzymes participates in the pathway of mannitol metabolism by
phosphorylating fructose to fructose 6-phosphate, or glucose to glucose 6-phosphate
(the glycolytic precursor of fructose 6-phosphate) and thereby providing the substrate
for mannitol 1-phosphate dehydrogenase. The term may refer to any or all of three
enzymes recognised by the Enzyme Commission. These are glucokinase (EC 2.1.1.2)
which is specific for conversion of glucose to glucose 6-phosphate; fructokinase (EC
2.1.1.4) which is specific for conversion of fructose to fructose 6-phosphate; and
hexokinase (EC 2.1.1.1) which can catalyse both reactions.
1.6.2.9 D-fructose-6-phosphate phosphatase (no EC number)
Synonyms: fructose-6-phosphatase
Reaction:
D-fructose 6-phosphate + H2O Æ D- fructose + Pi
Perhaps due to a general acceptance of the mannitol cycle (described below)
and its unidirectional operation, there has been less attention given to this reaction
which reverses the hexokinase-catalysed step. Activity corresponding to this enzyme
has been reported in three basidiomycetes – Agaricus bisporus (Morton et al., 1985b),
Lentinus edodes (Kulkarni, 1990) and Pleurotus ostreatus (Chakraborty et al., 2004).
The enzyme has not yet been purified and characterised, and there is no gene put
forward as encoding such an enzyme.
34
1.6.3 The Postulated Mannitol Cycle
The proposition that the metabolism of mannitol in fungi occurred in a
unidirectional enzymatic cycle was first proposed by Hult & Gatenbeck (1978). In
this cycle (Figure 1.3A) fructose 6-phosphate was converted to mannitol 1-phosphate
by mannitol 1-phosphate dehydrogenase, with the latter compound dephosphorylated
to mannitol by mannitol 1-phosphate phosphatase. Mannitol was then converted to
fructose by mannitol dehydrogenase, and in the final step fructose was phosphorylated
to fructose 6-phosphate by hexokinase (Hult and Gatenbeck, 1978). The net result of
the cycle was the stoichiometric regeneration of NADPH at the expense of NADH
and ATP, and NADPH regeneration was given as the main purpose of the cycle. Since
the mannitol 1-phosphate phosphatase-catalysed reaction was assumed to be
irreversible, the cycle could only proceed in one direction. A variation of this cycle
was proposed in which the direct conversion of fructose to fructose 6-phosphate was
replaced by several steps involving the conversion of fructose to glucitol (sorbitol),
followed by glucose, glucose 6-phosphate, and then fructose 6-phosphate (Jennings
and Burke, 1990) (Figure 1.3b).
The existence and importance of the mannitol cycle gained steady acceptance
(Martin et al., 1985; Niehaus and Jiang, 1989; Schmatz et al., 1989; Michalski et al.,
1992; Schmidt et al., 1998; Allocco et al., 1999; Ceccaroli et al., 2003) in the absence
of a means of scientifically falsifying the theory. There was some questioning of the
fact and significance of the cycle based on subcellular location of the enzymes, and an
observed lack of coordination in their maximal activities in Aspergillus nidulans
35
A
HEX
MDH
MPD
MPP
B
MPD
PGI
MPP
HEX
MDH
SDH
AR
Figure 1.3: (A) The mannitol enzymatic cycle as proposed by Hult and Gatenbeck
(1978). Figure as given in Hult et al. (1980).
(B) The modified mannitol cycle proposed by Jennings and Burke (1990).
The enzymes for each step have been added in red. Abbreviations: AR = aldose
reductase (EC 1.1.1.21); HEX = hexokinase; MDH = mannitol dehydrogenase (EC
1.1.1.138); MPD = mannitol 1-phosphate dehydrogenase (EC 1.1.1.67); MPP =
mannitol-1-phosphate phosphatase; PGI = phosphoglucose isomerase; SDH = sorbitol
(glucitol) dehydrogenase (EC 1.1.1.14). Note the proposed unidirectional operation of
the cycles.
36
(Singh et al., 1988). The involvement of NADH in a synthetic reaction was
questioned, as was the need for a pathway of NADPH re-generation in addition to that
of the pentose phosphate pathway, and the fact that the cycle required fructose to be a
better substrate for hexokinase than glucose (Jennings and Burke, 1990). Furthermore,
the proposers of the cycle themselves were among those who examined a range of
fungal species for existence of the cycle and found that basidiomycetes in general did
not have the full complement of enzymes (Hult et al., 1980; Ramstedt et al., 1987;
Kulkarni, 1990; Kitamoto et al., 2000; Deveau et al., 2008). Mannitol 1-phosphate
dehydrogenase activity was not detected in basidiomycetes in any of these studies and
mannitol 1-phosphate phosphatase was not detected in the majority. The only reported
exceptions to this were:
1. Pleurotus ostreatus, in which all enzymes of the proposed cycle were detected
(Chakraborty et al., 2004)
2.
Cryptococcus neoformans, from which Mpd1 was purified by Suvarna et al.
(2000), but the presence of other enzymes of the mannitol cycle was not
investigated. The recently released genome sequence contains two genes
reported as encoding Mpd1 (Loftus et al., 2005)
3. Phanerochaete chrysosporium, in which Mpd1 was found to be upregulated
following growth on benzoic acid (Matsuzaki et al., 2008)
The reported presence in many fungal species, of an NAD+-dependent
mannitol dehydrogenase, instead of, or in addition to, the NADP+-dependent enzyme
required for the operation of the cycle (Table 1.3), seems to have been unaddressed in
terms of its implications for such a cycle. Where the NAD+-dependent enzyme alone
was found, as reported for the ascomycete Chaetomium globosum (Adomako et al.,
37
1972) and the basidiomycetes Marasmius scorodonius and Mycena metata (Ramstedt
et al., 1987), the cycle would be unable to regenerate NADPH as proposed. An
alternative and extended model of the mannitol cycle was proposed in which the
direct conversion of fructose to fructose 6-phosphate was intercalated with steps
involving the conversion of fructose to sorbitol, followed by glucose, glucose 6phosphate, and then fructose 6-phosphate (Figure 1.3B) (Jennings and Burke, 1990).
Improved methods in targeted gene inactivation, and the existence of strains of
fungi which are amenable to such methods, have offered the means by which the
existence of the mannitol cycle may be conclusively investigated. The mannitol 1phosphate dehydrogenase gene was inactivated in Aspergillus nidulans (Ruijter et al.,
2003) and Stagonospora nodorum (Solomon et al., 2005a) prior to the
commencement of this study. While mannitol production was reduced in the mutant
strains to 30% and 20% respectively of the wild types, it was evident that mannitol
synthesis was possible in the absence of this enzyme. The first inactivation of a
mannitol 2-dehydrogenase occurred in S. nodorum and was performed by the author
as part of an Honours project (Waters, 2004). Although no mannitol 2-dehydrogenase
activity could be detected in the mutant strain, it was otherwise phenotypically
identical to the wild type, including the ability to grow on mannitol as a sole carbon
source, offering the first evidence that the mannitol cycle does not exist as proposed,
and suggesting a hitherto unknown means by which mannitol is catabolised. The
possibility of creating a mutant harbouring both disruption constructs presented the
opportunity of abolishing mannitol synthesis entirely and elucidating a role for
mannitol metabolism in the infection process.
38
1.7 Summary and Aims
Stagonospora nodorum is a potent and economically significant necrotrophic
pathogen of wheat. The availability of the genome sequence places an upper limit on
the number and identity of genes. Targeted gene deletions have demonstrated the
roles of several metabolites which are important during the three main stages of
infection. Some of these have been suggested by the work of other studies, while
others are the result of noted expression differences between EST libraries, alterations
in gene expression and changes in metabolite abundance over the course of infection.
The aims of this project were to further probe the relationship between
metabolism and infection in the Stagonospora nodorum-wheat pathosystem with
particular reference to mannitol. A genetics approach was used initially to create a
double mutant strain harbouring the disrupted mannitol dehydrogenase gene and
mannitol-1-phosphate dehydrogenase gene. It was hypothesised that this would
abolish the ability to synthesise or catabolise mannitol and would elucidate a role for
this compound in infection. The strain was phenotypically characterised using
standard in vitro and in planta growth assays and pathogenicity assays as established.
Nuclear magnetic resonance techniques, including the use of
13
C-labelled substrates,
were used to investigate changes in principal soluble metabolites between the wild
type and mutant strains, and to gain an understanding of their pathways of
metabolism. GC-MS metabolite profiling was used to observe changes in metabolic
profile over the course of an infection by the wild type strain. It was a further aim of
this study that it would suggest anti-fungal strategies by exposing weaknesses in the
fungus’ life-cycle.
39
CHAPTER 2 – GENERAL MATERIALS AND
METHODS
40
2.1 Fungal and Bacterial Strains
Stagonospora nodorum strain SN15 was supplied by the Department of
Agriculture, Western Australia, (now the Department of Agriculture and Food,
Western Australia).
A S. nodorum mutant strain, mpd1-1, harbouring a disrupted mannitol 1phosphate dehydrogenase gene was the generous gift of Dr. Peter Solomon (Solomon
et al., 2005a).
The S. nodorum mutant strains mdh1-67, mdh1-71, mdh1-73, mdh1-78, mdh179 and Mdh1-63e, each harbouring a disrupted mannitol 2-dehydrogenase gene, were
previously created by the author (Waters, 2004; Solomon et al., 2007).
Escherichia coli DH10B (Grant et al., 1990) (Invitrogen Corporation,
Carlsbad, CA) containing the disruption construct pGPSH-Mpd8 was the generous
gift of Dr. Peter Solomon. This construct comprises an insertionally mutagenised
mannitol 1-phosphate dehydrogenase gene and confers resistance to hygromycin
(Solomon et al., 2005a).
2.2 Wheat Variety
The SN15-susceptible wheat cultivar, Triticum aestivum L. subsp. aestivum
cv. Amery was used in pathogenicity assays and was supplied by the Department of
41
Agriculture, Western Australia, (now the Department of Agriculture and Food,
Western Australia).
2.3 General Media
All media used within this study are listed in Table 2.1. All reagents used were
of analytical grade. All water used was of milliQ quality and sterilised by autoclaving
unless otherwise indicated. All autoclaving was carried out for 20 min at 121 °C and
100 kPa.
2.4 Growth of Triticum aestivum cv. Amery
Seeds were surface-sterilised for 5 min in sterilisation solution (1% bleach, 5%
ethanol) followed by rinsing in sterile water. Sterile pots (125-130mm) were prepared
by adding Expanded Perlite – Coarse (The Perlite & Vermiculite Factory, Jandakot)
to the height of the drainage holes and filling the remainder of the pot to within
approximately 1.5 cm of the top with vermiculite (The Perlite & Vermiculite Factory,
Jandakot). Seeds were sown with 8 seeds per pot for a whole plant spray, or 50-60
seeds per pot for detached leaf assay – and covered with approximately 1 cm
vermiculite. Pots were placed 8 to a sterile tray (Nally, NSW) and the tray filled with
2-3 cm of tap water. Seedlings were grown for 2 weeks in a growth chamber at 23 °C
and a lighting regime of alternating 12 h light and 12 h dark. Lighting was provided
by fluorescent tubes, 50% of which were GRO- LUX® F36W/Grow T8 (Sylvania,
Germany) and the remainder Cool White L36 W/20 (Osram, Germany), with the two
different types being mounted alternately.12
42
Table 2.1: Media used in this study.
Medium
Ingredients
Benzimidazole Agar
1% w/v BBLTM agar (Becton, Dickinson & Co., USA)
100 mg.L-1 benzimidazole (ICN Biochemicals Inc, Aurora, USA)
Complete Supplement
20 g.L-1 Casamino acids (Becton, Dickinson & Co., USA)
(for CzV8CS media)
20 g.L-1 Peptone (Becton, Dickinson & Co., USA)
20 g.L-1 Yeast extract (Becton, Dickinson & Co., USA)
3 g.L-1 Adenine (Sigma Chemical Co., St. Louis, USA)
0.02 g.L-1 Biotin (Sigma-Aldrich Inc.., St. Louis, USA)
0.02 g.L-1 Nicotinic acid (Sigma Chemical Co., St. Louis, USA)
0.02 g.L-1 p-aminobenzoic acid (Sigma Chemical Co., USA)
0.02 g.L-1 Pyridoxine (Sigma Chemical Co., St. Louis, USA)
0.02 g.L-1 Thiamine (Sigma Chemical Co., St. Louis, USA)
Filter sterilise with 0.2 µm filter and store in fridge
CzV8CS Agar
45.4 g.L-1 Czapek Dox Agar (Oxoid)
10 g.L-1 BBLTM agar (Becton, Dickinson & Company)
3 g.L-1 CaCO3 (Chem-Supply, Gillman, South Australia)
200 mL.L-1 filtered V8 Juice (Campbell’s)
pH 6.0
Autoclave
50 mL.L-1 Complete Supplement (see above)
50 μg.mL-1 phleomycin (Cayla, Toulouse) where required
200 μg.mL-1 hygromycin-B (Roche, Mannheim) where required
CzV8CS Liquid Culture Medium
45.4 g.L-1 Czapek Dox Liquid Medium (Oxoid)
(for flask culture)
200 mL.L-1 centrifuged V8 juice (Campbell’s)
pH 6.0
Aliquot 95 mL per 250 mL flask
Autoclave
Add 5 mL.100 mL-1 complete supplement prior to use
CzV8-Proto Agar
45.4 g.L-1 Czapek Dox Agar (Oxoid)
10 g.L-1 BBLTM agar (Becton, Dickinson & Co., USA)
200 mL.L-1 centrifuged V8 Juice (Campbell’s)
182.2 g.L-1 sorbitol (Univar International Ltd, Poole, England)
pH 6.0
Autoclave and pour ~15 mL per plate
Continued on the following page
43
Table 2.1: (Continued)
Medium
Ingredients
CzV8-Proto Top Agar
45.4 g.L-1 Czapek Dox Agar (Oxoid)
7.5 g.L-1 BBLTM agar (Becton, Dickinson & Co., USA)
200 mL.L-1 centrifuged V8 Juice (Campbell’s)
182.2 g.L-1 sorbitol (Univar International Ltd, Poole, England)
pH 6.0
Autoclave
Luria-Bertani (LB) Broth
1% w/v bacto-peptone (Becton, Dickinson & Co., USA)
0.5% w/v yeast extract (Becton, Dickinson & Co., USA)
1% w/v NaCl (Univar, NSW)
pH 7.0
Autoclave
Luria-Bertani (LB) Agar
As for LB broth but with 1.5% w/v agar added
Minimal Medium (MM) – Liquid
30 g.L-1 sucrose (Univar, NSW)
(for flask culture)
2 g.L-1 NaNO3 (Chem-Supply, Gillman, South Australia)
1 g.L-1 K2HPO4 (Univar, NSW)
1 x trace stock solution (see below)
pH 6.0
Autoclave
Minimal Medium minus Carbon
As for liquid MM but without sucrose
(MM-C) - liquid
Minimal Medium (MM) - Solid
As for liquid MM but with the addition of
15 g.L-1 BBLTM agar (Becton, Dickinson & Co., USA)
50 μg.mL-1 phleomycin (Cayla, Toulouse) where required
200 μg.mL-1 hygromycin-B (Roche, Mannheim) where required
Top Agarose
10 g.L-1 BactoTM-Peptone (Becton, Dickinson & Co., USA)
5 g.L-1 NaCl (Univar, NSW)
6 g.L-1 agarose (Bio-Rad, Hercules, CA, USA)
Autoclave
100x Trace Stock Solution
50 g.L-1 KCl (Rowe Scientific, Australia)
(for Minimal Media)
50 g.L-1 MgSO4.7H2O (Chem-Supply, Gillman, South Australia)
1 g.L-1 ZnSO4.7H2O (BDH Laboratory Supplies, Poole, England)
1 g.L-1 FeSO4.7H2O (Univar, NSW)
0.25 g.L-1 CuSO4.5H2O (Univar, NSW)
Continued on the following page
44
Table 2.1: (Continued)
Medium
Ingredients
V8-Potato Dextrose Agar
150 mL L-1 V8 Juice (Campbell’s)
(V8-PDA)
10 g.L-1 Potato Dextrose Agar
3 g.L-1 CaCO3 (Chem-Supply, Gillman, South Australia)
15 g.L-1 Agar
pH 6.0
Autoclave
50 μg.mL-1 phleomycin (Cayla, Toulouse) where required
200 μg.mL-1 hygromycin-B (Roche, Mannheim) where required
45
2.5 Growth of Stagonospora nodorum
2.5.1 Routine Maintenance and Culture
Both wild type and transformant strains of S. nodorum were routinely grown
on solid media at 20 °C under alternating 12 h cycles of darkness and TL40W/05
(Philips, Holland) near-UV light. Liquid media were inoculated with 107-108 spores
into 100 mL of medium, shaken continuously at 140 rpm for 3 days on a Certomat® R
shaker (B. Braun, Melsungen, W. Germ.) at 20 °C in the dark. Strains of interest were
preserved by 250-350 mg (wet weight) of mycelium/spores being resuspended in 20%
glycerol, snap frozen in liquid nitrogen and stored at –80 °C.
2.5.2 Harvesting of Pycnidiospores
Pycnidiospores were harvested from sporulating plates, 2-3 weeks postinoculation. The plate was flooded with 5 mL sterile water, and the surface scraped
with a sterile pipette tip to remove aerial mycelium and ensure contact between the
water and pycnidia. The plate was left for 10 min to allow the water to swell and burst
pycnidia and release spores. A further 5 mL of sterile water was applied and the plate
re-scraped. The spore solution which was aspirated with a sterile syringe and filtered
through a sterile glass-wool-tipped 5 mL syringe. The solution was centrifuged for 5
min at 3500 g at room temperature and the pelleted spores resuspended in 1 mL sterile
water. Approximately 20 μL of spore solution, diluted to 1:10 or 1:100 if required,
were applied to a haemocytometer and spores counted under a light microscope to
determine the concentration of spores. A concentration of 1 x 107 spores.mL-1 was
sufficient for most procedures.
46
2.6 Growth of Escherichia coli
Escherichia coli cells were cultured overnight at 37 °C on solid or liquid LB
or liquid SOC media. Media were supplemented with ampicillin (100 μg.mL-1) as
required. Liquid media were shaken at 225 rpm on a Certomat® R shaker (B. Braun,
Melsungen, W. Germ.).
2.7 Nucleic Acid Extraction and Manipulation
2.7.1 Homogenisation of Fungal Mycelium/Pycnidiospores
Fungal mycelium was homogenised using a Retsch MM301 Mixer Mill
(Retsch GmbH, Haan, Germany) as per the manufacturer’s instructions.
Approximately 250-350 mg (wet weight) mycelia/pycnidiospores were harvested
from agar plates using a sterile scalpel blade. The mycelium was placed in a 2.0 mL
safety-capped tube, frozen in liquid nitrogen and stored at –80 °C until required.
Where pycnidiospores formed a significant portion of the tissue from which DNA was
to be extracted, the sample was lyophilised overnight in a Savant FDC206 freeze
drying chamber (Savant Scientific Instruments, Farmingdale, NY) attached to an Heto
Maxi-Dry Lyo freeze dryer (Heto-Holten, Allerød, Denmark). A single 3 mm
tungsten-carbide bead was placed in each tube prior to tissue lysis. Up to 5 tubes were
placed in the Adaptor Set (with balances as appropriate) in liquid nitrogen to render
the sample(s) metabolically inactive. Upon removal, the Adaptor Sets were knocked
against the bench to ensure the tungsten-carbide beads were mobile. The Adaptor Sets
were loaded into the Mixer Mill clamps and homogenised for 1 min at 30 Hz. This
tissue homogenisation was repeated up to a further 2 times with samples being re47
frozen in liquid nitrogen and Adaptor Sets being knocked against the bench to free the
beads each time before replacing in the Mixer Mill. Once the tissue resembled a fine
powder 300 µL of Buffer RLT was added to each tube followed by vortexing. Tubes
were centrifuged for 5 min at 6000 g at room temperature and lysate used
immediately for genomic DNA extraction.
2.7.2 Genomic DNA Extraction from Lysed Fungal Mycelium/Pycnidiospores
Genomic DNA was extracted from homogenised tissue using a Qiagen
BioSprint 15 (Thermo Electron Corporation, Finland) as per the manufacturer’s
instructions. The extracted DNA was stored at 4 °C until required.
2.7.3 Plasmid DNA Extraction
Plasmids were extracted using the Qiagen Midi-Prep Plasmid DNA Extraction
Kit (Qiagen Pty Ltd, Clifton Hill, Vic, Australia) as per the manufacturer’s
instructions. Briefly, an overnight 50 mL bacterial culture was centrifuged at 6000 g
for 15 min at 4 °C. The supernatant was discarded and the bacterial pellet resuspended
in 4 mL chilled Buffer P1 to which RNase A had been added. This was followed by
the addition of 4 mL Buffer P2. The tube was inverted gently 4-6 times to avoid
shearing genomic DNA and incubated for 5 min at room temperature. A further 4 mL
of chilled Buffer P3 was added, the tube inverted another 4-6 times and the entire
volume poured into a Qiagen Cartridge and incubated for 10 min at room temperature.
A Qiagen-Tip100 was equilibrated with 4 mL Buffer QBT after which it was removed
to a fresh vessel. The Qiagen Cartridge was decapped and placed into the Qiagen-Tip,
48
the plunger inserted in the cartridge and the lysate filtered into the Tip. The QiagenTip was then washed two times with 10 mL Buffer QC. DNA was then eluted into a
new tube with 5 mL Buffer QF and 3.5 mL room temperature propan-2-ol was added.
The volume was centrifuged at 5000 g for 60 min at 4 °C. The supernatant was
discarded and the pellet was washed with a 2 mL 70% ethanol at 5000 g for 60 min at
4 °C. The supernatant was discarded and the pellet air-dried for 5-10 min. The pellet
was then redissolved in 100 µL milliQ water.
2.7.4 Gel Electrophoresis of DNA
Agarose gels were prepared by dissolving 0.7% - 2.0% w/v Certified™
Molecular Biology Agarose (Bio-Rad, Hercules, CA) in 1x TAE Buffer (20 mM Tris,
10 mM glacial acetic acid, 1 mM EDTA). DNA gel electrophoresis was performed in
a horizontal DNA Sub Cell™, Wide Mini-Sub® Cell GT or Mini-Sub® Cell GT (BioRad, Hercules, CA) electrophoresis tank containing an agarose gel in 1x TAE Buffer.
DNA samples were mixed with 1x Blue/Orange Loading Dye (Promega, Madison,
WI) prior to loading into the gel. A lane containing 1 kb DNA Ladder (Promega,
Madison, WI) was loaded to enable the determination of the approximate molecular
weight of DNA bands. Electrophoresis tanks were attached to a Power Pac 300 (BioRad, Hercules, CA). Gels were electrophoresed until the dye front had progressed ¾
to 4/5 the length of the gel. The voltage used for electrophoresis varied with the size
of the gel, the sizes of the expected DNA bands, and the degree of
separation/definition of bands required. After completion of the run, gels were stained
with 0.5 μg.mL-1 ethidium bromide for 30 min and visualised under UV light in a
Bio-Rad Gel-Doc 1000 running Bio-Rad Molecular Analyst™ Version 1.4.
49
2.7.5 Determination of DNA Concentration
Prior to measurement, tubes containing extracted DNA were spun down in a
benchtop microcentrifuge at 6000 g for 1 min at room temperature to remove
interference in measurement by any MagAttract® Suspension G carried over from the
extraction process. DNA concentration was determined by a NanoDrop® ND-1000
Spectrophotometer (NanoDrop Technologies Inc., Wilmington, DE, USA) running
NanoDrop ND-1000 Version 3.1.2 (Coleman Technologies Inc., USA) as per the
manufacturer’s instructions. Briefly, 1-2 µL of DNA extract was pipetted onto the
lower measurement pedestal, the sampling arm lowered into position, and the DNA
concentration reading was taken. The upper and lower measurement pedestals were
cleaned prior to any measurement, and the NanoDrop was blanked with the same
solvent used to store the extracted DNA.
2.7.6 Restriction Endonuclease Digestion of DNA
Restriction digestion of DNA was performed with enzymes purchased from
Promega (Madison, WI) or Fermentas Life Sciences (Hanover, MD) as per the
manufacturer’s instructions. Each reaction contained 10x appropriate enzyme buffer,
10x BSA, excess enzyme(s) and DNA, with sterile water used to make up the final
reaction volume. Reactions were performed at the temperature specified for the
enzyme(s) used and incubation times ranged from 1.5 h to overnight. Reaction
volumes ranged from 15 μL (typically for analysis of plasmid minipreps) to 1 mL (for
linearisation of disruption constructs).
50
2.7.7 Purification of Linearised Plasmid DNA
Plasmid DNA was purified using a QIAquick PCR Purification Kit (QIAGEN
Pty Ltd, Clifton Hill, Vic, Australia) as per the manufacturer’s instructions. Briefly, 5
volumes of Buffer PB were added to 1 volume of PCR sample. The whole volume
was placed in a QIAquick spin column which was inserted into a 2 mL collection tube
and DNA bound by centrifugation at 17900 g for 30 sec at room temperature. The
eluent was discarded, 750 µL Wash Solution (Buffer PE) added and the tube
centrifuged at 17900 g for 30 sec at room temperature. The supernatant was discarded
and the tube again centrifuged at 17900 g for 60 sec at room temperature. The spin
column was removed to a clean 1.5 mL eppendorf tube and 10-30 µL STC buffer (1.2
M sorbitol, 10 mM CaCl2, 10 mM Tris-HCl pH 7.5) was added to the membrane and
the tube allowed to stand for 1 min. The tube was centrifuged at 17900 g for 60 sec at
room temperature and the purified DNA stored at -20 °C until required.
2.7.8 DNA Amplification by Polymerase Chain Reaction
PCR amplification of DNA was carried out in 25 μL reactions containing 2.5
μL 10x Buffer (Promega, Madison, WI), 1.0 μL 10 mM dNTPs (Promega, Madison,
WI), 0.5 μL 10 μM primer per primer used in the reaction, 0.2 μL Taq polymerase
(Promega, Madison, WI), a variable amount of DNA template and made up to 25 μL
with sterile water. Genomic DNA of transformants was routinely screened with actin
control primers actinF (5’-CTGCTTTGAGATCCACAT-3’) and actinR (5’GTCACCACTTTCAACTCC-3’) (Solomon et al., 2003) to confirm the presence of
DNA in DNA extractions. These primers amplified a band of approximately 300 bp.
51
PCRs were performed in a GeneAmp® PCR System 2400, 2700 or 9700 (Perkin
Elmer Applied Biosystems) thermocycler or a PTC-240 DNA Engine Tetrad® 2
Peltier Thermal Cycler (M J Research Inc., Waltham, Mass, USA; Bio-Rad
Laboratories Inc., Hercules, CA, USA) depending on the number of samples.
Thermocycler conditions consisted of an initial denaturing step of 2 min at 96 °C, 40
repeats of 10 sec at 96 °C, 20 sec at a variable annealing temperature, and 30 sec at 72
°C, a final extension step of 5 min at 72 °C, and a hold at 14 °C until ready for gel
electrophoresis.
2.8 Gas Chromatography – Mass Spectrometry
2.8.1 Extraction of Polar Metabolites
Biological material (fungal and/or plant) was harvested and snap frozen in
liquid nitrogen. Each sample was ground in liquid nitrogen in a pre-chilled sterile
mortar and pestle. Each ground sample was divided into four aliquots. Three aliquots
consisted of 10-100 mg sample and were placed into three pre-weighed eppendorf
tubes. The tubes were re-weighed to determine the weight of the sample in each tube.
These aliquots were later assayed for DNA concentration in order to normalise the
GC-MS sample data. The fourth aliquot for each sample consisted of 0.5-2.0 mg
sample and was transferred to a pre-weighed 2.0 mL eppendorf containing 1 mL
methanol and vortexed briefly to render the sample biologically inactive. This transfer
was performed as quickly as possible to minimise evaporation of methanol. The tube
was re-weighed to determine the weight of the sample. Each sample had 50 µL 0.2
mg.mL-1 ribitol (Sigma-Aldrich Inc., St Louis, MO, USA) added to act as an internal
52
standard, and 100 µL non-autoclaved milliQ water. Samples were inverted 5-6 times,
the tubes capped with safety caps, and incubated in an Eppedorf Thermomix
(Hamburg, Germany) shaking/heating block at 70 °C for 15 min with shaking at 1000
rpm. Tubes were centrifuged in an Eppendorf Centrifuge (Model 5417C, EppendorfNetheler-Hinz GmbH, Hamburg, Germany) at 14,000 g for 3 min and the supernatant
transferred to a fresh tube. The pellet was resuspended in 500 µL water plus 375 µL
chloroform, vortexed, and incubated at 37 °C for 5 min with shaking at 1000 rpm.
The tubes were centrifuged at 14,000 g for 3 min and the upper polar fraction
transferred to the fresh tube containing the supernatant from the first incubation.
Samples were then placed in a Savant SpeedVac® vacuum concentration chamber
(Savant Scientific Instruments, Farmingdale, NY) attached to a Savant RT400
Refrigerated Condensation Trap (Savant Scientific Instruments, Farmingdale, NY)
and evaporated to dryness overnight. Dried samples were stored at -80 °C for short
periods until required for derivatisation.
2.8.2 Derivatisation of Polar Metabolite Extracts
Methoximation of the carbonyl groups of the dried polar extracts was achieved
by the addition of 50 µL freshly prepared methoxylamine hydrochloride (Aldrich, St
Louis, MO, USA) (20 mg/mL-1 in pyridine (Univar, Seven Hills, NSW)) per sample,
followed by were incubation for 90 minutes at 30 °C with shaking at 1200 rpm.
Trimethylsilyl (TMS) esters were created by the addition of 80 µL N-methyl-N(trimethyl-silyl) trifluoroacetamide (MSTFA) (Sigma-Aldrich Inc., St Louis, MO,
USA) to the sample and incubation for 30 min at 37 °C with shaking at 1200 rpm.
53
Finally, 100 µL of the derivatised sample was transferred to an 11 mm crimp 2 mL
clear glass GC-MS vial containing a glass insert (Alltech, Deerfield, Il, USA), 10 µL
alkane mix added, and sealed with an 11 mm aluminium seal with rubber faced liner
(Alltech, Baulkham Hills, NSW). The alkane mix consisted of 62.5 µg.mL-1 each of
C10, C12, C15, C19, C22, C26, C32 and C36 dissolved in hexane. Samples were
stored at room temperature for at least 2 hours prior to GC-MS analysis. Where the
sample contained obvious suspended sediments, these were pelleted by centrifuging at
14,000 g for 1 min prior to loading the sample into the vial.
2.8.3 Gas Chromatography – Mass Spectrometry
Samples were injected as 1 µL derivatised metabolites in a 1:20
sample:methanol split ratio. The GC-MS system comprised an Agilent 7680
autosampler, Agilent 6890 gas chromatograph, and Agilent 5973N quadropole mass
spectrometer (all Agilent Technologies, Palo Alto, CA, USA). The system was
autotuned with perfluorotributylamine as per the manufacturer’s instructions. A 30-m
HP-50+ column with a 250 µm internal diameter and 0.25 µm film thickness was
used for gas chromatography (J&W Scientific, Folsom, CA, USA).
The operational temperatures were 230 °C (injection), 300 °C (interface) and
230 °C (ion source). The carrier gas used was helium with a flow rate retention time
locked to elute mannitol-TMS at 24.51 min. The GC oven temperature gradient
consisted of an initial temperature of 70 °C for 5 min, followed by ramping at 5
°C.min-1 to a final temperature of 300 °C held for 3 min.
54
Mass spectra and chromatograms were analysed using AnalyzerPro V2.2
(SpectralWorks Ltd, Runcorn, UK). Peak identification was based on comparison of
unknowns to the ACNFP GC-MS TMS Library, the NIST/EPA/NIH Mass Spectral
Library (NIST, Gaithersburg, MD, USA) or the Golm Metabolome Database (Kopka
et al., 2005) using NIST Mass Spectral Search Program 2.0 (NIST, Gaithersburg,
MD, USA).
2.8.4 Data Normalisation
All GC-MS samples contained an internal standard consisting of 50 µL 0.2
mg.mL-1 ribitol which was added during the polar metabolite extraction step. All
metabolite peak areas were divided by the peak area for ribitol, and then divided by
the wet weight of the sample to enable qualitative comparison between samples.
2.9 Software
Sequencing
and
homologous
recombinant
screening
primer
design,
CLUSTAL W alignments and Boxshading were performed using ANGIS Biomanager
(ANGIS, 2004). Restriction sites of nucleotide sequences were analysed using
NEBcutter V2.0 (Vincze et al., 2003). Sequence editing, manipulation and alignment
for base corrections and contig formation were performed using SEQtools version
8.2.067 (Rasmussen, 2004). Comparisons of nucleotide and amino acid sequences
against databases were performed with the BLAST suite of programs (Altschul et al.,
55
1997). Searches for protein motifs were performed using FingerPRINTScan (EBI,
2004a) and PPSearch (EBI, 2004b).
2.10 Statistical Analysis
Statistical analysis of data was performed with JMP IN V5.1 (SAS Institute
Inc, Cary, NC, USA) software using the Tukey-Kramer Honestly Significant
Difference (HSD) test with an alpha level of 0.05.
56
CHAPTER 3 - CONSTRUCTION AND
CHARACTERISATION OF A STRAIN OF
Stagonospora nodorum HARBOURING
DISRUPTED GENES FOR MANNITOL 2DEHYDROGENASE (mdh1) AND MANNITOL
1-PHOSPHATE 5-DEHYDROGENASE (mpd1).
57
3.1 INTRODUCTION
3.1.1 Nomenclature, Class and Structure of D-Mannitol
D-Mannitol belongs to the class of chemical compounds known as polyols or
polyhydric alcohols. These compounds are derived from precursor aldose or ketose
monosaccharides via the reduction of their carbonyl group to a hydroxyl group
(Brimacombe and Webber, 1972). D-Mannitol is a 6-carbon acyclic polyol which is
formed by the reduction of D-mannose or D-fructose, and derives its name from the
former of these two compounds. Since L-mannitol is not found in nature, the D-form
is commonly referred to as mannitol and this convention will be observed hereon. The
configuration of the hydroxyl groups on the carbon skeleton renders mannitol a
symmetric molecule (Figure 3.1).
3.1.2 Taxonomic Distribution
Mannitol, known previously as mannite or sugar of manna (Dunglison, 1856)
was first isolated from manna exuded from plants in 1806 by Proust (Ihde, 1984) and
was found to form a principal component of the basidiomycete Agaricus volvaceus in
1811 by Braconnot (Thomson, 1817; Gobley, 1856). One of the first systematic
reviews of the distribution of polyols concluded that mannitol was the most abundant
polyol in fungi and, with few exceptions, was present in all fungi studied (Lewis and
Smith, 1967). Subsequent reviews have shown this to be true for the fungi of the
phyla basidiomycota and ascomycota (with the exception of the yeasts and fission
yeasts of the subphyla saccharomycotina and taphrinomycotina), while mannitol was
58
Figure 3.1: The structure of D-mannitol (Fischer projection)
59
not found among the polyols of the phylum blastocladiomycota or the taxonomically
unclassifiable imperfect fungi assigned to the group fungi incertae sedis (Rast and
Pfyffer, 1989). Mannitol is reported as comprising up to 50% of the dry weight of the
fruiting bodies and 20% of the dry weight of mycelium of Agaricus bisporus (Rast,
1965) and 10-15% of the dry weight of the conidiospores of Aspergillus niger (Ruijter
et al., 2003). Mannitol is considered to be ubiquitous in lichens, found in the majority
of algae, and in the case of angiosperms it was the most widely distributed of the
polyols, being found in over 50 families (Lewis and Smith, 1967).
While only infrequently reported from animals, mannitol has been found to
occur naturally in insects (Wang et al., 2006) and humans (Servo et al., 1977a; Servo
et al., 1977b; Laker et al., 1982). Mannitol is also found in bacteria (Coyne and
Raistrick, 1932; Edwards et al., 1981; Wisselink et al., 2002) and apicomplexa
(Schmatz et al., 1989). It is considered to be the most abundantly occurring polyol in
nature (Wisselink et al., 2002). The near ubiquitous presence of mannitol as the major
soluble metabolite in phytopathogenic fungi has previously been recognised as
providing a potential antifungal target (Pfyffer et al., 1990).
3.1.3 Mannitol Metabolic Pathways in Stagonospora nodorum
There are a number of enzymes which have previously been reported as being
involved in mannitol metabolism in fungi (Table 1.3). Some of these have been
purported to occur as part of an enzymatic cycle (Figure 1.3). Two of the key
enzymes in this proposed cycle have been previously inactivated by targeted gene
deletion in S. nodorum. An EST library of S. nodorum genes expressed when grown
60
on wheat cell walls, produced a cDNA which encoded a putative mannitol 1phosphate dehydrogenase (Mpd1) (EC 1.1.1.17) (Solomon et al., 2005a). The gene
was disrupted by insertional mutagenesis using the knockout vector pGPSH-Mpd8
(Figure 3.2) and mutants lacked all detectable mannitol 1-phosphate dehydrogenase
activity in both directions. In vitro cultured mycelium contained approximately 20%
of the wild type levels of mannitol, and also contained less arabitol and more
trehalose than the wild type as shown by NMR (Solomon et al., 2005a). Similarly, the
conidia of the mpdA mutants in A. niger were found to contain 30% of the mannitol
content of wild type conidia (Ruijter et al., 2003). These results suggested that the
catabolic half of the mannitol cycle must be capable of operating in reverse in order to
synthesise mannitol. A fructose 6-phosphate phosphatase activity, which would be
required in order for this to happen, has previously been reported in fungi (Morton et
al., 1985b; Kulkarni, 1990; Chakraborty et al., 2004).
Using a reverse genetics approach, the gene encoding a putative mannitol
dehydrogenase gene (Mdh1) was cloned by degenerate PCR and insertionally
mutagenised using the knock-out vector pGPSP-Mdh1 (Figure 3.3) (Waters, 2004). It
was hypothesised, based on the mannitol cycle theory, that mutants in which this gene
was inactivated would be unable to catabolise mannitol and that this would elucidate a
clear role for mannitol in pathogenicity. The mdh1 mutants were found to lack all
mannitol dehydrogenase activity, yet were phenotypically identical to the wild type in
all other respects, including the ability to grow on mannitol as a sole carbon source,
and the ability to cause disease and sporulate in planta. The clear implication from the
mdh1 mutants was that the mannitol cycle could not exist as proposed, since
catabolism of mannitol would require the reversal of the dephosphorylation step from
61
Figure 3.2: Diagram outlining the construction of the knockout vector pGPSHMpd8 (taken from Solomon et al. (2005)). Note the transposon inserted at 354 bp
downstream of the predicted start codon, as determined by sequencing with primers
homologous to the transposon termini TnsL and TnsR.
62
TnsL
phleomycin
TnsR
335 bp
Mdh1
Genomic clone containing Mdh1
905 bp
335 bp
336 bp
phleomycin
pGPSP-Mdh1
Figure 3.3: Diagram outlining the construction of the knockout vector pGPSPMdh1. Note the transposon inserted at 335 bp downstream of the predicted start
codon, as determined by sequencing with primers homologous to the transposon
termini TnsL and TnsR.
63
mannitol 1-phosphate to mannitol, catalysed by mannitol-1-phosphate phosphatase.
This would be most simply achieved by the presence of a mannitol kinase enzyme.
However, apart from reports of a mannitol kinase in Microsporum gypseum (Leighton
et al., 1970) and Absidia glauca (Ueng et al., 1976; Ueng and McGuinness, 1977), no
such activity has previously been found in fungi (Lones and Peacock, 1964; Lee,
1967b; Strandberg, 1969; Adomako et al., 1972), including S. nodorum (P. Solomon,
pers. comm.).
The creation of a double transformant strain in which both the Mdh1 and
Mpd1 genes were disrupted was necessary for the further elucidation of the
metabolism of this compound. If these represented the only two mannitol metabolic
pathways, then it would be possible to abolish mannitol synthesis altogether, and also
produce a strain which would be unable to utilise mannitol as a sole carbon source.
Furthermore, it would be possible to finally demonstrate the role(s) of mannitol. In the
event that mannitol was still metabolised, there would be clear evidence for an
alternative pathway of mannitol biosynthesis. The first part of the current project was,
therefore, to create and characterise a double mutant strain which harboured both
inactivated gene constructs.
64
3.2 MATERIALS AND METHODS
3.2.1 Fungal Transformation
3.2.1.1 Preparation of Protoplasts
The S. nodorum strain mdh1-71, harbouring a disrupted mannitol 2dehydrogenase gene construct, pGPSP-Mdh1 (Figure 3.3 above), conferring
phleomycin resistance (Waters, 2004), was selected as the background for the double
mutant strain. Mycelium for preparation of protoplasts was obtained by inoculating
100 mL CzV8CS liquid medium with not less than 108 mdh1-71 pycnidiospores and
incubating for approximately 20 h in the dark at 20 ºC with shaking at 140 rpm. The
culture was centrifuged at 3000 g for 10 min at room temperature. The mycelium
pellets were resuspended in 50 mL 0.02 μm filter-sterilised wash solution (0.6 M
MgSO4) and washed at 3000 g for 5 min at room temperature. The pellet was
resuspended and well mixed in 25 mL 0.02 μm filter-sterilised Glucanex digestion
solution (1.2 M MgSO4, 10 mM NaH2PO4, 15 mg.mL-1 Glucanex (Novo Nordisk
Ferment Ltd., Dittingen, Switzerland), pH 5.8) and incubated for 2 h at 28 ºC without
agitation in a pre-warmed sterile glass petri dish to digest cell walls. The protoplasts
were transferred to a sterile 50 mL Falcon tube and the solution overlaid with 5 mL
Protoplast Overlay Solution (0.6 M sorbitol, 10 mM Tris-HCl pH 7.5) and centrifuged
at 3500 g for 15 min at room temperature. Up to 1 mL of protoplasts were removed
from the interface layer and transferred to a 2 mL eppendorf tube. An equal volume of
1 M Sorbitol Solution (1 M sorbitol, 10 mM Tris-HCl pH 7.5) was added, the
contents gently mixed by pipetting and centrifuged at 1500 g for 5 min at room
65
temperature. The pelleted protoplasts were resuspended in 2 mL STC buffer (1.2 M
sorbitol, 10 mM CaCl2, 10 mM Tris-HCl pH 7.5) and washed at 1500 g for 5 min at
room temperature. The pellet was resuspended in 0.5 mL STC buffer and kept on ice
until ready for transformation. A 1:500 dilution was prepared in STC buffer and the
protoplasts counted using an haemocytometer under an Olympus CX41RF (Olympus
Optical Co. Ltd., Philippines) light microscope. Ideally, a concentration of no less
than 5 x 108 protoplasts.mL-1 was required for transformation.
3.2.1.2 Transformation of Protoplasts
Transformation of protoplasts was essentially as described in Cooley et al.
(1988). Escherichia coli cells containing the pGPSH-Mpd8 (Figure 3.2) plasmid were
cultured overnight and plasmid DNA extracted. Sufficient DNA for two
transformations was linearised by digesting with the restriction endonuclease Apa1
(Promega, Madison, WI) for 3 h at 37 °C. The linearised DNA was purified using a
QIAquick PCR Purification Kit (QIAGEN, Clifton Hill, Vic, Australia) as described
above and resuspended in 10-30 μL STC buffer (1.2 M sorbitol, 10 mM CaCl2, 10
mM Tris-HCl pH 7.5). The concentration of DNA in the final solution was assayed
spectrophotometrically and 1 μL of the resuspended DNA was run on a 0.7% agarose
gel to ensure the plasmid was completely linearised and that only a single band was
present. Bands were visualised with a Bio-Rad Gel Doc 1000 using Perkin-Elmer™
UV Winlab Version 2.85.04. The linearised construct was stored at –20 ºC until
required.
66
Transformations were carried out in duplicate for each experiment. For each
transformation, approximately 108 protoplasts were transferred to a 2 mL eppendorf
tube, not less than 7.5 μg linearised DNA added, and the volume made up to 125 μL
with STC buffer if required. The contents were gently mixed with a P1000 pipette and
1 mL tip and incubated at room temperature for 15 min. PEG-mediated
transformation was conducted by adding 200 μL 60% PEG solution (60% PEG-4000,
10 mM CaCl2, 10 Tris pH 7.5) to the DNA with mixing by inversion and incubation
for 30 sec at room temperature. A further 200 μL 60% PEG solution was added,
mixed by inversion and incubated for 30 sec. Finally 800 μL 60% PEG solution was
added, mixed by inversion and incubated at room temperature for 15 min.
For each transformation experiment 5 mL 50 ºC CzV8-Proto Top Agar was
added to each of 4 pre-warmed (50 ºC) 10 mL sterile Falcon tubes, 310 μL
transformed protoplasts were added to each tube, the contents mixed and poured onto
a 15 mL CzV8-Proto Agar plate. For controls, 100 μL untransformed protoplasts were
added to each of an additional 2 pre-warmed (50 ºC) 10 mL sterile Falcon tubes
containing 5 mL 50 ºC CzV8-Proto Top Agar, mixed and poured onto 15 mL CzV8Proto Agar plates. All plates were wrapped in clingfilm, covered with aluminium foil
and incubated for 40 h at 20 ºC. After this incubation period 5 mL 50 ºC CzV8-Top
Agar containing 5.0 mg hygromycin per mL agar (i.e. 200 μg hygromycin per mL
agar on the plate) was overlaid on each transformed protoplast plate, and on one of the
untransformed protoplast plates. Plates were all rewrapped in clingfilm, covered in
foil and incubated at 20 ºC for 1-3 weeks, with transformant colonies being subcultured onto fresh CzV8CS + hygromycin plates as they appeared (3 to a plate). For
67
this particular transformation the CzV8-Proto Top Agar and CzV8-Proto Agar were
augmented with 1 mM mannitol.
3.2.1.3 Screening of Transformants
Approximately 2 weeks after sub-culturing, 200-250 mg (wet weight) of
mycelium were harvested from transformant colonies into a 1.5 mL eppendorf tube,
snap frozen in liquid nitrogen and stored at –80 ºC until DNA extraction was
performed as described (Section 2.7). Preliminary screening for homologous
recombinants was conducted by PCR amplification using the primers mpdkoF (5’GAGTTCACCATCGACCACT-3’) and mpdkoR (5’-TACTGCTTCTTCGCCTGG3’) predicted to amplify a band of 493 bp within a non-disrupted mannitol 1phosphate dehydrogenase gene. Presence of an amplified band indicated a putative
ectopic insertion and absence of an amplified band indicated a putative homologous
recombinant. PCR with actin primers was required in the latter instance to
demonstrate that there was genomic DNA in the sample. Putative homologous
recombinants were also screened with the primers mdhKOPcrF (5’-ACCGAGCTCA
AGGACCTCT-3’)
and
mdhKOPcrR
(5’-AACGAGGGAGCCAGTCTTG-3’)
predicted to amplify a band of 461 bp within a non-disrupted mannitol 2dehydrogenase gene. This was a precautionary step to confirm the stable integration
of the pGPSP-Mdh1 gene disruption construct. The annealing temperature for all
primer pairs used was 57 ºC.
68
3.2.1.4 Sub-Culturing of Transformant Colonies
Selected strains of putative ectopics and homologous recombinants identified
by PCR were subcultured in order to ensure a homogeneous culture for further
analysis. Approximately 100-150 mg (wet weight) mycelium/pycnidiospores were
harvested by scraping a sterile scalpel across the surface of the colony, transferred to a
1.5 mL eppendorf tube containing 1 mL sterile water and vortexed to resuspend the
fungal tissue. A 1:100 dilution was made with sterile water and 100 μL spread on a
fresh CzV8CS agar plate containing appropriate fungicides. Plates were wrapped in
clingfilm and incubated at 20 ºC under lighting conditions of alternating 12 h
darkness and 12 h TL40W/05 (Philips, Holland) near-UV light until individual
colonies could be seen. A discrete single colony was transferred to a fresh CzV8CS
agar plate containing appropriate fungicides, wrapped and incubated as previously.
The resulting colony was used as a source of inoculum for glycerol stocks from which
the strain was regenerated for experimental manipulation and Southern analysis to
confirm single integration of the disruption construct
3.2.2 Southern Hybridisation
3.2.2.1 PCR Amplification of DNA Probes
DNA probes for Southern analysis were amplified using the primer pairs
mdhSOUTHF
(5’-GTCGATGTCTTCATTGCCA-3’)
and
mdhSOUTHR
(5’-
GAAGTAGACGTAAGCGCCCT-3’), predicted to amplify a band of 393 bp, and
mpdSOUTHF (5’-AGTTCCTTCACAACTCTGGCT-3’) and mpdSOUTHR (5’-
69
GATGAAGCCCCTCCATGT-3’), predicted to amplify a band of 311 bp. PCR
amplification was carried out in 50 μL reactions containing 5.0 μL 10x Buffer
(Promega, Madison, WI), 1.0 μL 10 mM dNTPs (Promega, Madison, WI), 2.5 μL 10
μM primer per primer used in the reaction, 0.5 μL Taq polymerase (Promega,
Madison, WI), a variable amount of DNA template and made up to 50 μL with sterile
water. PCRs were performed in a GeneAmp® PCR System 2400, 2700 or 9700
(Perkin Elmer Applied Biosystems) thermocycler. Thermocycler conditions consisted
of an initial denaturing step of 2 min at 94 °C, 35 repeats of 20 sec denaturing at 94
°C, 20 sec annealing at 57 °C, and 40 sec extending at 72 °C, a final extension step of
5 min at 72 °C, and a hold at 14 °C until ready for gel electrophoresis.
3.2.2.2 DIG-Labelling of DNA Probes
This procedure was performed using the DIG High Prime DNA Labeling and
Detection Starter Kit II (Roche, Mannheim) as per the manufacturer’s instructions. A
volume containing 1 μg of PCR-amplified template DNA was made up to 16 μL with
sterile water, denatured by heating in a thermocycler at 99.9 °C for 10 min and
immediately chilled on ice. DIG-High Prime was thoroughly mixed by pipette and 4
μL was added to the denatured DNA and centrifuged briefly, followed by overnight
incubation in a thermocycler at 37 °C. The reaction was stopped by heating in a
thermocycler at 65 °C for 10 min and the DIG-labelled probe stored at –20 °C until
required.
70
3.2.2.3 Genomic DNA Digestion and Electrophoresis
Genomic DNA was extracted from the strains of interest (Section 2.7) and
digested in a 200 μL reaction containing 20 μg gDNA, 20 μL 10x buffer, 7 μL
restriction endonuclease and made up to the final volume with milliQ water. Strains
being analysed with the mdhSOUTH probe were digested with HindIII (Promega,
Madison, WI, USA) and strains being analysed with the mpdSOUTH probe were
digested with Pst1 (Promega, Madison, WI, USA). Digests were incubated overnight
at the manufacturer’s recommended temperature of 37 °C. The digested DNA was
concentrated in a Savant SpeedVac® vacuum concentration chamber (Savant
Scientific Instruments, Farmingdale, NY) attached to a Savant RT400 Refrigerated
Condensation
Trap
(Savant
Scientific
Instruments,
Farmingdale,
NY)
for
approximately 1 hour to produce a final volume of 25 μL. The concentrated DNA was
mixed with 1x Blue/Orange Loading Dye (Promega, Madison, WI, USA) and loaded
into the wells of a 250 mL 0.9% agarose gel and electrophoresed overnight at 30 V.
3.2.2.4 Southern Blot
Agarose gels for Southern blotting were electrophoresed until the dye front
had progressed 2/3 to 4/5 the length of the gel. Transfer of electrophoresed DNA from
the agarose gel to a membrane was performed on a VacuGene XL (Pharmacia
Biotech) vacuum tray attached to a VacuGene Pump (Pharmacia Biotech) using a
clean nylon mask which was approximately 5 mm smaller at each margin than the
agarose gel. An Hybond™ N+ positively charged nylon transfer membrane
(Amersham Pharmacia Biotech, Buckinghamshire, UK) was cut to the same
71
dimensions as the gel, and the apparatus assembled with the membrane beneath and
overlapping the nylon mask. The gel was placed such that it overlapped the mask and
the vacuum pump turned on and set to 50 mbar to ensure an intact seal. The gel was
serially covered with 50 mL Depurination Solution (0.2 M HCl), 50 mL Denaturing
Solution (1.5 M NaCl, 0.5 M NaOH) and 50 mL Neutralising Solution (1.5 M NaCl,
0.5 M Tris-HCl pH 8.0). Each solution was left in situ for 20 min following which it
was removed by pipette. Finally 20x SSC (3 M NaCl, 0.3 M sodium citrate, pH 7.0)
was used to flood the tray to a depth 2x the height of the gel. After 1 h the wells were
marked on the membrane with an HB pencil and the SSC was poured off. The top left
hand corner of the membrane was cut off and the membrane placed on Whatman
Chromatography Paper 3mm Chr. to dry. The membrane was then UV cross-linked in
a Gene-Linker (Bio-Rad, Hercules, CA) at 150 mJoule and stored at 4 °C until
required.
3.2.2.5 Hybridisation and Immunological Detection
Hybridisation and immunological detection was carried out using the DIG
High Prime DNA Labelling and Detection Starter Kit II (Roche, Mannheim) as per
the manufacturer’s instructions, in an Hybridization Oven/Shaker (Amersham
Pharmacia Biotech, Buckinghamshire, UK). Hybridisation membranes were prehybridised in an hybridisation tube with 20 mL DIG Easy Hyb and incubated for 30
min at 42 °C with constant rotation. The solution was replaced with the appropriate
denatured probe and hybridised overnight at 42 °C. Excess probe was removed by
stringency washes of 2 x 5 min in 100 mL 2x SSC, 0.1% SDS at 42 °C under constant
agitation, and 2 x 15 min in 100 mL 0.5 SSC, 0.1% SDS at 68 °C under constant
72
agitation. Immunological Detection commenced with washing the membrane for 1-5
min in Washing Buffer (0.1 M maleic acid, 0.15 M NaCl, pH 7.5 (20 °C), 0.3% v/v
Tween 20), 30 min incubation in 30 mL 1x Blocking Solution, and 30 min incubation
in 20 mL Antibody Solution. Membranes were then washed for 2 x 15 min in 100 mL
Washing Buffer, equilibrated for 5 min in 20 mL Detection Buffer, and placed DNAside up on an hybridisation bag. To the membrane was applied 1 mL CDP-Star, the
hybridisation bag was sealed and excess fluid expelled, and the membrane was
incubated for 5 min at room temperature. The membrane was exposed to Lumi-Film
Chemiluminescent Detection Film (Roche Diagnostics, Indianapolis) for 5 min to
overnight as required, and the film developed in a Fuji X-ray film processor FPM
3000.
3.2.3 In vitro Growth Assays
3.2.3.1 Growth on Solid Media
Selected strains were inoculated onto minimal medium, CZV8CS, and V8PDA agar plates (with phleomycin and/or hygromycin added as appropriate). The
inoculum consisted of 20 µL of a 106 spores.mL-1 spore suspension. Each strain was
prepared in triplicate and plates were wrapped in clingfilm and incubated at 20 °C
under a lighting regime of alternating 12 h darkness and 12 h TL40W/05 (Philips,
Holland) near-UV light. Measurements of colony diameter along a fixed line through
the centre of the colony were taken with a caliper at regular intervals and observations
made with regard to colony morphology.
73
3.2.3.2 Ability to Grow on Selected Carbon Sources
Selected strains were assayed for their ability to grow on selected carbon
substrates. All assays were performed in triplicate for each strain using a 96 well
microtitre plate with 180 μL medium and 20 μL of a 106 spores.mL-1 spore suspension
per well. Media consisted of liquid minimal medium as described (Table 2.1) but with
carbon being replaced with glucose, fructose, sucrose, trehalose or mannitol; or liquid
minimal media with no carbon or nitrogen, supplemented with either casamino acids
or casamino acids plus glucose. In all cases the final concentration of the compound
in the medium was 25 mM except for the casamino acids, where a final concentration
of 1 g.L-1 was used. A control treatment consisting of minimal medium with no
carbon source was also prepared. An initial reading of the plate at A595 was taken
using a Beckman Coulter® DTX 880 Multimode Detector (Wals, Austria) running
Multimode Detector Software Version 2.0.0.12 (Beckman Coulter Inc.). The plate
was covered and wrapped in clingfilm and incubated in the dark at 20 °C. After 1
week, a second reading at A595 was taken and fungal growth was measured by
deducting the first reading from the second.
3.2.3.3 Germination Assay
Glass slides were overlaid with 20 μL 1% agarose in milliQ water and covered
with a coverslip until dry and cool to produce a thin, even surface. The coverslips
were removed and the slides inoculated with 20 μL of a 106 spores.ml-1 spore
solution. Slides were placed in a humid chamber and incubated in the dark at 20 °C.
At 24 h incubation the slides were examined under a light microscope and the ratio of
74
germinated to ungerminated spores was calculated. Spores were considered to be
germinated if they had a visible germ tube.
3.2.4 Enzyme Assays
3.2.4.1 Preparation of Mycelium from Liquid Culture
Spores were harvested from each of the strains under investigation and 107
spores were inoculated into 100 mL minimal media in a 250 mL flask and incubated
for 3 days in the dark at 20 °C with shaking at 140 rpm. Cultures were harvested and
centrifuged for 10 min at 3000 g at 4 °C. Pellets were washed in 50 mM Tris-HCl pH
7.5 for 10 min at 3000 g at 4 °C. The supernatant was discarded and mycelium was
snap frozen by placing the tubes in liquid nitrogen. Samples were freeze dried
overnight in Savant FDC206 freeze-drying chamber (Savant Scientific Instruments,
Farmingdale, NY) attached to an Heto Maxi-Dry Lyo freeze dryer (Heto-Holten,
Allerød, Denmark) and tubes were replaced in liquid nitrogen and stored at -80 °C.
Each sample was ground in a sterile mortar and pestle containing 5-10 mL liquid
nitrogen. Once the liquid nitrogen had evaporated the tissue was further ground and
resuspended in 2 mL 50 mM Tris-HCl pH 7.5. The sample was transferred to a 2 mL
eppendorf tube, centrifuged at 20800 g for 45 min at 4 °C, and kept on ice thereafter.
The proteins were desalted using PD-10 Desalting Columns (GE Healthcare, Uppsala,
Sweden) as per the manufacturer’s instructions. Briefly, the PD10 columns were
equilibrated by eluting 25 mL 50 mM Tris-HCl pH 7.5 buffer and discarding the
flow-through. The centrifuged supernatant was made up to 2.5 mL with 50 mM TrisHCl pH 7.5 buffer and applied to the equilibrated column with the flow-through being
75
discarded. A further 3.5 mL buffer was applied to the column and the eluent collected
and kept on ice for enzyme assay.
3.2.4.2 Determination of Protein Concentration
The protein concentration in the samples was determined using a modified
bicinchoninic acid (BCA) method (Smith et al., 1985). Briefly, a fresh 50:1
BCA:CuSO4 working solution was made up. Each sample was assayed in triplicate
with 1 mL working solution and up to 50 μL desalted cell free extract. Where dilution
was required the sample was made up to 50 μL with milliQ water. A blank
comprising 50 μL sterile water was included. Bovine serum albumin (BSA) dilutions
of 0, 0.2, 0.4, 0.6, 0.8 and 1.0 mg BSA.mL-1 milliQ water were prepared in triplicate
to formulate a protein standard curve. The samples were incubated at 37 °C for 30
min and absorbance at A562 was measured on a Lambda 25 UV/VIS
Spectrophotometer (Perkin Elmer) running Perkin Elmer™ UV Winlab Version
2.85.04. Absorbance readings for blanks were subtracted from those of tests to
produce a net absorbance and the BSA protein standard curve used to determine
protein concentration in mg protein.mL-1 CFE.
3.2.4.3 Measurement of Relative Enzyme Activity
Samples were maintained on ice between all the following procedures.
Cuvettes for spectrophotometric assay were prepared in triplicate for each sample.
Each cuvette contained 50 μL desalted sample supernatant, unless otherwise stated,
and the reagents and volumes of each used for each enzyme assayed are as detailed
76
below. Each assay volume was made up to 1 mL with milliQ water. Enzyme
substrates were not added until the samples were equilibrated and in the case of
negative controls, water was added instead of the substrate. The oxidation of
NADH/NADPH or reduction of NAD+/NADP+ was measured at 340 nm to determine
activity in terms of U.mL-1 extract. One unit of activity was defined as the amount of
enzyme required to oxidise 1 μmol NADH/NADPH in 1 min, or to reduce 1 μmol
NAD+/NADP+ in 1 min at 30 ºC.
3.2.4.3.1 NADP+-dependent glucose 6-phosphate oxidation (glucose 6-phosphate
dehydrogenase)
This protocol was based on that of Langdon (1966). Each reaction volume
consisted of 500 μL 0.1 M Tris-HCl buffer pH 7.5, 45 μL 20 mM NADP+, 140 μL 0.1
M MgCl2, 50 μL CFE supernatant (milliQ water for control) and made up to 960 μL
with milliQ water. After equilibration 40 μL 50 mM glucose 6-phosphate was added
to start the reaction. Glucose 6-phosphate oxidation was measured by proxy using the
reduction of NADP+ as determined by change in absorbance at 340 nm.
3.2.4.3.2 NADPH-dependent fructose reduction (mannitol dehydrogenase)
This protocol was based on Noeldner et al. (1994). Each reaction volume
consisted of 500 μL 0.1 M Tris-HCl buffer pH 7.5, 10 μL 25 mM NADPH, 50 μL
CFE supernatant (milliQ water for control) and made up to 667 μL with milliQ water.
After equilibration 333 μL 2.4 M fructose was added to start the reaction. Fructose
77
reduction was measured by proxy using the oxidation of NADPH as determined by
change in absorbance at 340 nm.
3.2.4.3.3 NADP+-dependent mannitol oxidation (mannitol dehydrogenase)
This protocol was based on Trail and Xu (2002). Each reaction volume
consisted of 200 μL 0.5 M Tris-HCl buffer pH 9.0, 100 μL 20 mM NADP+, 50 μL
CFE supernatant (milliQ water for control) and made up to 500 μL with milliQ water.
After equilibration 500 μL 0.8 M mannitol was added to start the reaction. Mannitol
oxidation was measured by proxy using the reduction of NADP+ as determined by
change in absorbance at 340 nm.
3.2.4.3.4 NADH-dependent fructose reduction (NAD-mannitol dehydrogenase)
This protocol was based on the NADPH-dependent fructose reduction assay
above. Each reaction volume consisted of 500 μL 0.1 M Tris-HCl buffer pH 5.95, 10
μL 25 mM NADH, 50 μL CFE supernatant (milliQ water for control) and made up to
667 μL with milliQ water. After equilibration 333 μL 2.4 M fructose was added to
start the reaction. Fructose reduction was measured by proxy using the oxidation of
NADH as determined by change in absorbance at 340 nm.
3.2.4.3.5 NAD+-dependent mannitol oxidation (NAD-mannitol dehydrogenase)
This protocol was based on the NADP+-dependent mannitol oxidation assay
above. Each reaction volume consisted of 250 μL 0.2 M Tris-HCl buffer pH 9.0, 36
78
μL 10 mM NAD+, 50 μL CFE supernatant (milliQ water for control) and made up to
875 μL with milliQ water. After equilibration 125 μL 0.8 M mannitol was added to
start the reaction. Mannitol oxidation was measured by proxy using the reduction of
NAD+ as determined by change in absorbance at 340 nm.
3.2.4.3.6 NAD+-dependent sorbitol oxidation (sorbitol dehydrogenase)
This protocol was based on the NADP+-dependent mannitol oxidation assay
above. Each reaction volume consisted of 250 μL 0.2 M Tris-HCl buffer pH 9.0, 36
μL 10 mM NAD+, 50 μL CFE supernatant (milliQ water for control) and made up to
875 μL with milliQ water. After equilibration 125 μL 0.8 M sorbitol was added to
start the reaction. Sorbitol oxidation was measured by proxy using the reduction of
NAD+ as determined by change in absorbance at 340 nm.
3.2.4.3.7 NADH-dependent fructose 6-phosphate reduction (mannitol 1-phosphate
dehydrogenase)
This protocol was based on Solomon et al. (2005a). Each reaction volume
consisted of 500 μL 0.1 M Tris-HCl buffer pH 7.5, 10 μL 25 mM NADH, 50 μL CFE
supernatant (milliQ water for control) and made up to 900 μL with milliQ water.
After equilibration 100 μL 20 mM fructose 6-phosphate was added to start the
reaction. Fructose 6-phosphate reduction was measured by proxy using the oxidation
of NADH as determined by change in absorbance at 340 nm.
79
3.2.4.3.8 NAD+-dependent mannitol 1-phosphate oxidation (mannitol 1-phosphate
dehydrogenase)
This protocol was based on Kiser and Niehaus (1981). Each reaction volume
consisted of 500 μL 0.1 M Tris-HCl buffer pH 7.5, 25 μL 10 mM NAD+, 50 μL CFE
supernatant (milliQ water for control) and made up to 900 μL with milliQ water.
After equilibration 100 μL 10 mM mannitol 1-phosphate was added to start the
reaction. Mannitol 1-phosphate oxidation was measured by proxy using the reduction
of NAD+ as determined by change in absorbance at 340 nm.
3.2.4.4 Calculation of Specific Enzyme Activity
Specific enzyme activity was calculated by dividing the relative enzyme
activity (U.mL-1 extract) by the protein concentration (mg protein.mL-1 extract) to
produce specific activity in U.mg protein-1.
3.2.5 Stress Tolerance Assays
3.2.5.1 Osmotic Stress Assay
Selected strains were prepared as for the liquid growth assays above. Liquid
media consisted of minimal medium with the carbon replaced with glucose at a final
concentration of 25 mM and supplemented with 0, 0.25, 0.5, 0.75 and 1.0 M NaCl
(Merck Pty Ltd, Kilsyth, Victoria).
80
3.2.5.2 Oxidative Stress Assay
Selected strains were prepared as for the liquid growth assays above. Liquid
media consisted of minimal medium with the carbon replaced with glucose at a final
concentration of 25 mM and supplemented with 0 mM, 0.1 nM, 0.33 nM, 0.10 μM,
0.33 μM, 0.10 mM and 0.33 mM tert-butyl hydroperoxide (Sigma-Aldrich, St. Louis,
USA) as an oxidant.
3.2.6 Pathogenicity Assays
3.2.6.1 Detached Leaf Assay
Detached leaf assays were set up using a method modified from that described
in Benedikz et al. (1981). Leaves were harvested from wheat cv. Amery plants
approximately 2 weeks after potting. Harvesting was confined to the first true leaf.
The first 2 cm from the tip were cut off and the next 4 cm of blade were used for the
assay. Trimmed leaves were placed vein down onto a benzimidazole agar plate with
the two ends being embedded into the agar using a blunt forceps and sterile cotton tip.
(Benedikz et al., 1981b).
Leaves were prepared in triplicate for each treatment. Leaves were inoculated
in the centre of the blade with a 5 μL droplet of 106 spores.mL-1 resuspended in 0.02
% Tween 20. Once inoculated, plates were wrapped with clingfilm and incubated at
23 °C under an alternating light regime of 12 h Lumilux® Plus Eco (L 36W/21-840,
Osram, Germany) fluorescent lighting and 12 h dark. Leaves were monitored every 1-
81
2 days for chlorosis and appearance of lesions, and lesions were measured by a
caliper. Pycnidia were counted every 1-2 days.
3.2.6.2 Whole Plant Spray
Wheat cv. Amery seeds were surface sterilised and sown 8 seeds per pot, 8
pots per treatment, and grown for 2 weeks as described above (Section 2.4). On the
day of spraying, spores were harvested for each strain of interest and 13 mL of 106
spores.mL-1 in 0.02% v/v Tween 20 was made up for each strain (13 mL 0.02% v/v
Tween 20 for the negative control). The spore suspensions were kept on ice and 1 mL
of each was removed and stored at 4 °C for an assay of ability to grow on different
carbon sources as described above (Section 3.2.3.2). Pots were numbered and placed
randomly 8 pots to a tray, with the numbers for each treatment being noted. Each set
of 8 pots was sprayed evenly with the remaining 12 mL spore suspension in an
AirClean 600 Workstation (AirClean Systems, Australia) running at 270 Pa. The
spore suspension was applied with a Vivair 300 Mini Compressor (Vivaz) attached to
an SMC air brush sprayer at 140 kPa in two equal lots and allowed to dry between
applications. After the second application, the tray was filled with water to about 1 cm
deep, covered with a lid and sealed with Parafilm. The sealed trays were left for 2
days and then uncovered. An Ultrasonic Humidifier KT-100A (Ultrasonic, Taiwan)
was used to maintain a moist environment in the growth chamber for the following 5
days. All other growth conditions were as set out above (Section 2.4). On the 7th day
post-inoculation the plants were subjected to blind scoring with scores assigned as per
Figure 3.4.
82
9
8
7
6
5
4
3
2
1
Figure 3.4: Score chart for assigning disease scores to wheat cv. Amery seedlings
infected with strains of Stagonospora nodorum.
Source: ACNFP.
83
3.2.6.3 Latent Period Assay
On the day following the scoring for the whole plant spray, five leaves with
lesions were harvested per treatment for a latent period assay. Only first true leaves
were harvested, avoiding those which showed dead or yellowing tips, but which were
representative of the treatment. Harvested leaves were trimmed and embedded in
benzimidazole agar as described for detached leaf assay (Section 3.2.6.1 above).
Leaves were examined daily and scored when the number of pycnidia at stage 4 or 5
(Figure 3.5) exceeded 50.
3.2.6.4 Microscopic Examination of Host Penetration
A detached leaf assay was set up as above (Section 3.2.6.1) using SN15 and
the double mutant strain mpd1mdh1-107. Three leaves per strain were harvested at 1,
2, 4, 6 and 7 dpi. Leaves were cleared and stained with trypan blue using a method
modified from those of Bruzzese and Hasan (1983) and Shipton and Brown (1962).
Briefly, trypan staining solution was made up with 15 mL lactic acid, 15 g phenol,
28.8 mL 50% glycerol and 3.75 mL 0.4% trypan blue (all reagents from SigmaAldrich Inc., St. Louis, USA). Sampled leaves were boiled for 5 min in a 1:1 mixture
of trypan blue staining solution and 100% ethanol, and left at room temperature
overnight. Leaves were mounted and viewed under an Olympus BH-2 light
microscope (Olympus Optical Co. Ltd.) and images captured with an Olympus DP12
camera (Olympus Optical Co. Ltd., Japan).
84
All black
Swollen
Pink/black
Round exudate
Glistening pink
Burst bubble of
spores
Now both
different
shapes
Nipple like
Pink/black
Figure 3.5: Criteria for assigning developmental stages in Stagonospora nodorum
pycnidia on leaves of wheat cv. Amery.
Source: ACNFP.
85
3.2.7 Mannitol Supplementation Assays
3.2.7.1 In vitro Response to Mannitol Supplementation
3.2.7.1.1 In vitro sporulation response to altered mannitol concentration
The wild type strain SN15 and double mutant strain mpd1mdh1-107 were
selected for analysis. The background strain mdh1-71 and ectopic double transformant
Mpd1mdh1-101 were included as controls. Pycnidiospores were harvested for each
strain from CZV8CS agar cultures. Spore solutions of 106 spores.mL-1 were prepared
and 20 μL inoculated per plate onto minimal medium agar plates supplemented with
0, 1, 3, 10, 30 and 100 mM mannitol. Each treatment was prepared in triplicate. Plates
were incubated under the standard growth conditions and spores harvested and
counted after 20 dpi.
The strains SN15, mdh1-71, mpd1-1, mpd1mdh1-102, mpd1mdh1-107 and
Mpd1mdh1-101 were all grown on minimal media agar and minimal media agar
supplemented with 3 mM mannitol. The inoculum for the mpd1mdh1-107 strain was
sourced from minimal medium plates which had been serially sub-cultured for 1, 2,
and 3 generations to determine the effect of depleting residual mannitol from spores.
All treatments were prepared in triplicate. Plates were incubated under the standard
growth conditions and spores harvested and counted after 20 dpi.
86
3.2.7.1.2 Assay of mannitol content of spores
Pycnidiospores were harvested for SN15, mdh1-71, mpd1-1 and mpd1mpdh1107 cultures grown on minimal medium plates, and on minimal medium plates
supplemented with 3 mM mannitol. Spores were lyophilised overnight and
metabolites extracted, derivatised, and analysed by GC-MS as per the protocol above
for mannitol content.
3.2.7.2 In planta Response to Mannitol Supplementation
The wild type strain SN15 and the mutant strains mdh1-71, mpd1-1,
mpd1mdh1-102 and mpd1mdh1-107 were selected for analysis. Three detached leaf
assays were prepared as per the standard conditions with 4 replicate leaves per strain
and Tween 20-inoculated and uninoculated controls. Once the inoculation droplets
had all dried at 3 dpi, one of the DLAs had 5 μl drops of 3 mM mannitol added to
each inoculation site on a daily basis. A second DLA had 5 μl drops of milliQ water
added, and the third DLA had no addition. Lesion development was monitored and
measured and the number of pycnidia per leaf counted every 1-2 days.
87
3.3 RESULTS
3.3.1 Isolation of the mpd1mdh1 Double Mutant Strain
3.3.1.1 Transformation of Protoplasts
Two transformation experiments were conducted. The first experiment yielded
a total of 7 colonies and the second yielded 47 colonies. In both experiments the
negative control plates yielded zero colonies and the positive control plates exhibited
confluent growth.
3.3.1.2 PCR Screening
PCR screening was performed on strains including the wild type (SN15),
transformants harbouring either of the disruption constructs, and transformants
harbouring both disruption constructs. The results of PCR using the mdhkoF/R
primers and mpdkoF/R primers are shown (Figures 3.6 and 3.7 respectively). PCR
screening identified 2 colonies out of the 54 transformants as putative homologous
recombinants giving a recombination frequency of 3.7%. The putative homologous
recombinants were designated mpd1mdh1-102 and mpd1mdh1-107. A double
transformant created during a previous experiment and designated mpd1mdh1-51 (P.
Solomon, pers. comm.) was included in the analysis. For the purposes of control and
comparative analysis, an ectopic strain designated Mpd1mdh1-101 was retained, as
were the single mutant strains mdh1-71 (used as the genetic background for the
double mutant) and mpd1-1.
88
1 kb
Mdh1
actin
1
2
3
4
5
6
7
8
9
10
11
Figure 3.6: Duplex PCR amplification of gDNA from SN15 and mutant strains
transformed with pGPSP-Mdh1 or having this construct as their background. PCR
amplification was conducted using actinF/R primers (~300 bp) and mdhkoF/R
primers (~461 bp), with an annealing temperature of 57 °C. Lanes: 1: 1 kb MW
markers;
k
2 SN15;
2:
S 1 3:
3 mdh1-71;
dh1 1 4:
4 mdh1-73;
dh1 3 5: mdh1-79;
dh1 9 6:
6 mpd1mdh1-51;
d1 dh1 1 7:
mpd1mdh1-102; 8: mpd1mdh1-107; 9: Mpd1mdh1-101; 10: Mdh1-63; 11: negative
89
1 kb
Mpd1
actin
1
2
3
4
5
6
7
Figure 3.7: Duplex PCR amplification of gDNA from SN15 and mutant strains
transformed with pGPSH-Mpd8. PCR amplification was conducted using actinF/R
primers (~300 bp) and mpdkoF/R primers (~500 bp). Lanes: 1: 1 kb MW markers;
2: SN15; 3: mpd1-1; 4: mpd1mdh1-51; 5: mpd1mdh1-102; 6: mpd1mdh1-107; 7:
M d1 dh1 101
Mpd1mdh1-101.
90
3.3.1.3 Southern Hybridisation
A PCR of SN15 gDNA using the MdhSOUTHF/R and MpdSOUTHF/R
primers confirmed that the bands to be used as probes were of the predicted size
(Figure 3.8). Southern analysis of selected strains using a probe homologous to the
Mpd1 gene demonstrated that this is a single-copy gene and that the doubly
transformed strains mpd1mdh1-102 and mpd1mdh1-107 and the singly transformed
control strain mpd1-1 each contained a single insertion of the pGPSH-Mpd8
disruption construct (Figure 3.9A). The double transformant strain mpd1mdh1-51
appeared to have multiple copies of the pGPSH-Mpd8 disruption construct and was
discarded for further analysis. The hygromycin-/phleomycin-resistant ectopic mutant
Mpd1mdh1-101 showed the presence of the intact native gene.
Southern analysis of selected strains using a probe homologous to the Mdh1
gene demonstrated that this is a single-copy gene and that the background strain
mdh1-71, the singly transformed strains mdh1-73 and mdh1-79 and the doubly
transformed strains mpd1mdh1-102 and mpd1mdh1-107 each had a single insertion of
the pGPSP-Mdh1 disruption construct. The ectopic control strain Mdh1-63 showed
the presence of both the native and mutagenised versions of the gene (Figure 3.9B).
3.3.2 In vitro Phenotype
The strains SN15, mdh1-71, mpd1-1, mpd1mdh1-107 and Mpd1mdh1-101
were characterised on three different media as shown (Figure 3.10). The general
morphology of colonies could be divided into three zones. Firstly there was an inner
91
1 kb
mdhSOUTH
mpdSOUTH
dSOUTH
1
2
3
Figure 3.8: PCR amplification of gDNA from SN15 for use as a probe for Southern
analysis. PCR amplification was conducted using mdhSOUTHF/R primers (~393
bp) or mpdSOUTHF/R (~311 bp).
Lanes: 1: 1 kb MW markers; 2: mdhSOUTH
band; 3: mpdSOUTH band.
92
A
3 kb
2.5 kb
1
2
3
4
5
6
7
B
4.5 kb
4 kb
1
2
3
4
5
6
7
8
Figure 3.9:
A: Southern analysis of ApaI-digested gDNA transformed with the pGPSH-Mpd8
disruption construct, using probes homologous to Mpd1. Lane 1: MW markers; lane
2: SN15; lane 3: mpd1-1; lane 4: mpd1mdh1-51; lane 5: mpd1mdh1-102; lane 6:
mpd1mdh1-107; lane 7: Mpd1mdh1-101.
B: Southern analysis of HindIII-digested gDNA transformed with the pGPSP-Mdh1
disruption construct, using probes homologous to Mdh1. Lane 1: MW markers; lane
2: SN15; lane 3: mdh1-71; lane 4: mdh1-73; lane 5: mdh1-79; lane 6: mpd1mdh1102; lane 7: mpd1mdh1-107; lane 8: Mdh1-63.
93
Column 1
Column 2
Column 3
SN15
mdh1-71
dh1 71
mpd1 1
mpd1-1
mpd1mdh1-107
Mpd1mdh1-101
1 cm
Figure 3.10: Phenotypic characterisation of strains of Stagonospora nodorum grown
on three different media.
media Column 1: minimal medium (29 dpi); Column 2: V8
V8-PDA
PDA
(28 dpi); Column 3: CZV8CS (24 dpi).
94
zone consisting of the main body of the colony. This was often marked by dense
pycnidiation with no diurnal rings discernable and could be covered with dense aerial
mycelium. Next came an intermediate zone which tended to be covered in shorter
aerial mycelium (~0.16 cm) if any was present, but with occasional denser clumps.
This was the area in which diurnal rings of pycnidia occurred if they were present.
These were formed in response to the light-dark cycle under which the fungus was
grown. Finally, there was an outer zone or perimeter in which pycnidia were less
dense or absent, and which usually had greatly reduced aerial mycelium. Specific
differences between strains and media are as set out below.
3.3.2.1 Minimal Media Agar
When grown on minimal media agar plates SN15 produced colonies which
were covered in short aerial mycelium which varied in density over the surface of the
plate but formed no particular pattern. The colour of the aerial mycelium ranged from
white, where it was thickest, to a creamy colour where it was thinner, and greyish
where the thinner layer occurred over areas of dense pycnidiation. Pycnidia occurred
ubiquitously but were denser in the main body of the colony which extended to within
6 mm of the edge of the petri dish. Thereafter, pycnidia occurred as randomly
scattered single entities. Pycnidia located closer to the centre of the plate were oozing
cirrhus.
The mdh1-71 mutant was similar to the wild type although aerial mycelium
seemed to be more evenly distributed and of a more even density. While some white
clumps were evident, the general appearance was of a general tan/khaki colour
95
tending to more white/grey in the centre of the colony where pycnidiation was
heaviest. The perimeter of the colony appeared to be less even and darker than was
the case for SN15. Pycnidia towards the centre of the plate were observed to be
oozing cirrhus.
The most outstanding feature of the mpd1-1 mutant grown on minimal media
was the darkness of the agar beneath the colony. The density of the aerial mycelium
also decreased from the centre of the colony to the perimeter with a consequent
change in the colour of the mycelium from white at the centre to tan/khaki over the
intermediate portion and grey or translucent towards the perimeter. Pycnidia appeared
to be abundant but were not oozing as profusely as SN15 at the centre of the colony.
Moisture droplets were observed on the surface of the mycelium but it was not clear
whether these were the result of condensation or produced by the mycelium.
The mpd1mdh1-107 double mutant strain exhibited short white/beige aerial
mycelium of uniform density apart from the occasional dense white clump. Pycnidia
appeared abundant but rather than occurring ubiquitously in the main body of the
colony as the other strains, they occurred in clumps, giving the colony a ‘spotted’
appearance. The pycnidia closer to the centre of the petri dish were more dense and
were oozing cirrhus.
The ectopic strain Mpd1mdh1-101 was similar to the background strain mdh171.
96
3.3.2.2 CZV8CS Agar
On CzV8CS agar plates, SN15 produced a circular, symmetrical colony which
was basically flat on the plate. The perimeter of the colony consisted of a translucent
region of dense, fine, short hyphae extending into the medium, followed by an
opaque, non-sporulating zone of dense, pink-white mycelium. Behind this was a
short, sparsely sporulating zone of orange-brown mycelium with sparse aerial hyphae,
followed by the main body of the colony, consisting of dark brown diurnal rings of
heavy sporulation beneath pink-white aerial hyphae which varied between being
uniformly dense to being scattered and patchy.
The mdh1-71 mutant was virtually identical to SN15 although seemed more
prone to sectoring under these growth conditions, with areas of non-pycnidiation
sometimes occurring within the intermediate zone. The mpd1-1 mutant was also
similar to SN15 but usually had a thicker covering of aerial mycelium over most of
the colony.
The mpd1mdh1-107 double mutant had quite a variable phenotype on this
medium. This ranged from having dense pink-white mycelium around the central and
intermediate portion of the colony with less dense hyphae around the perimeter, to
having little aerial hyphae at all with the colony having bright pink colour, to having
patchy pink aerial hyphae and with parts of the colony having no aerial hyphae and
showing a ‘scalded skin’ look – bright red and wet looking. The strain also appeared
to be prone to sectoring on this medium and some plates exhibited combinations of
the above three phenotypes. The abundance and distribution of pycnidia was also very
97
variable with the first of the phenotypes above having normal looking pycnidia but
being reduced in abundance, occurring around the main body of the colony only. The
second phenotype produced more reduced numbers of pycnidia which tended to be
scattered in no particular pattern. The ‘scalded skin’ phenotype produced almost no
pycnidia.
The ectopic strain Mpd1mdh1-101 was similar to the background strain mdh171 except that while the amount of aerial mycelium was variable, it didn’t seem as
prone to sectoring.
3.3.2.3 V8-PDA
Almost no aerial mycelium was observed for SN15 when grown on this
medium. There was the occasional dense white clump, usually found towards the
centre of the colony, but this was atypical for this strain. Pycnidia were dense and
ubiquitous with formation of diurnal rings in the intermediate zone between the centre
of the colony and the perimeter. Occasionally aerial mycelium occurred above the
spaces between the pycnidial rings, but these were not profuse enough to render the
pycnidial rings indistinguishable.
The mdh1-71 mutant exhibited the same growth pattern as SN15 except that
the pycnidial rings were obvious closer to the centre of the colony, perhaps suggesting
that pycnidia were less dense than was the case for SN15. Formation of dense aerial
hyphae towards the centre of the colony was more common but not universal. The
mpd1-1 mutant was also very similar to SN15 with commonly more aerial hyphae
98
formation towards the centre, and possibly slightly darker in colour – but not to the
same extent as this strain growing on minimal media. Aerial hyphae in the outer part
of the colony were sometime seen to occur as lines radiating out from the intermediate
zone, but this was not universal.
The mdh1mpd1-107 double mutant often had a dense pink-white clump of
aerial mycelium at the very centre of the colony with short tan/khaki aerial hyphae
generally occurring over the intermediate region. Where this central clump was
absent, the centre consisted of clumps of pycnidia. In the outer 1/3 to ¼ of the colony
aerial hyphae occurred as small, discrete pink-white spots roughly arranged in rings.
Pycnidia were visible but did not appear to be as dense as the other three strains,
although they may have been partly obscured by the aerial hyphae.
The ectopic strain Mpd1mdh1-101 was similar to the background strain mdh171.
3.3.2.4 Mean Daily Growth Rates on Solid Medium
The strains SN15, mdh1-71, mpd1-1 and mpd1mdh1-107 were cultured on the
three solid media above in triplicate. Colony diameters were measured on a daily
basis from 3 to 14 days post inoculation when the majority of strains had grown to the
limit of the plate. The daily growth rate was calculated for each strain on each
medium and the Tukey-Kramer HSD test applied to compare the treatments (Figure
3.11). There was no statistically significant difference between the four strains on any
one medium. There was also no statistically significant difference between the growth
99
Mean
n Daily Growth Rate (cm
m/day)
0.7
ABC ABC ABC ABC
ABC BC C ABC
AB ABC A ABC
CZV8CS Agar
V8PDA
0.6
0.5
0.4
0.3
0.2
0.1
0
Minimal Medium Agar
Medium
SN15
mdh1-71
mpd1-1
mpd1mdh1-107
Figure 3.11: Mean daily growth rate (cm/day) (±SE) of strains of Stagonospora
nodorum on solid media. Statistical significance was calculated using the TukeyKramer HSD test, groups sharing a common letter were not significantly different.
The means were calculated from the daily growth rates of triplicate samples
measuredd daily
d il from
f
3 dpi
d i to
t 14 dpi.
d i N=33.
N 33
100
rates on minimal medium agar or the other two solid media. SN15 and mpd1-1 on V8PDA had a significantly greater daily growth rate than mpd1-1 on CZV8CS agar, and
mpd1-1 on V8-PDA also had a significantly greater daily growth rate than mdh1-71
on CZV8CS agar.
3.3.2.5 Ability to Grow on Selected Carbon Sources
The strains SN15, mdh1-71, mpd1-1 and mpd1mdh1-107 were investigated.
Growth on most media was equivalent for all strains with the exception that
mpd1mdh1-107 grew more poorly than the other strains on glucose and trehalose,
while it showed equivalent growth on fructose and was intermediate on sucrose
(Table 3.1). The three mutant strains grew poorly on mannitol compared to the wild
type, with mpd1mdh1-107 showing no growth on mannitol as a sole carbon source.
3.3.2.6 Germination Assay
The strains SN15, mdh1-71, mpd1-1 and mpd1mdh1-107 were investigated for
the ability of pycnidiospores to germinate. The wild type strain and the double mutant
were not significantly different from any other strains (Figure 3.12). The mpd1-1
strain had a significantly higher mean percentage germination rate than the mdh1-71
strain.
101
Table 3.1: Relative growth of Stagonospora nodorum strains SN15, mdh1-71, mpd11 and mpd1mdh1-107 on selected media in liquid culture. The base medium was
minimal medium with no carbon source (MM-C) or no carbon or nitrogen source
(MM-C-N) with supplements as indicated below.
Medium
SN15
mdh1-71
mpd1-1
mpd1mdh1-107
MM-C + 25 mM glucose
***
***
***
**
MM-C + 25 mM fructose
***
***
***
***
MM-C + 25 mM sucrose
***
***
***
**
MM-C + 25 mM trehalose
***
***
***
**
MM-C + 25 mM mannitol
***
**
*
-
***
***
***
***
MM-C-N + 1 g.L-1 casamino acids * * *
***
***
***
-
-
-
-1
MM-C-N + 1 g.L casamino acids
+ 25 mM glucose
MM-C
-
* * * = good growth
**
= medium growth
*
= poor growth
-
= no growth
102
Mean percentage of germinated spores
80.0
AB
B
SN15
mdh1-71
A
AB
70.0
60.0
50.0
40.0
30.0
20.0
10.0
0.0
mpd1-1
mpd1mdh1-107
Strain
Figure 3.12: Mean percentage of germinated spores (±SE) for selected strains of
Stagonospora nodorum at 24 hpi on 1% agarose. Statistical significance was
calculated using the Tukey-Kramer HSD test, groups sharing a common letter were
not significantly different. N=3.
103
3.3.3 Enzyme Assays.
Enzyme assays were performed on desalted cell-free extracts of the strains
SN15, mdh1-71, mpd1-1, mpd1mdh1-107 and Mpd1mdh1-101 (Table 3.2). Mannitol
dehydrogenase activity was present in all strains in both directions except those
strains in which the Mdh1 gene was inactivated. Mannitol 1-phosphate exhibited the
same behaviour except that fructose 6-phosphate reduction was not seen in the ectopic
strain (although mannitol 1-phosphate oxidation was seen in this strain). NADHdependent fructose reduction was observed in all strains and was an order of
magnitude less than the NADPH-dependent fructose reduction where both activities
were observed. There was, however, no NAD+-dependent mannitol oxidation detected
in any strain, while a low level of NAD+-dependent sorbitol oxidation was observed
in three of the strains. Glucose 6-phosphate dehydrogenase activity, used as a control
to demonstrate enzyme activity in the samples, was detected in all strains.
3.3.4 Stress Tolerance Assays
3.3.4.1 Osmotic Stress Assay
There was no significant difference observed in the response of the strains to
increasing concentrations of NaCl (Figure 3.13A).
104
Table 3.2: Specific enzyme activities for selected Stagonospora nodorum strains. All activities are shown as U/mg protein unless otherwise indicated.
Enzyme Activity
SN15
NADPH-dependent fructose reduction
1.44 ± 0.39
nd
1.24 ± 0.42
nd
nd
NADP+-dependent mannitol oxidation
11.8 ± 1.76
nd
3.07 ± 0.22
nd
nd
NADH-dependent fructose 6-phosphate reduction
0.21 ± 0.09
0.21 ± 0.09
nd
nd
nd†
NAD+-dependent mannitol 1-phosphate oxidation*
5.36 ± 1.06
6.59 ± 2.11
nd
nd
1.31 ± 2.18
NADH-dependent fructose reduction
0.11 ± 0.01
0.10 ± 0.01
0.17 ± 0.02
0.15 ± 0.01
0.81 ± 0.05
NAD+-dependent mannitol oxidation
nd
nd
nd
nd
nd
NAD+-dependent sorbitol oxidation
0.02 ± 0.0
0.01 ± 0.01
nd
0.08 ± 0.04
nd
NADP+-dependent glucose 6-phosphate oxidation
2.18 ± 0.21
0.64 ± 0.05
0.48 ± 0.06
1.27 ± 0.12
1.23 ± 0.20
mdh1-71
mpd1-1
mpd1mdh1-107
Mpd1mdh1-101
nd = not detected
* Activity expressed in mU/mg protein
†
While activity was not detected in this particular set of assays, activity equivalent to WT levels was seen in previous assays
A
B
Figure 3.13: Assays of the ability of strains of Stagonospora nodorum to grow
under conditions of osmotic stress (A) and oxidative stress (B). N=8.
106
3.3.4.2 Oxidative Stress Assay
There was no significant difference observed in the response of the strains to
increasing concentrations of tert-butyl hydro-peroxide (Figure 3.13B).
3.3.5 Pathogenicity Assays
3.3.5.1 Detached Leaf Assay
A detached leaf assay was set up using the strains SN15, mdh1-71, mpd1-1,
mpd1mdh1-102, mpd1mdh1-107 and Mpd1mdh1-101 with Tween 20-inoculated and
uninoculated controls (Figures 3.14 and 3.15). Using the Tukey-Kramer Test to
compare mean lesion formation for each day of measurement, the strains fell into four
groups for the duration of the experiment, with only three exceptions. The strains
SN15, mdh1-71 and Mpd1mdh1-101 exhibited the fastest rate of lesion formation, and
their lesions were indistinguishable in appearance. The mpd1-1 strain was
intermediate in its rate of lesion formation between the wild type and the double
mutants, and was significantly different to each. While it resembled a slowerprogressing version of the wild type lesion, it produced no pycnidia (Figure 3.16). The
double mutant strains both showed significantly reduced lesion formation, were
darker in appearance, and formed no pycnidia (Figure 3.16). There was no lesion
formation on either of the negative controls. The exceptions to the above groupings
occurred at 4 dpi, where the mdh1-71 strain was not significantly different from the
mpd1-1 strain; 6 dpi, where the mpd1mdh1-107 strain was not significantly different
107
2.5
Mean Lesion Length (cm)
2.0
A
1.5
AB
B
1.0
C
0.5
AB
BC
D
0.0
4
6
8
10
12
Days Post Inoculation
Figure 3.14: Mean lesion size (±SE) on detached wheat Amery leaves
inoculated with SN15 (♦), mdh1-71 (■), mpd1-1 (▲), mpd1mdh1-102 (●),
mpd1mdh1-107 (О) Mpd1mdh1-101 (X), Tween control (*) and uninoculated
control (). Statistical significance was calculated for each day of
measurement using the Tukey-Kramer HSD test, groups sharing a common
letter were not significantly different. The groups to which strains were
assigned at 12 dpi are shown using black letters. Apart from three exceptions
shown in red, all strains remained in the same groups for the duration of the
experiment. N=4.
108
SN15
mdh1-71
mpd1-1
mpd1mdh1-102
mpd1mdh1-107
Tween
Mpd1mdh1-101
Uninoculated
Figure 3.15: Detached leaf assay at 12 days post infection with strains of
Stagonospora nodorum as noted above. (Bar = 1 mm).
109
SN15
mpd1-1
mdh1-71
mpd1mdh1-107
Figure 3.16: Lesion formation on a detached leaf assay at 12 days post-inoculation
with selected strains of Stagonospora nodorum on wheat as noted above. Note the
absence of pycnidia in the mpd1 mutants. (Bar = 1 mm).
110
from the mpd1-1 strain, and 9 dpi, where the Mpd1mdh1-101 strain was not
significantly different from the mpd1-1 strain.
3.3.5.2 Whole Plant Spray
The results of a whole plant spray assay are presented (Figure 3.17). As
determined by the Tukey-Kramer test, there was no significant difference in the
ability of the mutants strains tested to cause disease from that of the wild type strain.
The only significant difference noted between the strains was that the double mutant
mdh1mpd1-102 caused significantly less disease than its parent background strain
mdh1-71 and the ectopic double transformant strain Mpd1mdh1-101. The Tween 20inoculated negative control exhibited a statistically lower mean disease score
compared to all infected treatments.
3.3.5.3 Latent Period Assay
The strains SN15, mdh1-71 and Mpd1mdh1-101 all scored with more than 50
pycnidia at stage 4/5, five days after the LPA was set-up i.e. 12 days after the whole
plant spray was inoculated. There were no pycnidia to be seen on the leaves
inoculated with the strains mpd1-1 or mpd1mdh1-107. There were 2 stage 4/5
pycnidia observed on one of the leaves inoculated with mpd1mdh1-102.
111
B
AB
C
mdh1mpd1-107
Tween
Mdh1mpd1-101
AB
mdh1mpd1-102
A
mpd1-1
A
mdh1-71
AB
10
Mean disease score
9
8
7
6
5
4
3
2
0
SN15
1
Strain
Figure 3.17: Mean disease scores (±SE) for wild type and selected mutant strains
of Stagonospora nodorum from a whole plant spray pathogenicity assay. Statistical
significance was calculated using the Tukey-Kramer HSD test, groups sharing a
common letter are not significantly different. N=8 (N=7 for the strain mpd1-1).
112
3.3.5.4 Microscopic Examination of Host Penetration
The double mutant strain mpd1mdh1-107 was compared with the wild type for
its ability to invade the host via stomata or direct penetration of the cuticle by hyphae
or hyphopodia. It was found to use all three methods of penetration (Figure 3.18).
3.3.6 Mannitol Supplementation Assays
3.3.6.1 In vitro Response to Mannitol Supplementation
3.3.6.1.1 In vitro sporulation response to altered mannitol concentration
The variability of the mpd1mdh1-107 double mutant to produce pycnidia and
spores was investigated by growing the strain on minimal media supplemented with 0,
1, 3, 10, 30 and 100 mM mannitol. Spore production was maximal at 3 mM mannitol
(Figure 3.19A) and pycnidia formation was observed to increase with increasing
concentration of mannitol up to 3 mM and to decrease thereafter (Figure 3.19B). The
response of SN15, mdh1-71 and Mpd1mdh101 to changes in mannitol concentration
were negligible by comparison.
When all strains under investigation were grown on minimal media agar with
and without 3 mM mannitol supplementation, it was found that there was no effect
upon strains with an intact mpd1 gene, but that all mpd1-disrupted strains had spore
counts increased to almost wild type levels by the addition of mannitol (Figure 3.20).
The double mutant strain mpd1mdh1-107 was subcultured for three generations onto
113
A
C
B
D
Figure 3.18: Trypan blue-stained lesions from detached leaves infected with
Stagonospora nodorum strains SN15 and mpd1mdh1-107. Arrows indicate
penetration attempts.
A: mdp1mdh1-107 hyphae entering stomata (2 dpi)
B: mdp1mdh1-107 hyphae penetrating host cuticle (2 dpi)
C: mdp1mdh1-107 differentiating hyphopodia (green arrow shows a hyphopodium
penetrating the host cuticle) (7 dpi)
D: SN15 (6 dpi) showing entry by stomate (red arrow), direct penetration by hyphae
(yellow arrow), and formation of hyphopodia (green arrows)
114
A
1.00E+09
Mean Spores/mL
1.00E+08
SN15
mdh1-71
1.00E+07
mpd1mdh1-107
Mpd1mdh1-101
1.00E+06
1.00E+05
0
5
10
15
20
25
30
Mannitol (mM)
B
0 mM
1 mM
3 mM
10 mM
30 mM
100 mM
1 cm
Figure 3.19:
A: The effect of mannitol supplementation upon sporulation of the strains SN15,
mdh1-71, mpd1mdh1-107 and Mpd1mdh1-101. Mean spores/mL (±SE) for strains
grown on minimal media agar supplemented with 0, 1, 3, 10 and 30 mM mannitol
are shown. N=3.
B: Pycnidia production by mpd1mdh1-107 in response to changes in mannitol
concentration in supplemented minimal media agar.
115
1.00E+09
1.00E+08
Mean Spores/mL
1.00E+07
1.00E+06
1.00E+05
1.00E+04
1.00E+03
1.00E+02
1.00E+01
1.00E+00
SN15
mdh1-71
mpd1-1
mpd1mdh1-102 mpd1mdh1-107- mpd1mdh1-107- mpd1mdh1-107- Mpd1mdh1-101
1
2
3
Strain
Figure 3.20: Comparison of mean spores/mL ( ±SE) for strains of Stagonospora
nodorum as shown. Blue columns result from growth on minimal media agar.
Purple columns result from growth on minimal media agar supplemented with 3
mM mannitol. The inoculum for the double mutant strain mpd1mdh1-107 came
from minimal medium agar plates on which the strain had been serially subcultured for 1, 2 and 3 generations as indicated by the suffix. N=3. Note that the y
axis has a logarithmic scale.
116
the two media treatments prior to inoculation and while zero spores were recorded for
some plates, sporulation was not completely abolished in all biological repeats for any
of these generations.
3.3.6.1.2 Assay of mannitol content of spores
GC-MS analysis demonstrated that spores of SN15 cultured on minimal
medium agar contained large amounts of mannitol, while the mpd1-disrupted mutants
grown on the same medium had only traces to undetectable amounts of mannitol in
their spores (Figure 3.21). When mpd1mdh1-107 was grown on minimal media
supplemented with 3 mM mannitol, large amounts of mannitol were detected. Due to
the small numbers of spores obtained from the unsupplemented cultures of the mpd1disrupted mutants, only qualitative comparisons could be made.
3.3.6.2 In Planta Response to Mannitol Supplementation
Addition of exogenous mannitol to lesions on leaves infected with SN15,
mdh1-71, mpd1-1 and mpd1mdh1-107 in a detached leaf assay resulted in the
formation of pycnidia on leaves infected by mpd1mdh1-107 and mpd1-1 by 12 dpi
(Figure 3.22). While these pycnidia were not as abundant as those formed on leaves
infected with SN15 or mdh1-71, and appeared to be smaller, it was the only condition
in which pycnidia were produced in a DLA by the mpd1-disrupted strains. To exclude
the possibility that there was cross-infection, pycnidia were picked off the leaves and
inoculated onto media containing appropriate antibiotics. In all cases there was
117
mannitol
SN15
mpd1-1
mpd1mdh1m107
mpd1mdh1m107
Figure 3.21: GC-MS chromatograms demonstrating the amount of mannitol present
in spores of SN15, mpd1-1 and mpd1mdh1-107 harvested from minimal medium
agar plates. The bottom chromatogram came from spores cultured on medium
supplemented with 3 mM mannitol. The retention time of mannitol was 24.51 in
these chromatograms.
118
Figure 3.22: Chemical complementation of the in planta sporulation defect of the
Stagonospora nodorum double mutant strain mpd1mdh1-107. Lesions were
inoculated with 5 μL 3 mM mannitol on a daily basis from 3 days post infection
(right). Mocks (left) were inoculated with 5 μL water. Top row: SN15; bottom row:
mpd1mdh1-107. Pictures were taken at 12 dpi. (Bar = 1 mm).
119
growth on the plates, thus confirming that the pycnidia were formed by the strain used
for the leaf infection.
3.4 DISCUSSION
3.4.1 Isolation of the mpd1mdh1 Double Mutant Strain
The transformation rate of 3.7% observed in this study was comparable with
the 3% homologous recombinant strains reported during the transformation of SN15
with the pGPSH-Mpd8 disruption construct (Solomon et al., 2006c). The recent
release of the S. nodorum genome sequence (Hane et al., 2007) has confirmed the
Southern analysis result that the two genes of interest were single-copy. The gene
encoding
mannitol
2-dehydrogenase
(Mdh1)
is
identified
by
the
record
SNOG_09898.2 and is found on scaffold 38. The gene encoding mannitol 1phosphate dehydrogenase (Mpd1) is identified by the record SNOG_12666.2 and is
found on scaffold 40.
3.4.2 Enzyme Assays
Cell-free extracts from all strains were positive for the constitutive enzyme
glucose 6-phosphate dehydrogenase, demonstrating that there was active protein.
Strains which had an intact Mdh1 gene (SN15 and mpd1-1) demonstrated mannitol
dehydrogenase activity in both the forward and reverse reaction. Strains which had an
intact Mpd1 gene (SN15, mdh1-71 and Mpd1mdh1-101) demonstrated mannitol 1phosphate dehydrogenase activity in both the forward and reverse directions, with the
120
exception of the ectopic strain. In this particular set of assays, the protein
concentration of the CFE for the ectopic strain was less than half that of the other
strains and the reduction of fructose 6-phosphate was not detected, although the
oxidation of mannitol 1-phosphate was detected. Fructose 6-phosphate reduction was
previously seen in this strain during the optimisation of the assay with an activity of
0.02 U/mg protein. Strains in which the Mdh1 and/or Mpd1 genes were inactivated
demonstrated no activity for the disrupted gene product.
A putative NAD-dependent mannitol dehydrogenase activity was assayed.
While activity was demonstrated in all strains using fructose as the substrate and
NADH as the co-factor, there was no activity in any strain using mannitol as the
substrate and NAD+ as the co-factor. There was activity in some strains, however,
using sorbitol as the substrate and NAD+ as the co-factor. Sorbitol does not appear as
a significant metabolite in S. nodorum in NMR or GC-MS analyses (Chapters 4 and 5,
this study). It is possible that there is an alternative endogenous substrate for the
oxidative reaction of this enzyme. Since this was not a focus of this study it was not
investigated further.
3.4.3 Mannitol Synthesis can Occur by Two Pathways
The previous inactivation of the Mpd1 gene in S. nodorum and A. niger
showed that mutant strains were still able to synthesis mannitol, although to only 2030% of wild type levels (Ruijter et al., 2003; Solomon et al., 2005a). The further
characterisation of the mdh1 mutant in S. nodorum showed that this strain is
unaffected in mannitol synthesis (Chapter 5, this study). The double mutant strain was
121
unable to synthesise mannitol. Furthermore, no strain was able to oxidise mannitol
using NAD+ as a co-factor. These data demonstrate that there was no additional
pathway of mannitol synthesis apart from the two under investigation. Secondly,
mannitol synthesis can be facilitated by either of the “halves” of the mannitol cycle.
This suggests that there is an as yet unidentified fructose 6-phosphate phosphatase
gene in S. nodorum. The reduced ability of the mpd1-1 mutant to accumulate mannitol
suggests that synthesis of mannitol occurred primarily via mannitol 1-phosphate in S.
nodorum. This is also consistent with the findings in A. niger (Ruijter et al., 2003). It
cannot be determined on the basis on these studies whether these pathways operate
simultaneously in vivo or whether they may be subject to regulation.
3.4.4 Mannitol Catabolism is Facilitated Primarily via Mannitol 1-Phosphate
The mpd1 inactivated strains were essentially unable to grow on mannitol as a
sole carbon source indicating that the catabolic step of the mannitol cycle is unable to
utilise mannitol in the absence of mannitol 1-phosphate dehydrogenase. It could be
claimed that this was perhaps due to some catabolism-inhibiting compound produced
when Mdh1 is used as the sole pathway for mannitol synthesis. The mdh1-71 strain,
however, was able to grown on mannitol as a sole carbon source, albeit at a reduced
rate compared to the wild type. This indicates that mannitol is catabolised primarily,
perhaps exclusively via mannitol 1-phosphate. This further suggests that mannitol
metabolism does not occur in an enzymatic cycle, as described above, in S. nodorum.
The implication is that the step from mannitol 1-phosphate to mannitol, catalysed by
mannitol 1-phosphate phosphatase must be reversible. The simplest reaction
achieving this would be facilitated by a mannitol kinase. While activity for such an
122
enzyme has been described in bacteria (Klungsøyr, 1966; Mehta et al., 1977), it has
not been convincingly demonstrated in fungi (Lones and Peacock, 1964; Lee, 1967b;
Strandberg, 1969; Adomako et al., 1972).
Recent independent confirmation that these enzymes do not operate in a cycle
in other fungi comes from studies in A. niger (Aguilar et al., 2008). Gene expression
studies with GFP and dTomato fusions to the promoters of MpdA and MtdA indicated
that while MpdA is expressed in mycelium, MtdA expression is restricted to conidia.
3.4.5 Mannitol is Required for Asexual Sporulation
Previous studies have shown that Mdh1 is dispensable for asexual sporulation,
with mutant strains producing wild type levels of pycnidia and spores both in vitro
and in vivo (Waters, 2004), while Mpd1 was required for sporulation in planta, but on
standard growth medium was able to sporulate normally in vitro (Solomon et al.,
2005a). Manipulation of the medium to exclude mannitol, and serial subculturing to
deplete exogenously accumulated mannitol from spores revealed that mpd1 strains
were compromised in their ability to sporulate in vitro. This was partially
compensated for in the mpd1 strain, which was previously shown to be able to
synthesise low levels of mannitol. However, sporulation in the non-mannitol
synthesising double mutant strain was abolished. This deficiency could be chemically
complemented, and addition of 3 mM mannitol to the growth medium restored
sporulation to wild type levels. The double mutant was shown to be unable to
sporulate in planta. Addition of exogenous mannitol to the leaf lesions was able to
partially complement this deficiency, with pycnidia being produced, although
123
sporulation did not reach wild type levels. The germination assay demonstrated that
spores produced by the various mutant strains were not significantly affected in
germination rates from the wild type. The difference noted between the strains where
mpd1-1 spores recorded a significantly higher germination rate in 24 h than mdh1-71
spores, did not explain the slower rate of lesion development of the former strain
compared to the latter.
There is a clear requirement for mannitol in asexual sporulation, and this is the
first conclusive evidence for a role for this compound in fungi. The mode of action
cannot be determined from this study, however. Mannitol has previously been
proposed as having a role in stress tolerance, since it has been shown to be a potent
quencher of reactive oxygen species (Smirnoff and Cumbes, 1989). Fungal mutants in
which mannitol production has been reduced have shown increased sensitivity to heat
and/or osmotic stress (Chaturvedi et al., 1996; Ruijter et al., 2003). The mpd1 mutants
did not show a significantly different response to oxidative or osmotic stress, but there
may be some other stress encountered during the infection process for which mannitol
is required. It is also possible that the presence of mannitol may be involved in a
sensory pathway involved in pycnidiogenesis, in a similar way to the proposed
extracellular sporulation factor(s) induced by FluG in A. nidulans (D'Souza et al.,
2001), although this would require further investigation.
124
3.5 CONCLUSION
3.5.1 Mannitol is Required for Pathogenicity
The major finding of this investigation is that the phytopathogen S. nodorum
has a requirement for mannitol in order to complete the process of infection on wheat.
The abolition of mannitol synthesis in a double mutant harbouring disruption
constructs for mannitol dehydrogenase and mannitol 1-phosphate dehydrogenase
resulted in an inability to sporulate in planta which could be partially rescued by the
addition of exogenous mannitol. This is the first demonstrated role for mannitol in a
fungal phytopathogen and suggests that if a means of inhibiting mannitol synthesis
could be devised, the polycyclic infection process could be arrested, thus significantly
reducing the impact of the disease in a wheat crop.
3.5.2 Enzymatic Cycling of Mannitol is Physiologically Unimportant
A major conclusion arising from this portion of the study was that the
mannitol cycle is unlikely to operate in S. nodorum as proposed by Hult and
Gatenbeck (1978). Instead, the evidence indicates that the metabolism of mannitol in
S. nodorum occurs by two separate pathways (Figure 3.23).
One of these pathways consists of the dephosphorylation of glycolytic/
gluconeogenic fructose 6-phosphate to fructose by fructose 6-phosphatase (and which
can be catalysed in the reverse direction by hexokinase), followed by the reversible
reduction of fructose to mannitol by mannitol 2-dehydrogenase. It is also possible that
125
glucose 6-phosphate
fructose 1,6-bisphosphate
fructose 6-phosphate
F6PP
MPD
HEX
mannitol 1-phosphate
M1PP
?
mannitol
fructose
MDH
mannitol
Figure 3.23: The two pathways for mannitol metabolism in Stagonospora nodorum
showing the enzymes involved in each step including a putative mannitol
phosphorylation step catalysed by unknown enzyme(s). The thicker arrows of the
mannitol 1-phosphate-mediated pathway indicate that this is the major pathway for
mannitol synthesis and catabolism. The glycolytic and gluconeogenic precursors of
fructose 6-phosphate are also shown. Abbreviations: MPD = mannitol 1-phosphate
dehydrogenase; M1PP = mannitol-1-phosphate phosphatase; Hex = hexokinase;
F6PP = fructose-6-phosphate phosphatase; MDH = mannitol dehydrogenase.
126
host fructose, resulting from the action of fungal/host invertase on host sucrose, could
also feed directly into this pathway. Disruption of this pathway produced no
phenotype. This indicated that while it may be possible for mannitol to be metabolised
in an enzymatic cycle in the wild type, there is no demonstrated physiological
requirement for such a cycle.
The other pathway consists of the reversible reduction of fructose 6-phosphate
to mannitol 1-phosphate by mannitol 1-phosphate dehydrogenase, followed by the dephosphorylation of mannitol 1-phosphate to mannitol by mannitol 1-phosphate
phosphatase. This final step is also reversible but the exact reaction pathway and
enzyme(s) involved are not yet determined.
Of these two demonstrated pathways, it is the mannitol 1-phosphate mediated
route which is the major catabolic pathway. It remains to be determined whether
mannitol is maintained as a single pool or not. Flux analysis would be required to
demonstrate whether the mannitol synthesised by either pathway is temporally or
spatially separated in the fungus as has been suggested for A. niger.
127
CHAPTER 4: METABOLOMICS ANALYSIS
OF HEALTHY AND DISEASED LEAVES
128
4.1 INTRODUCTION
4.1.1 The Metabolome and Antimicrobial Metabolites
The metabolome refers to the entire complement of low-molecular-weight
metabolites in a biological sample under a given set of conditions. The term was
simultaneously coined by Oliver et al. (1998) and Tweeddale et al. (1998) as a
concept complementary to the transcriptome and proteome (the complete set of genes
transcribed
or
proteins
synthesised
under
given
conditions
respectively).
Metabolomics refers to the analysis of the metabolome and encompasses contextsensitive changes in the metabolome which occur with developmental progress,
altered growth conditions, and resulting from genetic mutation. Typically the
metabolome pertains to a single organism, but investigations of heterogeneous
systems are informative, particularly when metabolites are specific to one of the
organisms involved. In the study of a pathosystem, metabolomics can be used to
identify changes in the metabolite profile which characterise the progress of an
infection. Detection of such changes could offer a means for improved understanding
of host-pathogen interactions. Provided the metabolites synthesised are sufficiently
abundant, it will be possible to detect them by use of an appropriate metabolomics
technology. The two main groups of methods currently employed for studies of this
nature are spectroscopic methods such as nuclear magnetic resonance (NMR)
spectroscopy, and chromatographic methods such as liquid/gas chromatography
coupled to mass spectrometry (LC/GC-MS) (Christensen and Nielsen, 1999). The
former of these is described more fully in Chapter 5 below and was not considered
suitable for use in this part of the study due to its lower sensitivity, requiring
129
millimolar concentrations of metabolites (Chatham et al., 2003). GC-MS has been
used to detect metabolites at nanomolar concentrations, and was the method used for
this investigation.
4.1.2 Overview of Technique
4.1.2.1 Gas Chromatography-Mass Spectrometry
Gas chromatography-mass spectrometry (GC-MS) is a popular platform for
metabolomics. This is due to well established techniques for sample preparation and
analysis, and the availability of two large online public domain metabolite databases,
the
NIST/EPA/NIH
Mass
Spectral
Library
(192,108
compounds)
(NIST,
Gaithersburg, MD, USA) and the Golm Metabolome Database (Kopka et al., 2005).
Other advantages include the ease of use, small sample size, high reproducibility and
robustness, and the relatively low costs compared to other spectroscopic and
chromatographic methods (Allwood et al., 2008; Garcia et al., 2008).
GC-MS requires analytes to be volatile and thermostable up to 250 ºC. A
necessary preparatory step is the derivatisation of the extracted metabolites. This
involves a methoximation step to confer thermal stability, and treatment with a
silylating compound such as TMS to form volatile trimethylsilyl esters (Halket et al.,
2005). The derivatised metabolites are separated by elution through a GC capillary
column with a polar stationary phase and using an inert carrier gas (helium in this
experiment) as the mobile phase. Retention time (RT) locking to a standard
compound (mannitol in the case of this experiment) enables the RTs of metabolites to
130
be precisely measured. The use of an internal standard not present in the samples
(ribitol in the case of this experiment), in conjunction with the sample weights,
enables the data to be normalised and permit valid comparisons of metabolite
abundances between samples (Solomon et al., 2006b). The mass spectrometer (MS)
consists of an ion source, mass analyser, and detection system. As samples are
introduced into the MS they are ionised by one of a variety of ion sources including
flame ionisation, chemical ionisation or electron ionisation. The latter method was
used here and involved bombardment of the gaseous sample molecules with a beam
of energetic electrons to generate ions. Mass analysers separate ions based on their
mass-to-charge (m/z) ratio, and range from the simple and inexpensive linear
quadropole (Q) and time-of-flight (TOF) systems, to hybrid Q-TOF and Orbitrap
systems, up to the more sensitive and expensive Fourier transform ion cyclotron
resonance (FT-ICR) analyser (Dunn, 2008). The first of these systems was used here.
A disadvantage of GC-MS is the requirement for derivatisation. Besides
requiring additional sample preparation time, there are a number of artefacts and
unexpected by-products which can be generated by the derivatisation step (Little,
1999). It is limited to the detection of low-molecular-weight, volatile compounds.
Other MS techniques such as liquid chromatography (LC)-MS are better suited for
detection of non-volatile and larger molecular weight metabolites.
4.1.2.2 Principal Components Analysis
Principal Components Analysis (PCA) is a multivariate analysis technique
devised by Pearson (1901), and independently developed by Hotelling (1933a;
131
1933b). The fundamental aim of the technique is to reduce the dimensionality of a
data set consisting of a large number of correlated factors (p), to a smaller number of
uncorrelated factors (m), whilst retaining as much of the variation present in the
original data set as possible. This is achieved by transforming the data set to a new set
of variables, the principal components (PCs), which are numbered in order of
decreasing ability to explain the total remaining variation. In a data set in which the
variation is non-random, the majority of the variation should ideally be accounted for
by the first few principal components. Where m is substantially smaller than p, the
PCs may be amenable to ready interpretation, although this is by no means necessarily
the case in all circumstances (Jolliffe, 1986).
4.1.3 Aims of the Study
The aim of this experiment was to use GC-MS, coupled with multivariate
analysis, to analyse the metabolites of infected and uninfected tissue of wheat leaves
inoculated with Stagonospora nodorum. This was designed to compare the two
conditions to reveal the presence of lesion-specific metabolites, in addition to any
metabolites resulting from a systemic reaction to infection. Mock-inoculated and
uninoculated controls were also included.
132
4.2 MATERIALS AND METHODS
4.2.1 Sample Collection
Benzimidazole agar plates were set up as per a detached leaf assay (Section
3.2.6.1), except that 8 cm sections of leaf were embedded in the agar, following the
trimming of 2 cm from the tips of the leaves. A separate plate was prepared for each
sample and consisted of 14 leaves per plate. Three biological replicates were prepared
for each harvesting time point. Each leaf was inoculated at 2.5 cm from one end of the
leaf with 5 μL of a SN15 spore solution consisting of 106 spores.mL-1 made up in
0.02% Tween 20 in milliQ water. Controls were prepared including leaves mockinoculated with 0.02% Tween 20 only, and uninoculated leaves. Plates were wrapped
in clingfilm and incubated under the standard culture conditions described above
(3.2.6.1). Samples were harvested at 0.5, 1, 3, 5 and 8 dpi. At each time point, the
portion of the leaves containing the lesion was excised and the 14 lesions per replicate
placed in a 1.5 mL eppendorf tube and snap frozen in liquid nitrogen. A second
portion from each leaf was harvested from the uninoculated, healthy part of the leaf,
with these 14 segments placed in a 1.5 mL eppendorf tube and snap frozen. To reduce
the risk of contamination between the diseased and non-diseased leaf portions, the
non-diseased portions were harvested before the inoculated/diseased portions.
Samples were stored at -80 ºC until ready for metabolite extraction. Samples were
defined as:
1. Infected – tissue taken from the site of inoculation with spore solution. In the
early stages this comprised a sample of leaf with the attached inoculation
droplet whilst for the later stages this comprised the entire lesion formed.
133
2. Uninfected – healthy tissue taken from a leaf inoculated with spore solution.
3. Mock-inoculated – tissue taken from the site of inoculation with Tween 20 (8
dpi only).
4. Uninoculated – tissue taken from an uninoculated leaf (8 dpi only).
4.2.2 Sample Preparation for GC-MS
Harvested tissue was homogenised in a Retsch MM301 Mixer Mill as
described above (Section 2.7.1). Wet weights for each sample were recorded
following addition of the homogenate to pre-weighed eppendorf tubes containing 1
mL methanol. Extraction of polar metabolites, derivatisation of samples, and GC-MS
spectra acquisition and analysis were performed as described above (Section 2.8).
4.2.3 Data Analysis
Normalisation of metabolite abundances in samples was performed using the
wet weight of the samples and the peak area of the ribitol internal standard in each
sample. The normalised data set was log-transformed following the addition of “1” to
each value in order to account for missing (i.e. zero) values. The transformed data was
analysed by Principal Component Analysis using The Unscrambler® v9.8 (Camo
Software AS, Oslo). The model was validated using the cross-validation method.
Outliers were identified via the Hotelling T2 ellipse 95% confidence limit. The top 20
variables (metabolites) contributing to the variation accounted for in each of the
principal components were subjected to statistical analysis as described previously
(Section 2.10).
134
4.3 RESULTS
4.3.1 GC-MS Peaks
A total of 194 peaks were detected across all samples by GC-MS (Table 4.1).
Of these, 99 were identified as a result of comparing fragmentation patterns with the
NIST and Golm libraries. These included the ribitol internal standard, and 67
endogenous
compounds,
some
with
multiple
derivatives,
and
including
methoxyaminated and dephosphorylated versions of some compounds. Two samples
(one 1 dpi infected and one 3 dpi uninfected replicate) had to be discarded due to
sample spillage in one case, and misinjection by the GC-MS automatic loader in the
other.
4.3.2 Principal Components Analysis
Analysis using the Hotelling T2 ellipse 95% confidence limit resulted in two
of the 8 dpi diseased leaf samples being identified as outliers. These were retained in
the analysis.
Principal components analysis showed that the first four principal components
cumulatively accounted for 76% of the total variance (PC1 = 43%, PC2 = 15%, PC3 =
10%, and PC4 = 8%). The best separation among samples in score plots was achieved
by combining PC1 with PC2 (Figure 4.1A).
135
Table 4.1: Library of retention times (RT) and identities for metabolites detected by
GC-MS from healthy and diseased tissue of wheat leaves infected with Stagonospora
nodorum and harvested at 0.5, 1, 3, 5 and 8 days post infection. Metabolites from
negative controls including mock-inoculated and uninoculated leaves are included.
RT
5.1447
5.9261
6.0850
6.3168
8.7803
9.1313
9.4491
10.1445
10.3167
11.1047
11.7073
12.1378
12.3299
12.3696
12.3762
12.8000
12.8663
12.9789
13.1444
13.1643
13.4756
13.6344
13.8663
13.9125
13.9920
14.0847
14.1642
14.4092
14.8065
15.2768
15.6077
16.0284
16.6211
16.8859
17.0713
17.4556
17.5349
17.7203
17.8197
Identity
Unknown
Unknown
Unknown
Unknown
Unknown
L-Valine_2TMS
Glycine_3TMS
Unknown
Glycerol_3TMS
Malonic_acid_2TMS
Glycine_3TMS
L-Serine_2TMS
L-Serine
Unknown
Unknown
Phosphoric_acid_3TMS
Unknown
L-Alanine_3TMS
Glyceric_acid_3TMS
Unknown
Unknown
L-Serine_3TMS
Unknown
Unknown
Nicotinic_acid_1TMS
L-Threonine_3TMS
Succinic_acid_2TMS
Unknown
Maleic_acid_2TMS
Unknown
Unknown
Threonic_acid-1,4-lactone_2TMS
Alanine_3TMS
Propanedioic_acid
Unknown
Unknown
L-Aspartic_acid_3TMS
4-Aminobutyric_acid_3TMS
Malic_acid_3TMS
Continued on following page
136
Table 4.1: Cont.
RT
18.0515
18.1178
18.2701
18.3031
18.3760
18.4686
18.6209
18.8527
19.2037
19.7136
19.9586
20.2831
20.5613
20.7136
20.8195
20.9586
21.3362
21.5347
21.7996
21.8261
21.8857
21.9453
22.1970
22.4156
22.5481
22.8327
22.8461
22.9189
23.0447
23.1638
23.3096
23.3361
23.6009
23.6803
23.8063
24.3094
24.3161
24.3427
24.3756
24.3823
24.4088
24.4154
24.4551
24.5346
24.6208
24.6405
Identity
Unknown
Unknown
Unknown
Unknown
Unknown
L-Aspartic_acid_3TMS
Unknown
Erythronic_acid_4TMS
Unknown
Xylitol_5TMS
Unknown
N-Acetylglutamic_acid_2TMS
Ribitol_5TMS
Arabitol_5TMS
L-Glutamic_acid_3TMS
Pyroglutamic_acid_2TMS
Unknown
Galactonic_acid_6TMS
Ribonic_acid_5TMS
L-Phenylalanine_2TMS
Unknown
Unknown
Unknown
Unknown
Unknown
2-keto-L-Gluconic_acid_5TMS
L-Asparagine_3TMS
Unknown
Unknown
Glucaric_acid_6TMS
Unknown
Ornithine_4TMS
Unknown
Unknown
Unknown
Unknown
Mannitol_6TMS
L-Glycerol-3-phosphate_4TMS
D-Quinic_acid_5TMS
Unknown
Unknown
3,1,11,15-Tetramethyl-2-hexadecen-1-ol
cis_Aconitic_acid_3TMS
Fructose_methoxyamine_5TMS
Mannitol_6TMS
Unknown
Continued on following page
137
Table 4.1: Cont.
RT
24.6538
24.6605
24.6671
24.6803
24.8657
24.9717
24.9849
25.0446
25.0843
25.3094
25.3690
25.4021
25.4950
25.5147
25.6140
25.8326
25.9650
26.0511
26.1835
26.2696
26.3226
26.3756
26.4220
26.5082
26.5744
26.6605
26.7663
26.9186
27.0908
27.1835
27.3160
27.5411
27.6008
27.6868
27.6935
27.7796
27.8127
28.0444
28.1571
28.2100
28.3556
28.7132
28.7861
28.8125
28.8656
29.0642
Identity
Fumaric_acid_2TMS
Phosphoric_acid_3TMS
Unknown
Unknown
Fructose_methoxyamine_5TMS
Unknown
Unknown
Glucose_methoxyamine_5TMS
Citric_acid_4TMS
Glucose_methoxyamine_5TMS
L-Lysine_4TMS
Unknown
Unknown
Isocitric_acid_4TMS
cis-Aconitic_acid_3TMS
Xylitol_5TMS
Glyceric_acid-3-phosphate_4TMS
Unknown
Unknown
Unknown
Unknown
L-Asparagine_4TMS
Unknown
Unknown
1-Ethylglucopyranoside_4TMS
Glucopyranose_5TMS
Glucose_5TMS
Unknown
Galactonic_acid_6TMS
Unknown
Glucaric_acid_6TMS
myo_Inositol_6TMS
L-Aspartic_acid_3TMS
Galactinol_9TMS
Unknown
L-Tyrosine_3TMS
Gulose_5TMS
Unknown
β-1-Galactopyranoside
Unknown
Galactose_methoxyamine_5TMS
Glucaric_acid_6TMS
L-Aspartic_acid_3TMS
Sedoheptulose_6TMS
Hexadecanoic_acid_1TMS
1-Methyl-β-D-galactopyranoside_4TMS
Continued on following page
138
Table 4.1: Cont.
RT
29.3961
29.3027
29.4086
29.9053
30.5344
30.5411
30.6801
30.7794
31.0112
31.3489
31.7793
32.0641
32.1701
32.2893
32.4548
32.4614
32.8986
33.0707
33.4415
34.0243
34.2429
34.4350
34.6070
35.2760
35.5542
35.5739
35.8387
36.1568
36.5342
36.7262
37.0043
37.0838
37.2228
37.3751
38.4479
40.1300
40.6599
40.6796
41.0704
41.1963
41.3418
41.7723
41.7791
41.9115
42.0438
42.6928
Identity
Unknown
Unknown
Unknown
Unknown
2-O-Glycerol-β-D-galactopyranoside_6TMS
Unknown
Unknown
2-O-Glycerol-β-D-galactopyranoside_6TMS
1-Methyl-β-D-galactopyranoside_4TMS
Unknown
1-Methyl-β-D-galactopyranoside_4TMS
Unknown
Unknown
Octadecanoic_acid_1TMS
Unknown
Ribitol_5TMS
Unknown
α-Linolenic_acid
Unknown
Glucose-6-phosphate_methoxyamine_6TMS
L-Tryptophan_3TMS
myo-Inositol-2-phosphate_7TMS
D-Glucuronic_acid_5TMS
Glucopyranose_5TMS
Unknown
Unknown
Unknown
Unknown
Unknown
Maltose_methoxyamine_8TMS
Sucrose_8TMS
Unknown
Unknown
Galactinol_9TMS
Trehalose_8TMS
Unknown
myo_Inositol_6TMS
Unknown
Unknown
Unknown
Galactinol_9TMS
Unknown
Unknown
Unknown
Unknown
Unknown
Continued on following page
139
Table 4.1: Cont.
RT
42.8451
43.2557
43.6795
44.3022
44.3352
44.5537
46.1497
47.6795
48.0767
48.6793
51.3017
52.0368
53.8777
53.9308
54.5996
54.6593
54.6726
Identity
Unknown
Unknown
Melibiose_8TMS
Octacosanol_1TMS
Lanost-8-ene-3-β,7-α-diol,3-acetate
D-Glucuronic_acid_5TMS
Sucrose_8TMS
Unknown
Unknown
β-Sitosterol_1TMS
Unknown
Gulose_5TMS
Sucrose_8TMS
Unknown
Unknown
Unknown
Sucrose_8TMS
140
PC2 (15%)
A
PC1 (43%)
B
Figure 4.1: Principal components analysis (PCA) score plot (A) and loading plot
(B) for PC1 versus PC2 from a PCA of polar metabolites processed by GC-MS.
Samples consisted of tissue harvested at 0.5, 1, 3, 5 and 8 days post inoculation
(dpi) from a detached leaf assay. Tissue was harvested from diseased (inf) and
healthy (noninf) tissue of leaves inoculated with Stagonospora nodorum spores.
Control leaves were either mock inoculated (mock) or uninoculated (uninoc) (both
8 dpi only).
141
Considering
the
principal
components
individually,
PC1
exhibited
differentiation between the samples both in terms of time to sample harvest, and
infection. While the separation in the uninfected samples was not large, there was
nevertheless an observable continuum with the early stage samples at one end and the
late stage samples at the other. All diseased tissue samples from 3 dpi and later had
positive scores with respect to PC1 and with a positive trend between the score and
time to harvest. All remaining samples had negative scores with respect to PC1 with
the exception of one 5 dpi uninfected sample and three 8 dpi samples (one uninfected,
one mock inoculated and one uninoculated). Three of these four had low positive
scores, while the 8 dpi uninfected sample had a score which placed it among the 3 dpi
infected samples. There was a trend observed whereby score increased with time to
harvest. Loadings for each of the top 20 variables (metabolites) contributing to PC1
are shown (Figure 4.2A).
The samples were clustered into three groups by PC2. The early stage
uninfected samples up to and including 1 dpi formed one cluster all with positive
scores, while the remaining uninfected, mock-inoculated and uninoculated samples
formed a second cluster all with negative scores. Apart from a couple of interpolated
samples, all of the infected samples formed a discrete cluster between these two.
Loadings for each of the top 20 metabolites contributing to PC2 are shown (Figure
4.2B).
The PCA loading plot for PC1 versus PC2 (Figure 4.1B) gave emphasis to
some of the factors separating the samples. Mannitol and trehalose were key
metabolites defining the infected samples particularly in the later stages of infection,
142
Metabolite
A:
Mannitol (6TMS)
Citric acid (4TMS)
Sucrose (8TMS)
Malic acid (3TMS)
Trehalose (8TMS)
Unknown – 26.3226
Glucose MeOx (5TMS)
Sucrose (8TMS)
Arabitol (5TMS)
cis-Aconitic acid (3TMS)
Unknown – 30.5411
Glucose MeOx (5TMS)
Unknown – 29.4086
Pyroglutamic acid (2TMS)
Galactinol (9TMS)
Melibiose (8TMS)
L-Serine
L-Glutamatic acid (3TMS)
L-Aspartic acid (3TMS)
Unknown – 10.1445
-0.3
-0.2
-0.1
0.0
0.1
0.2
0.3
0.4
0.2
0.3
0.5
Factor Loading
B:
Metabolite
Glucose MeOx (5TMS
Unknown – 24.3094
Citric acid (4TMS)
Mannitol (6TMS)
Glucose MeOx (5TMS)
Isocitric acid (4TMS)
cis-Aconitic acid (3TMS)
Malic acid (3TMS)
Unknown 37.3751
Glucose (5TMS)
Unknown – 27.1835
Erythronic acid (4TMS)
D-Glucuronic acid (5TMS)
Unknown – 13.1643
Fructose MeOx (5TMS)
2-O-Glycerol-β-D-galactopyranoside (6TMS)
myo-Inositol (6TMS)
D-Quinic acid (5TMS)
Trehalose (8TMS)
Unknown – 28.0444
-0.4
-0.3
-0.2
-0.1
0.0
0.1
0.4
Factor Loading
Figure 4.2: The top 20 variables (metabolites) contributing to the variation
accounted for by PC1 (A) and PC2 (B) in a PCA of healthy and Stagonospora
nodorum-infected wheat leaf tissue. Metabolites are arranged on the Y-axis in order
of magnitude of factor loading (regardless of sign). Unidentified metabolites are
shown with their retention time. Note that the scales on the X-axis are not identical.
143
while sucrose and glucose were defining features of the uninfected/mock
inoculated/uninoculated and early stage infection samples. The early stage samples of
all treatments were associated with cis-aconitate. They also had high levels of
glucose, although other derivatives of glucose were also found in later samples. The
later stage uninfected/mock inoculated/uninoculated samples were also characterised
by isocitrate, and the unknown compound with a RT of 24.3094. Malate and citrate
were both negatively correlated with the early samples and positively correlated with
the later samples, particularly where infection was present. Out of the top 20
metabolites for each principal components, seven metabolites contributed to both PC1
and PC2 (malate, mannitol, cis-aconitic acid, glucose methoxyamine (both
derivatives), citrate, and trehalose).
4.3.3 Statistical Analysis of Metabolites Identified by PCA
There were 33 unique GC-MS peaks in the top 40 variables contributing to
PC1 and PC2 as determined by their factor loadings. When the normalised GC-MS
data was analysed for each of these, 9 exhibited no statistically significant differences
between samples. The remaining 24 exhibited a number of expression profiles.
4.3.3.1 Metabolites Present Only in Diseased Samples
There were three metabolites present only in infected tissue (Figure 4.3). The
presence of these metabolites was only statistically significant at 8 dpi, although
arabitol and mannitol were present at earlier stages of infection and displayed a trend
whereby they increased with time of infection. Trehalose was only present at 8 dpi.
144
Arabitol
Normalised abundance
40.00
B
B
B
0.5
1
3
B
A
B
B
B
B
B
8
0.5
1
3
5
8
B
B
30.00
20.00
10.00
0.00
5
diseased
mock uninoc
healthy
Treatment
Mannitol
Normalised abundance
900.00
B
B
B
1
3
B
A
B
B
B
B
B
1
3
5
8
B
B
750.00
600.00
450.00
300.00
150.00
0.00
0.5
5
8
0.5
diseased
mock uninoc
healthy
Treatment
Trehalose
Normalised abundance
160.00
B
B
B
B
A
0.5
1
3
5
8
B
B
B
B
B
1
3
5
8
B
B
140.00
120.00
100.00
80.00
60.00
40.00
20.00
0.00
0.5
diseased
mock uninoc
healthy
Treatment
Figure 4.3: Mean normalised abundance (±SE) for metabolites present only in
diseased tissue. Treatments consisted of diseased and healthy tissue sampled at 0, 1,
3, 5 and 8 dpi from wheat leaves inoculated with Stagonospora nodorum. Mockinoculated and uninoculated controls were harvested at 8 days. GC-MS peak areas
were normalised by division using the peak area of an internal standard (ribitol)
followed by division by the sample wet weight. Statistical significance was
calculated for each group using the Tukey-Kramer HSD test, groups sharing a
common letter were not significantly different. N=3 for all treatments except 1 dpi
infected and 3 dpi uninfected (N=2).
145
4.3.3.2 Metabolites Increasing with Time of Infection in Diseased Samples
Six metabolites were present in all treatments, but were significantly higher in
later stage diseased samples (Figure 4.4). None of these metabolites exhibited any
significant difference in abundance within the healthy tissue samples with the
exception of glucose. In this case the 8 dpi uninfected samples had significantly more
glucose than the 0.5 dpi samples.
4.3.3.3 Metabolites Significantly Higher in Healthy Tissue than Diseased Tissue
Six metabolites were significantly more abundant in some healthy samples
than in any diseased samples (Figure 4.5). In the case of the two glucose
methoxyamine derivatives, sucrose, and an unknown (RT 28.0444) the compound
was highest in the early stage uninfected (0.5 to 1 dpi) samples. In the case of Dglucuronic acid and isocitric acid, the compound was highest in the late stage (3 to 8
dpi) samples.
4.3.3.4 Metabolites Significantly Lower in Late Stage Diseased Tissue
Nine metabolites were present in both diseased and healthy tissue, but were
significantly lower or absent in 5 dpi and/or 8 dpi infected samples (Figures 4.6 and
4.7). All were compounds with low abundances. Six were compounds with no match
in the MS databases. One unknown with an RT of 24.3094 was not identified in any
diseased sample.
146
L-Aspartic acid
Citric acid
250.00
AB AB AB AB
A AB AB AB AB AB AB
B
10.00
8.00
6.00
4.00
2.00
Normalised abundance
Normalised abundance
12.00
B
B
B
1
3
1
3
5
8
0.5
1
diseased
B
B
B
1
3
5
8
B
B
150.00
100.00
50.00
3
5
8
mock uninoc
0.5
healthy
0.5
mock uninoc
healthy
L-Glutamic acid
Glucose
50.00
C BC AB AB AB AB AB
C C
C C
Normalised abundance
AB A
C
8
Treatment
10.00
BC AB C
C
5
diseased
Treatment
Normalised abundance
B
0.00
0.5
AB AB AB AB A
B
AB AB AB AB AB B
40.00
30.00
6.00
20.00
4.00
10.00
2.00
0.00
0.5
1
3
5
8
0.5
1
diseased
3
5
8
0.00
0.5
mock uninoc
1
3
5
8
B AB AB
A
B
B
3
5
8
mock uninoc
healthy
Pyroglutamic acid
B
20.00
B
B
B
B
B
Normalised abundance
B
1
Treatment
Malic acid
300.00
0.5
diseased
healthy
Treatment
Normalised abundance
B
200.00
0.00
8.00
A
B
B
B
B
A
5
8
B
B
B
1
3
B
B
B
B
16.00
250.00
200.00
12.00
150.00
100.00
50.00
8.00
4.00
0.00
0.00
0.5
0.5
1
3
5
8
0.5
1
diseased
3
healthy
5
8
1
3
0.5
5
8
mock uninoc
mock uninoc
diseased
Treatment
healthy
Treatment
Figure 4.4: Mean normalised abundance (±SE) for metabolites significantly higher
in later stage infected tissue. Treatments consisted of diseased and healthy tissue
sampled at 0, 1, 3, 5 and 8 dpi from wheat leaves inoculated with Stagonospora
nodorum. Mock-inoculated and uninoculated controls were harvested at 8 days.
GC-MS peak areas were normalised by division using the peak area of an internal
standard (ribitol) followed by division by the sample wet weight. Statistical
significance was calculated for each group using the Tukey-Kramer HSD test,
groups sharing a common letter were not significantly different. N=3 for all
treatments except 1 dpi infected and 3 dpi uninfected (N=2).
147
Glucose Methoxyamine
Glucose methoxyamine 2
40.00
CD BC D
D
D
B
A
D
D
D
D
D
150.00
120.00
90.00
60.00
30.00
Normalised abundance
Normalised abundance
180.00
BC B
C
C
C
C
C
3
5
C
C
20.00
10.00
0.00
0.5
1
3
5
8
0.5
1
diseased
3
5
8
mock uninoc
0.5
healthy
1
3
5
C
C
2.50
AB
C
2.00
1.50
1.00
0.50
12.00
1
3
5
8
0.5
1
3
diseased
5
8
BC BC CD D CD
D D
4.00
2.00
0.00
0.5
mock uninoc
1
3
5
B
B
B
A
8
0.5
1
3
diseased
healthy
5
8
mock uninoc
healthy
Treatment
Unknown 28.0444
B
AB AB
B AB AB
75.00
60.00
45.00
30.00
15.00
0.00
4.00
Normalised abundance
B
A AB AB
C
6.00
Sucrose
B
BC BC AB AB
D D CD CD
8.00
Treatment
90.00
mock uninoc
10.00
0.00
0.5
8
Isocitric acid
C BC AB BC AB A
C
Normalised abundance
BC BC
1
Treatment
14.00
C
0.5
healthy
D-Glucuronic acid
3.00
8
diseased
Treatment
Normalised abundance
A
30.00
0.00
Normalised abundance
B
C
D
B
D
D D
BC
A
D
D
CD CD D
0.5
1
3
5
8
3.00
2.00
1.00
0.00
0.5
1
3
5
8
0.5
1
diseased
3
5
8
mock uninoc
healthy
Treatment
0.5
1
3
5
8
diseased
mock uninoc
healthy
Treatment
Figure 4.5: Mean normalised abundance (±SE) for metabolites significantly higher
in healthy tissue. Treatments consisted of diseased and healthy tissue sampled at 0,
1, 3, 5 and 8 dpi from wheat leaves inoculated with Stagonospora nodorum. Mockinoculated and uninoculated controls were harvested at 8 days. GC-MS peak areas
were normalised by division using the peak area of an internal standard (ribitol)
followed by division by the sample wet weight. Statistical significance was
calculated for each group using the Tukey-Kramer HSD test, groups sharing a
common letter were not significantly different. N=3 for all treatments except 1 dpi
infected and 3 dpi uninfected (N=2).
148
Galactinol
5.00
B AB AB AB AB AB A
AB AB AB AB
AB
7.00
6.00
5.00
4.00
3.00
2.00
1.00
Normalised abundance
Normalised abundance
Erythronic acid
8.00
AB AB AB
0.5
1
3
5
8
0.5
1
diseased
3
5
8
AB AB AB
2.00
1.00
0.5
mock uninoc
1
3
5
8
B
B
A
3
5
8
mock uninoc
healthy
Unknown 13.1643
AB AB AB AB AB AB
4.00
3.00
2.00
1.00
10.00
Normalised abundance
B
1
Treatment
Melibiose
AB AB
0.5
diseased
healthy
5.00
Normalised abundance
A
3.00
Treatment
AB AB AB AB B AB AB AB AB
A AB
0.5
8
A
8.00
6.00
4.00
2.00
0.00
0.00
0.5
1
3
5
8
0.5
1
diseased
3
5
8
mock uninoc
1
3
5
8
0.5
1
diseased
healthy
3
5
mock uninoc
healthy
Treatment
Treatment
Unknown 27.1835
Unknown 26.3226
4.00
AB AB AB B
B AB
A
AB AB AB AB AB
20.00
15.00
10.00
5.00
0.00
Normalised abundance
Normalised abundance
A
4.00
0.00
0.00
25.00
B AB AB
B
AB AB
C C
C
C
C
BC AB AB AB A
C
C
AB AB
C
8
0.5
mock
3.00
2.00
1.00
0.00
0.5
1
3
5
8
0.5
1
diseased
3
healthy
5
8
mock uninoc
0.5
1
3
5
1
diseased
Treatment
3
5
8
healthy
Treatment
Figure 4.6: Mean normalised abundance (±SE) for metabolites significantly lower
in late stage diseased tissue. Treatments consisted of diseased and healthy tissue
sampled at 0, 1, 3, 5 and 8 dpi from wheat leaves inoculated with Stagonospora
nodorum. Mock-inoculated and uninoculated controls were harvested at 8 days.
GC-MS peak areas were normalised by division using the peak area of an internal
standard (ribitol) followed by division by the sample wet weight. Statistical
significance was calculated for each group using the Tukey-Kramer HSD test,
groups sharing a common letter were not significantly different. N=3 for all
treatments except 1 dpi infected and 3 dpi uninfected (N=2).
149
uninoc
Unknown 24.3094
Normalised abundance
8.00
B
B
B
B
B
5
8
B
B
0.5
1
AB
A
A
5
8
A
A
6.00
4.00
2.00
0.00
0.5
1
3
3
diseased
mock uninoc
healthy
Treatment
Unknown 29.4086
Normalised abundance
14.00
AB AB AB B
B AB
A
0.5
8
1
AB AB AB AB AB
12.00
10.00
8.00
6.00
4.00
2.00
0.00
1
3
5
0.5
3
diseased
5
8
mock uninoc
healthy
Treatment
Unknown 37.3751
Normalised abundance
10.00
B
B
B
B
B
B
B
B
B
B
0.5
1
3
5
8
0.5
1
3
5
8
A
B
8.00
6.00
4.00
2.00
0.00
diseased
mock uninoc
healthy
Treatment
Figure 4.7: Mean normalised abundance (±SE) for metabolites significantly lower
in late stage diseased tissue. Treatments consisted of diseased and healthy tissue
sampled at 0, 1, 3, 5 and 8 dpi from wheat leaves inoculated with Stagonospora
nodorum. Mock-inoculated and uninoculated controls were harvested at 8 days.
GC-MS peak areas were normalised by division using the peak area of an internal
standard (ribitol) followed by division by the sample wet weight. Statistical
significance was calculated for each group using the Tukey-Kramer HSD test,
groups sharing a common letter were not significantly different. N=3 for all
treatments except 1 dpi infected and 3 dpi uninfected (N=2).
150
4.4 DISCUSSION
A non-targeted GC-MS approach was used to examine the polar metabolomes
of healthy and diseased wheat leaf tissue from infected leaves, compared to tissue
from mock-inoculated and uninoculated leaves. PCA was used to identify metabolites
which contributed to the principal components explaining most of the variation in the
data set. Metabolites of interest were subjected to statistical analysis to determine
those whose abundance was significantly different between treatments.
4.4.1 Compounds Associated with Infected Tissue Only
The statistically significant difference represented by the presence of mannitol,
trehalose and arabitol in the 8 dpi infected samples is the most likely reason for two of
these replicates being identified as outliers. Since the changes in abundance of these
compounds is of biological significance, they were not excluded from the data set.
4.4.1.1 Mannitol
Mannitol levels rose as the disease progressed, although this increase was only
statistically significant in the 8 dpi samples. Mannitol was not detected in the healthy
tissue samples. This compound was demonstrated as being required for sporulation in
Chapter 3 of this study. The concentration of mannitol was previously shown to
increase dramatically with infection in this pathosystem (Lowe, 2006). Mannitol is the
most abundant polyol found in fungi, and has also been described in over 50 plant
families (Lewis and Smith, 1967), but has not been reported in wheat. In a study
151
involving the ectopic expression of the mannitol-1-phosphate dehydrogenase (mtlD)
gene of Escherichia coli in wheat, there was no mannitol detected in –mtlD plants,
while +mtlD plants constitutively produced mannitol (Abebe et al., 2003). Mannitol
has previously been suggested as a fungal-specific compound in the Cladosporium
fulvum-tomato and Sclerotinia sclerotiorum-sunflower pathosystems based on the
presence of the compound in infected, but not uninfected tissue (Clark et al., 2003;
Jobic et al., 2007). It is likely that mannitol is a fungal-specific compound in the S.
nodorum-wheat pathosystem.
4.4.1.2 Trehalose
Trehalose is a glucose dimer which is widely distributed among bacteria,
fungi, insects, invertebrates and plants (Elbein et al., 2003). While trehalose
accumulation was noted for many microorganisms including phytopathogenic fungi, it
was not considered to occur widely in plants until the unexpected discovery in
Arabidopsis thaliana of a plethora of trehalose biosynthesis genes (Leyman et al.,
2001). Prior to this trehalose had only been associated with the “resurrection” plants
Selaginella lepidophylla and Myrothamnus flabellifolia under conditions of water
stress (Müller et al., 2001). Putative trehalose biosynthetic genes have since been
identified in wheat transcripts/ESTs induced by abiotic stress (Ramalingam et al.,
2006; Mohammadi et al., 2007), and trehalose and enzyme activities corresponding to
trehalose biosynthetic enzymes were reported from wheat subjected to salt and water
stress (El-Bashiti et al., 2005). The generally low levels of trehalose reported from
higher plants has been suggested as being due to the ubiquitous production of the
trehalose-degrading enzyme trehalase (El-Bashiti et al., 2005). The observation here
152
of trehalose only in the 8 dpi infected samples accords with a previous report that
trehalose concentration dramatically increased in the S. nodorum-wheat pathosystem
in concert with pycnidia production (Lowe, 2006). Trehalose was considered to be a
fungal-specific compound in the S. sclerotiorum-sunflower pathosystem since it was
detected in fungal extracts and infected tissue, but not in uninfected plant tissue (Jobic
et al., 2007). Disruption of trehalose 6-phosphate synthase (Tps1) in S. nodorum
reduced trehalose levels in infected tissue to 1% of wild type levels with markedly
reduced sporulation, but little effect on the ability of the mutants to cause lesions
(Lowe, 2006). Similarly, traces of trehalose were still detectable in tps1 mutants in M.
grisea (Foster et al., 2003) and Botrytis cinerea (Doehlemann et al., 2006). These
reports suggest that while trehalose production by the host in response to pathogen
attack cannot be ruled out, it would appear that the nearly all of the trehalose observed
in infected tissue is likely to be of fungal origin.
4.4.1.3 L-Arabitol
Levels of the pentitol L-arabitol rose from 3 dpi on as the disease progressed,
although this increase was only statistically significant in the 8 dpi diseased tissue
samples. This compound was previously reported as having a role in osmotolerance in
S. nodorum (Lowe et al., 2008), Magnaporthe grisea (Dixon et al., 1999) and C.
fulvum (Clark et al., 2003). In the 13C NMR study presented in Chapter 5, arabitol was
the second most abundant metabolite detected in in vitro cultures of the wild type.
Pentitols other than ribitol have rarely been reported from plants (Lewis and Smith,
1967). L-arabitol was reported to be converted to L-ribulose in tobacco, pea and
wheat, although L-arabitol was not considered to be a natural substrate in any of these
153
plants (Kocourek et al., 1964). L-Arabitol was considered to be a fungal-specific
compound in the C. fulvum-tomato pathosystem since it was detected in infected
tissue, but not in uninfected plant tissue (Clark et al., 2003). While it is most likely to
be fungal-specific in the pathosystem investigated in this study, this question could be
resolved by abolishing L-arabitol production in the fungus. However, mutants in
which genes for L-arabitol dehydrogenase (Abd1) and/or L-xylitol dehydrogenase
(Xdh1) were disrupted, were still able to produce basal levels of L-arabitol, and an
additional putative L-arabitol synthesis gene was identified in the S. nodorum genome
(Lowe et al., 2008). These mutants were all fully pathogenic. Targeted gene
replacement of the osmosensory MAP kinase-encoding gene OSM1 in M. grisea
resulted in drastically reduced arabitol production and sensitivity to osmotic stress,
but had no effect on pathogenicity (Dixon et al., 1999). Thus, while this compound
was seen here to increase significantly in the late stages of infection, there was no
evidence that it is required for pathogenicity.
4.4.2 Plant Specific Compounds
Sucrose was significantly higher in the 0.5 dpi healthy tissue samples than in
any of the diseased samples. This is consistent with the conversion of this compound
to its glucose and fructose moieties by host and/or fungal invertase in infected tissue,
and with the fungus acting as a carbon sink. Sucrose was considered to be a plantspecific metabolite in this pathosystem. This was based on the fact that:
1. Sucrose was present in all healthy tissue samples.
2. Sucrose was not detected in the
13
C NMR study of the fungus described in
Chapter 5 below.
154
3. There has been no report of a sucrose synthase in any fungal species and a
BLAST of the S. nodorum genome sequence using the Tuber solanum sucrose
synthase protein sequence (Accession #P10691) produced no hits.
Melibiose is a disaccharide of glucose and galactose and is a hydrolysation
product of the plant trisaccharide raffinose (Hepworth, 1924). Galactinol is formed
from UDP-galactose and myo-inositol and has no known function other than as a
precursor for the formation of the raffinose family oligosaccharides (Zhao et al.,
2004). The observed trend whereby these metabolites decreased with time of infection
to their statistically significant absence in the 8 dpi samples is consistent with their
representing a plant carbon resource which was being consumed by the pathogen.
4.4.3 Miscellaneous Metabolites
The majority of the remaining compounds which exhibited a significant
difference between treatments, were detected in all samples. The exception to this
observation was the unknown with an RT of 24.3094 which was strongly associated
with the later stage non-diseased samples. This unknown was not particularly
abundant in any sample and it would appear that its proximity to the RT of mannitol
resulted in the presence of this unknown compound in the infected samples being
obscured. An extracted ion chromatogram for this unknown suggested that it was also
present in at least some of the infected samples, but the abundance was so low that its
detection was not certain. It obtained very poor matches from the metabolite
databases with the best score being for sorbitol. If this unknown represents an
155
authentic metabolite, it would appear to be a sugar or sugar alcohol of uncertain
identity.
The remaining metabolites consisted of amino acids and amino acid
derivatives (aspartate, glutamate, pyroglutamate), tricarboxylic acid (TCA) cycle
intermediates (citrate, isocitrate, malate), glucose/glucose derivatives, organic acids
(D-glucuronic acid, erythronic acid) and seven unknowns. None of the identified
compounds were specific to either the host or the pathogen. There are a number of
interpretations that could be made in terms of the observed changes in abundance. For
instance, compounds which appeared to decrease with time of infection could be
represented as
1. plant metabolites which were being consumed by the pathogen
2. fungal metabolites which were required during the early stages of infection but
not at the later stages
3. a combination of the above
Without a means of discriminating between the two potential sources of these
metabolites, it was not possible to draw a conclusion regarding the biological
significance of changes in their abundances during the infection process. Organic
acids have previously been shown to be pathogenicity factors/toxins in some
pathosystems e.g. fumaric acid and oxalic acid (Scheffer, 1983). It is therefore of
interest that several of these have shown up as being significantly more abundant in
the 8 dpi infected samples. The fact that these tissues were necrotic, the significant
reduction in or absence of major plant oligosaccharides, and the significant increase in
fungal-specific metabolites, is strongly suggestive that the metabolites present in these
156
tissues would either be of fungal origin, or have been maintained by the fungus rather
than degraded. It is likely then that the metabolites falling into this category warrant
further investigation by targeted gene disruption in the fungus.
4.4.4 No Evidence Found For an Induced Defence Response in the S. nodorumWheat Pathosystem
Many pathosystems are characterised by the production by the host of
antimicrobial metabolites such as the constitutively synthesised phytoanticipans, and
the pathogen-attack-induced phytoalexins (VanEtten et al., 1994). Previous studies on
wheat phytoalexins have focused on the larger, non-polar secondary metabolites and
were investigated by LC-MS and HPLC (Hashimoto et al., 1995; Rémus-Borel et al.,
2005; Rémus-Borel et al., 2006). A number of polar phytoalexins have previously
been described in other plants including resveratrol in tomato (Ragab et al., 2006),
rishitin and lubimin in potato (Fanelli et al., 1992), β-ionone, geranylacetone and
terpinyl acetate in cantaloupe (Lamikanra et al., 2002), momilactone A in rice
(Atawong et al., 2002), and galactinol in cucumber (Kim et al., 2008). Such a
compound would have featured as a significant difference between the mockinoculated/uninoculated controls and the healthy tissue from the infected leaves.
There was no such compound observed. While galactinol was seen to be significantly
higher in the later stage healthy versus diseased tissue from infected leaves, it was not
significantly higher than the non-infected control. It may be that S. nodorum does not
elicit a defence response from the host, or that the levels of any such response were
below the detection limits of the method used. Since the wheat plants were not grown
157
under conditions of absolute sterility, it is also possible that endogenous microorganisms may already have primed any inducible defence response.
4.5 CONCLUSION
An undirected GC-MS metabolomics analysis of healthy and diseased tissue
from wheat leaves infected with S. nodorum was undertaken. PCA of the data set
highlighted metabolites which contributed to the principal components explaining the
variation between the treatments. Statistical analysis of these metabolites showed that
the fungus-specific or -associated metabolites mannitol, trehalose and arabitol were
significantly higher in the later stage infected samples. The plant-specific metabolites
sucrose, galactinol and melibiose were absent in the later stage infected samples. A
number of amino acids, organic acids, TCA cycle intermediates and unknown
metabolites showed some significant changes in abundance in healthy versus infected
tissue and may warrant further investigation. It was not possible to conclusively
determine the organism of origin of these latter compounds in the infected tissue.
There were no metabolites which differentiated healthy tissue from pathogeninoculated versus non-inoculated leaves so as to suggest an inducible host defence
response in the former.
158
CHAPTER 5: 13C-NMR INVESTIGATION OF
MANNITOL METABOLISM IN
Stagonospora nodorum
159
5.1 INTRODUCTION
5.1.1 Overview of Technique
The technique of nuclear magnetic resonance (NMR) spectroscopy is founded
on the existence of a magnetic moment in the nuclei of atoms with an odd mass
number, or an even mass number but odd atomic number, with the result that these
behave like spinning magnetic bodies (Stothers, 1972). When a strong external
magnetic field, H0, is applied to such nuclei, their magnetic fields align either parallel
or antiparallel to H0. Upon irradiation with radiofrequency energy of the correct
frequency, the lower-energy-state, parallel-aligned nuclei absorb energy and spin-flip
to the higher energy state, and are said to be in resonance with the applied radiation
(McMurry, 1984). The spectra produced by modern NMR spectrometers plot the
effective magnetic field strength applied to nuclei against their intensity of absorption
of energy (Ratcliffe, 1996).
The measurement of the nuclear magnetic moment was first achieved by use
of the molecular beam resonance method (Rabi et al., 1938; Rabi et al., 1939)
involving the changes in orientation of nuclear spin exhibited by molecular beams in a
strong, externally applied magnetic field and in a high vacuum. It was subsequently
and simultaneously demonstrated by refinements in the technique, that NMR
spectroscopy could be applied to bulk solid (Purcell et al., 1946) and liquid samples
(Bloch, 1946; Bloch et al., 1946) and with improvements in both sensitivity and
precision. Purcell and Bloch were awarded the Nobel Prize for physics in 1952 for
their work on NMR.
160
An unexpected and surprising observation was next made, that the frequency
of resonance of
19
F and
14
N nuclei was dependent upon the chemical compound in
which they were contained (Dickinson, 1950; Proctor and Yu, 1950). The term
“chemical shift” was coined in order to describe the observed differences in
radiofrequency required to bring isotopic nuclei in different chemical environments
into resonance (Arnold et al., 1951).
The progress of
13
C-NMR spectroscopy was initially limited by the high
detection threshold of the spectrometers, and the low natural abundance (1.1%) of the
13
C isotope (Aubert et al., 1996a). Improvements in instrument sensitivity and
computer techniques have led to NMR being used extensively for structural analysis
of novel compounds, and for metabolite profiling (McMurry, 1984). When used in
conjunction with growth on
13
C-enriched substrates, it provides a powerful tool for
following the metabolism of that substrate, for illustrating differences in metabolism
on different substrates and between different strains and species, and for elucidating
pathways of carbon metabolism (Jeffrey et al., 1991; Ratcliffe, 1996). In conjunction
with other techniques of magnetic resonance, it currently has a wide range of
applications in physiology, biology, chemistry, pharmacy and medicine (Shulman and
Rothman, 2005; Webb, 2006).
5.1.2 Advantages and Disadvantages of NMR
One of the advantages of NMR compared to other metabolome
characterisation methods such as GC-MS, is that no chemical alteration or isolation of
161
the compounds is required, requiring relatively simple sample preparation and
reducing the incidence of experimental artefacts (Yoshida et al., 1984; Pfeffer and
Shachar-Hill, 1996). The technique is also non-destructive (Yoshida et al., 1984) ,
allowing spectra of different nuclei to be acquired for the same sample, and permitting
the sample to be subsequently analysed by a different technique (Last et al., 2007).
13
C-NMR spectra can be acquired from intact mycelium or from mycelial extracts,
and the two procedures have been shown to deliver equivalent spectra (Martin et al.,
1985; Ratcliffe, 1996). The use of
13
C-labelled substrates turns the low natural
abundance of 13C to advantage, since it allows the identification of multiply-labelled
metabolites and the quantitation of the label distributed in the spectrum (Ratcliffe,
1996; Pfeffer et al., 2001).
Its main disadvantage is that it is relatively insensitive technique requiring
milligram amounts of sample with metabolites required to be present at millimolar
concentrations in order to be detected (Pfeffer et al., 2001; Chatham et al., 2003).
There are some issues involved with comparisons between spectra on the basis of
chemical shifts (Wishart and Sykes, 1994; Wishart and Case, 2001) which are
discussed below. The quantitation of the proportion of
13
C accruing to different
resonances in a spectrum, and comparing this between spectra, can also be
problematic. There are a number of approaches to handling this issue and these are
also discussed below.
162
5.1.3 13C-NMR Studies in Filamentous Fungi
13
C-NMR studies have been conducted on a range of filamentous fungi
including ectomycorrhizal ascomycetes and basidiomycetes, saprophytes and
phytopathogens. The purposes of these studies have ranged from providing baseline
information for fungicidal mechanism-of-action studies (Forgue et al., 2006), to
furthering the understanding of carbon assimilation and cycling pathways in fungi
(Martin et al., 1984; Dijkema et al., 1985; Dijkema and Visser, 1987; Thomas and
Baxter, 1987; Martin et al., 1988; Ramstedt et al., 1989; Peksel et al., 2002; RangelCastro et al., 2002), including how these pathways are affected in mycorrhizal fungi
and phytopathogens under free living versus host-associated conditions (Shachar-Hill
et al., 1995; Martin et al., 1998; Bago et al., 1999; Jobic et al., 2007). Key
conclusions from these studies have been that mannitol is amongst the most abundant
soluble metabolites (Yoshida et al., 1984; Dijkema et al., 1985; Martin et al., 1985;
Dijkema and Visser, 1987; Ramstedt et al., 1989; Peksel et al., 2002; Jobic et al.,
2007). Secondly it has been noted in studies using [1-13C]-glucose as a growth
substrate, that there is a “scrambling” of the label originating from the C1, which
results in label appearing on the C1 and C6 of molecules such as glucose and
trehalose (Martin et al., 1985; Martin et al., 1988; Ramstedt et al., 1989; Peksel et al.,
2002; Rangel-Castro et al., 2002). This has been explained as occurring via the
metabolism of the labelled substrate via mannitol, with the operation of the purported
mannitol cycle, and the symmetry of the compound, resulting in the observed
scrambling (Martin et al., 1985; Martin et al., 1988; Ramstedt et al., 1989; RangelCastro et al., 2002). Given the evidence against the mannitol cycle in Stagonospora
nodorum, the mannitol mutants created presented an opportunity to re-examine the
163
metabolic fate of
13
C-labelled carbon, and to further elucidate the mechanism of
mannitol metabolism.
5.1.4 Aims of the Study
The aims of this study were to characterise the differences between the S.
nodorum wild type strain SN15, and mutants with a disrupted mannitol
dehydrogenase gene (mdh1-71), a disrupted mannitol 1-phosphate dehydrogenase
gene (mpd1-1), or with both of these genes disrupted (mpd1mdh1-107) using
13
C-
NMR spectroscopy. It was hypothesised that differences between the natural
abundance spectra, and spectra acquired after growth on [1-13C]-labelled substrates
would reveal differences in the pathway(s) of carbon metabolism between the strains
which would firstly elucidate how the mdh1-71 strain is able to catabolise mannitol.
It was further hypothesised that the scrambling of the
13
C label seen in other
studies, resulting in labelling of both terminal carbons of glucose and trehalose, can
occur by either of the mannitol metabolic pathways. In the double mutant, however,
this scrambling mechanism will be inoperable, and the presence or absence of label in
the C6 of trehalose and glucose will indicate whether other pathways such as the
aldose/triosephosphate isomerase/pentose phosphate pathway contribute significantly
to scrambling.
164
5.2 MATERIALS AND METHODS
5.2.1 Preparation of Standards
Compound standards were prepared by making a 60-250 mM solution of the
compound in 1 mL D2O (99 atom %, Sigma-Aldrich Co., St. Louis, MO, USA). The
solution was lyophilised overnight in a Savant FDC206 freeze-drying chamber
(Savant Scientific Instruments, Farmingdale, NY) attached to an Heto Maxi-Dry Lyo
freeze dryer (Heto-Holten, Allerød, Denmark) and stored at -80 °C until required.
Prior to NMR spectroscopy the freeze-dried standard was resuspended in 1 mL D2O,
centrifuged at 20800 g for 10 min in a benchtop Eppendorf Centrifuge (Model 5417C,
Eppendorf-Netheler-Hinz GmbH, Hamburg, Germany) to pellet any debris, and 700
μL transferred to a NMR tube for spectroscopy.
5.2.2 Flask Culture of Fungal Strains
The strains SN15, mdh1-71, mpd1-1 and mpd1mdh1-107 were used in this
experiment.
5.2.2.1 Natural Abundance Cultures
Natural abundance (NA) cultures were inoculated with mycelium and spores
scraped from 1/3 of a CZV8CS agar plate culture into a 250 mL flask containing 50
165
mL MM-C liquid medium with 40 mM glucose. An additional set of mpd1mdh1-107
NA cultures were started with inoculum from minimal medium agar plates. The flasks
were incubated on a Certomat® R shaker (B. Braun, Melsungen, W. Germ.) operating
at 140 rpm, for 3 days at 20 °C in the dark. Mycelium was harvested after 3 days into
a pre-weighed sterile 50 mL Falcon tube and centrifuged for 10 min at 3000 g at 4 °C.
The supernatant was discarded and the pellet resuspended in 50 mL milliQ H2O and
centrifuged for a further 10 min at 3000 g at 4 °C. The supernatant was discarded and
the pelleted mycelium was snap frozen by placing the tubes in liquid nitrogen.
Samples were lyophilised overnight in a Savant FDC206 freeze-drying chamber
(Savant Scientific Instruments, Farmingdale, NY) attached to an Heto Maxi-Dry Lyo
freeze dryer (Heto-Holten, Allerød, Denmark). Tubes were re-weighed to determine
the dry weight of the mycelium, prior to being replaced in liquid nitrogen and stored
at -80 °C. Three replicates of each treatment were prepared.
5.2.2.2 [1-13C]-Glucose-Labelled Cultures
Cultures were prepared as for NA cultures (5.2.2.1) except that the carbon
source added to the MM-C liquid medium was 40 mM D-[1-13C]-glucose (SigmaAldrich Inc., St. Louis, MO).
5.2.2.3 [1-13C]-Mannitol-Labelled Cultures
166
5.2.2.3.1 Assay of mannitol uptake
Three 50 mL minimal media flask cultures were prepared for each of the
strains SN15, mdh1-71, mpd1-1 and mpd1mdh1-107 and incubated for four days.
Cultures were harvested and centrifuged as described above and 1.5 mL of the
minimal medium retained for assay from each culture. The cultures were then washed
in 50 mM Tris-HCl pH 7.5 and transferred to fresh flasks containing MM-C with 40
mM mannitol. For control purposes a set of flasks were prepared containing MM-C or
MM-C with 40 mM mannitol, but to which no fungal culture was added. An aliquot
of 1.5 mL was taken from each of these control flasks and retained for assay. All
flasks were incubated under standard growth conditions. At 24 h and 48 hours
incubation a 1.5 mL aliquot was taken from each flask and retained for assay.
A flask culture of SN15 was grown for three days in 50 mL MM-C + 40 mM
sucrose and harvested and prepared for enzyme assay as described above (Section
3.2.4). The mannitol oxidation activity of the mannitol dehydrogenase enzyme
(Section 3.2.4.3.3) in the desalted extract was used to determine the amount of
mannitol in the spent medium and controls. A standard curve was prepared using 50
μL per 1 mL assay volume of 0, 10, 20, 30, 40 and 50 mM mannitol. The activity of
all collected samples was similarly assayed using 50 μL per assay volume. Additional
controls using 50 μL milliQ water or 50 μL 20 mM or 40 mM mannitol as the
substrate were included in each run. The mannitol standard curve was used to
calculate the amount of mannitol in each sample. The allowed the determination of
the amount of mannitol uptake for each culture.
167
5.2.2.3.2 Preparation of [1-13C]-mannitol-labelled cultures
Cultures were prepared as for the natural abundance cultures (5.2.2.1) except
that strains were harvested and washed after two days incubation on 40 mM glucose.
The harvested mycelium was washed and transferred to a fresh 250 mL flask
containing 50 mL MM-C with 20 mM D-[1-13C]-mannitol (Omicron Biochemicals,
Inc., South Bend, IN). Cultures were incubated for a further 24 h and then harvested
and stored as for natural abundance cultures.
5.2.2.4 [1-13C]-Glucose Feed-Chase Cultures
A set of 12 SN15 flask cultures were incubated for three days, harvested and
washed as per the natural abundance cultures (5.2.2.1) The washed cultures were
transferred to fresh 250 mL flasks containing 50 mL MM-C with 40 mM D-[1-13C]glucose (Sigma-Aldrich Inc, St. Louis, MO) and returned to the shaker. At one hour
and four hours incubation on the labelled medium, three cultures were harvested,
washed and snap frozen in liquid nitrogen for lyophilisation. The six remaining
cultures were also harvested and washed after 4 hours on the labelled medium, and
then transferred to a further set of 250 mL flasks containing 50 mL MM-C with 40
mM unlabelled glucose. At one hour and four hours post-transfer to the unlabelled
medium, three cultures were harvested, washed and snap frozen in liquid nitrogen for
lyophilisation. All cultures were lyophilised overnight, weighed, and stored as per the
natural abundance cultures.
168
5.2.3 Methanol/Water Extraction of Polar Metabolites
Samples were retrieved from the -80 ºC freezer and ground in a sterile mortar
and pestle with liquid nitrogen to a fine powder. Once ground, 20 mL of a -20 °C
70:30 methanol:water solution was added and the sample was ground for a further 1
minute. The resuspended sample was poured into a funnel lined with #1 Whatman
paper filter and the filtrate collected into a flask. The mortar and pestle were rinsed
twice with a further 5 mL -20 °C 70:30 methanol:water solution and this was added to
the funnel and the filtrate collected. The filtrate volume was reduced by rotary
evaporation using an RE111 Rotavapor (Büchi, Switzerland) with the water bath
temperature set to 45 °C until about 2-3 mL remained. The samples were transferred
to a sterile 10 mL Falcon tube, snap frozen in liquid nitrogen and lyophilised
overnight as described above (5.2.2.1) and stored at -80 °C until ready for NMR
analysis.
5.2.4 NMR Tube Preparation
Prior to use, NMR tubes (Kontes Glass Company, Vineland, New Jersey,
USA) were rinsed 2-3 times with de-ionised water. Tubes were then rinsed 2-3 times
with 100% acetone to expel any remaining water and dried overnight in a 150 °C
oven.
169
5.2.5 Sample Preparation for NMR Analysis
Samples were retrieved from the -80 °C freezer, resuspended in 1 mL D2O (99
atom %, Sigma-Aldrich Inc., St. Louis, MO, USA) and transferred to a pre-weighed
1.5 mL Eppendorf tube. Samples were lyophilised overnight as described previously
(5.2.2.1), reweighed to determine the dry weight of the polar extract, and stored at
room temperature until ready for NMR analysis. Just prior to NMR spectrometry,
samples were resuspended in 1 mL D2O and centrifuged in a benchtop Eppendorf
microcentrifuge at 20800 g for 10 min to pellet any particulate debris. The liquid
fraction was transferred to a fresh 1.5 mL Eppendorf tube, re-centrifuged, and 600 μL
transferred to a clean NMR tube and taken for NMR spectroscopy. The pellet in the
pre-weighed Eppendorf tube was evaporated to dryness in a 68 °C heating block and
the tube re-weighed to determine the dry weight of the pellet. This weight was
deducted from the original weight of the dried extract to determine the weight of the
soluble polar extract in the NMR sample.
5.2.6 NMR Spectra Acquisition
The
13
C NMR spectra were acquired using a Bruker Avance DPX-300
Spectrometer (Bruker Instruments Inc, Billerica, MA) operating at 75.5 MHz,
equipped with a 5 mm 1H/multinuclear probe, and interfaced with a console running
Xwin-NMR Version 3.5. Spectra were routinely run at 300 K with locking on the
D2O solvent. For samples containing carbon standards or
13
C-labelled samples, 512
scans were sufficient to obtain a satisfactory spectrum. For natural abundance or very
low dry weight samples 10,000 scans were routinely acquired.
170
5.2.7 NMR Spectra Analysis
5.2.7.1 Software
Spectra were analysed using MestReNova 5.2.1 NMR analysis software
(MestreLab Research, A Coruña, Spain). Assignment of spectral peaks to compounds
was based upon comparison of their chemical shifts and relative intensities with those
of prepared standards or with compounds listed in the Spectral Database for Organic
Compounds (SDBS) (AIST, 2008), the Aldrich Library of
13
C and 1H FT NMR
Spectra (Pouchert and Behnke, 1993) and the Chemical Shift tables compiled by Fan
(1996).
5.2.7.2 Compound Identity and Label Quantification
5.2.7.2.1 Internal referencing of chemical shifts
The MestReNova sensitivity threshold for peak selection was adjusted to that
level which was sufficiently low to pick up the majority of biologically relevant
peaks, but without picking random noise peaks. Negative peaks and peaks with a
chemical shift of less than zero were automatically precluded. The chemical shifts of
the selected peaks were referenced to the largest peak of the most abundant compound
in the spectrum using the values from the compound standard spectra. Typically this
was the β-anomer of the carbon 1 (C1) of glucose (95.83 ppm), the C1,6 of mannitol
(63.16 ppm) or the C1 of trehalose (93.16 ppm). When possible, the compound which
was used as the substrate (glucose or mannitol) was used for this referencing process.
171
In those instances where the substrate had been exhausted, the principal metabolic
product was used (mannitol or trehalose). These were all 6-carbon compounds and the
peaks selected had an unambiguous chemical shift, which facilitated their
identification.
The re-referenced peak data was downloaded and compared to the library of
compound standards. Peaks were initially assigned to compounds where the
difference between the chemical shifts of the sample peak and standard peak were less
than 0.1 ppm. Peaks which were assigned to the same compound were assessed on the
basis of the maximum variation of their relative chemical shifts (MaxVar(RCS)) and
the maximum relative peak intensity (Max(RPI)) to confirm their identity.
5.2.7.2.2 MaxVar(RCS)
The MaxVar(RCS) was determined by calculating the differences between the
actual chemical shifts for each peak assigned to a compound, and their ideal chemical
shifts as established by the compound standard. The smallest difference value was
deducted from the largest difference value and where the net difference was less than
0.05 ppm, the peaks were considered to be a good match for the compound. This can
be expressed as:
MaxVar(RCS) = Max(CS1-ICS1, CS2-ICS2,…CSn-ICSn) - Min(CS1-ICS1, CS2ICS2,…CSn-ICSn)
Where MaxVar(RCS) = maximum variation in relative chemical shift
172
Max = maximum of calculated differences
Min = minimum of calculated differences
CS1-n = chemical shift for carbons 1-n of candidate peaks for a compound
ICS1-n = ideal chemical shift for carbons 1-n from compound standard
5.2.7.2.3 Max(RPI)
A set of ideal NA relative peak intensities (RPI) for each compound was
calculated based on the compound standard NA spectra, assigning the most intense
peak (the base peak) a value of 100%. The RPI for all other resonances in the
compound standard were determined in relation to the base peak. The RPI for the
peaks assigned to a compound in a biological spectrum were determined by applying
the intensity of the compound standard base peak (i.e. 100%) to the peak which best
matched the base peak in terms of chemical shift. This was not necessarily the most
intense peak in the biological candidate peaks. The RPI were calculated for all other
peaks assigned to the compound in the biological spectrum. The Max(RPI) was
determined by dividing the calculated RPI for each candidate peak by its ideal RPI
and deducting 1. Given that there was an observed variation between standards from
different sources as noted below (Section 5.3.1), and given further that two peaks
from the same compound could have their intensity altered in opposite directions,
there was some flexibility required in the use of this parameter. Generally, as long as
the RPI of particular candidate peak was within the range of -50% to +80% of the
standard, it was not rejected as belonging to that compound. The Max(RPI) can be
expressed as:
173
Max(RPI) = Max((ORPI1/IRPI1)-1), (ORPI2/IRPI2)-1),…(ORPIn/IRPIn)-1))
Where Max(RPI) = maximum difference in observed versus ideal relative
peak intensities
ORPI1…n = observed relative peak intensity for candidate peaks 1…n
IRPI1…n = ideal relative peak intensity for peaks 1…n of a compound
If a peak was rejected as belonging to the assigned compound, the remaining
peaks were reassessed using Max(RPI) to determine whether they constituted
acceptable candidates for the compound.
5.2.7.2.4 Missing peaks
Where some peaks were identified for a compound, but others were not
present, an iterative search process was undertaken. Where the missing peak(s) were
of low intensity, the sensitivity threshold could be lowered to pick up the missing
peak(s). If this resulted in too much noise being picked up then the peak was
discounted. Where the chemical shift of a candidate peak resulted in MaxVar(RCS)
being greater than 0.05 ppm, but less than 0.15 ppm, it was only considered if the
peak could be assigned unambiguously, if there was evidence from other biological
spectra that this peak was subject to variation in its chemical shift, and if the
Max(RPI) did not result in its rejection.
174
5.2.7.2.5 Comparison of relative abundances between spectra
Direct comparison of the relative abundance of compounds between spectra on
the basis of resonance peak intensity was not possible since the scale used is an
arbitrary one. To enable quantitative comparisons, the sum of the intensities for all
peaks above the detection threshold was calculated. For a compound in the spectrum,
the ratio of the sum of the intensities of peaks assigned to the compound to the total
intensity for the spectra, enabled the percentage of total intensity to be determined.
This can be expressed as:
%TI = (Σ(I1, I2,…In))/(ΣI)
Where %TI = percentage of total intensity
I1…n = intensity of peak 1…n of a compound
ΣI = sum of all resonance peak intensities above the detection threshold
Unidentified peaks which comprised a high percentage of total intensity were
the subject of further analysis to provide information concerning their structure and/or
identity. This included visual examination of spectra, and comparison of relative
chemical shifts and relative peak intensities between spectra, to determine
unidentified peaks which might be associated with each other, and which accounted
for consistent proportions of total intensity.
175
5.2.7.2.6 Quantification of 13C-labelling
Peaks assigned to the same compound were compared to determine whether
13
C-labelling, above NA levels, had occurred as a result of growth on
13
C-labelled
substrate. The RPI of the peaks were calculated using the same formula as for
determining Max(RPI) above (Section 5.2.7.2.3). A peak was not considered to be
labelled unless it had a Max(RPI) of 100% or more i.e. one-fold labelling or more.
There were circumstances where this approach required modification. These
occurred where the base peak in the compound standard was the putatively labelled
peak in the biological spectrum, where the base peak was co-located with a peak from
another compound preventing unambiguous assignment, or where it was subject to
peak splitting under the influence of a heavily labelled neighbouring carbon. In these
cases the RPI of all sibling peaks were adjusted using the next most intense peak from
the compound standard. This procedure was compromised where there were no
sibling peaks to form the basis of the comparison, or where the putatively labelled
peak had the same chemical shift as the peak of another compound.
5.3 RESULTS
5.3.1 Standards
Natural abundance 13C-NMR spectra were acquired for a number of compound
standards as listed (Table 5.1). While the general position of the resonance peaks
176
Table 5.1: Standard compounds for which 13C natural abundance NMR spectra were
acquired.
Compound
250 mM L-alanine
250 mM L-arabinose
250 mM D-arabitol
200 mM L-arginine
200 mM L-asparagine
37.6 mM L-aspartate
200 mM citric acid
250 mM D-fructose
100 mM D-fructose 6-phosphate
200 mM D-galactose
200 mM D-gluconate
200 mM D-glucose
200 mM D-[1-13C]-glucose
75 mM D-glucose 6-phosphate
200 mM L-glutamate
200 mM L-glutamine
200 mM glycerol
60 mM inosine
200 mM L-malate
200 mM D-mannitol
200 mM D-[1-13C]-mannitol
100 mM D-mannitol 1-phosphate
200 mM D-mannose
200 mM meso-erythritol
200 mM L-methionine
200 mM L-ornithine
200 mM L-phenylalanine
200 mM pyruvate
200 mM L-serine
200 mM D-sorbitol
200 mM D-sucrose
200 mM L-threonine
200 mM D-trehalose
200 mM L-tryptophan
200 mM xylitol
Supplier
Sigma-Aldrich Inc., St. Louis, MO, USA
Sigma-Aldrich Inc., St. Louis, MO, USA
Sigma-Aldrich Inc., St. Louis, MO, USA
Sigma-Aldrich Inc., St. Louis, MO, USA
Sigma-Aldrich Inc., St. Louis, MO, USA
Sigma-Aldrich Inc., St. Louis, MO, USA
Merck Pty. Ltd., Kilsyth, Vic.
BDH Laboratory Supplies, Poole, UK
Sigma-Aldrich Inc., St. Louis, MO, USA
Sigma-Aldrich Inc., St. Louis, MO, USA
Sigma-Aldrich Inc., St. Louis, MO, USA
Merck Pty. Ltd., Kilsyth, Vic.
Sigma-Aldrich Inc., St. Louis, MO, USA
Sigma-Aldrich Inc., St. Louis, MO, USA
Sigma-Aldrich Inc., St. Louis, MO, USA
Sigma-Aldrich Inc., St. Louis, MO, USA
Sigma-Aldrich Inc., St. Louis, MO, USA
Sigma-Aldrich Inc., St. Louis, MO, USA
Sigma-Aldrich Inc., St. Louis, MO, USA
BDH Laboratory Supplies, Poole, UK
Omicron Biochemicals, Inc., South Bend, IN
Sigma-Aldrich Inc., St. Louis, MO, USA
Sigma-Aldrich Inc., St. Louis, MO, USA
Sigma-Aldrich Inc., St. Louis, MO, USA
VWR International Ltd, Poole, UK
VWR International Ltd, Poole, UK
VWR International Ltd, Poole, UK
Sigma-Aldrich Inc., St. Louis, MO, USA
Sigma-Aldrich Inc., St. Louis, MO, USA
VWR International Ltd, Poole, UK
Univar, Seven Hills, NSW
Sigma-Aldrich Inc., St. Louis, MO, USA
Sigma-Aldrich Inc., St. Louis, MO, USA
Sigma-Aldrich Inc., St. Louis, MO, USA
The Sweet Life, Perth, WA
177
correlated with those reported in the SDBS database, the Aldrich Library, and Fan’s
Chemical Shift tables, the actual chemical shifts of the peaks were not identical. For
example, the C1,6 peak of mannitol was reported as being located at 64.36 ppm
(SDBS), 65.99 ppm (Aldrich Library), 73.60 ppm (Fan) and 63.16 ppm (this study).
Fan’s Chemical Shift tables were compiled from a wide variety of sources and were
generally found to agree poorly with the other two resources and from the data
collected in this study. Apart from Fan’s Chemical Shift tables, the distances between
the peaks of mannitol for other spectra, including those quoted in some other studies
(Table 5.2), were largely in agreement, with the largest variation being 0.3 ppm.
It was also observed that the relative heights of the peaks exhibited some
variation between the spectra of different sources (Figure 5.1). To account for the
contribution of peak width to intensity, the relative peak intensities for each
compound standard in this study were calculated, assigning the most intense peak a
value of 100%. The ACNFP library of compound standards is included in the
appendix (Table 8.3). There was still some disagreement between published data in
terms of relative peak intensities. For example, while this study and the SDBS agreed
that the C2,5 of mannitol was the most intense peak of the three, they disagreed on the
order of intensity for the other two peaks of this compound.
There were some instances where the chemical shifts of resonance peaks in
different compounds were co-located. This is illustrated by the C5 of L-arabitol and
one of the spinning sidebands of [1-13C]-D-mannitol (Figure 5.2).
178
Table 5.2:
13
C-NMR chemical shifts (ppm) for the peaks of D-mannitol from Standard Compound compilations and from reported
experimental observations. The calculated differences in the relative positions of the C2,5 and C3,4 – and C2,5 and C1,6, and the net
difference in published chemical shifts for each peak, are shown
Data Source
C2,5
C3,4
C2,5 – C3,4
C1,6
C2,5 – C1,6
700 mM Mannitol in D2O - SDBS (AIST, 2008)
72.05
70.48
1.57
64.36
7.69
Mannitol in D2O (concentration not given) - Aldrich Library
73.57
72.00
1.57
65.99
7.58
Mannitol (no concentration or solvent given) (Fan, 1996)
76.3
75.3
1.0
73.6
2.7
200 mM Mannitol in D2O (this study)
70.74
69.18
1.56
63.16
7.58
Agaricus bisporus (Donker and Braaksma, 1997)
71.7
70.1
1.6
63.9
7.8
Aspergillus nidulans (Dijkema et al., 1985)
71.8
70.2
1.6
64.4
7.4
Sphaerosporella brunnea (Martin et al., 1988)
72.2
70.6
1.6
64.6
7.6
Magnaporthe [syn. Pyricularia] oryzae (Yoshida et al., 1984)
72.0
70.7
1.3
64.3
7.7
70.82 (±0.05)
69.25 (±0.05)
1.57
63.22 (±0.05)
7.6
5.56
6.12
-
10.44
-
(Pouchert and Behnke, 1993)
Stagonospora nodorum (this study)*
Maximum chemical shift – minimum chemical shift
* mean (±SE) prior to re-referencing to the internal standard. N ≥ 55.
C1,6
A
B
C1,6
C
24000
23000
22000
C1,6
21000
20000
19000
18000
17000
16000
15000
14000
13000
12000
11000
10000
9000
8000
7000
6000
5000
4000
3000
2000
1000
0
75
Figure 5.1:
13
74
73
72
71
70
69
68
f1 (ppm)
67
66
65
64
63
62
61
C NMR spectra for D-mannitol illustrating source-dependent
differences in relative height of peaks.
A: Spectral Database for Organic Compounds (AIST, 2008)
B: Aldrich Library of 13C and 1H FT NMR Spectra (Pouchert and Behnke, 1993).
C: This study.
Note the relative height of the C1,6 peak which is the least intense of the three
peaks in this study, the most intense in the Aldrich spectrum and of median
intensity in the SDBS spectrum.
180
6 3 .4 2
6 3 .1 6
6 2 .9 5
66 22 .8
.8 67
6 3 .6 9
7 0 .4 7
7 0 .3 6
7 0 .1 7
7 1 .0 2
.8 35
77 00 .7
7 1 .4 7
6 9 .1 9
6 9 .1 7
M1,6
M3/M4
M5
SB
SB
A1
M2
M2
A5
A3
A4
A2
74
73
72
71
70
69
68
67
66
65
64
63
62
61
f1 (ppm )
Figure 5.2:
13
C-NMR spectra showing co-location of the chemical shifts of the C1
resonance peak of L-arabitol (red) and a spinning sideband of the C1,6 resonance
peak of [1-13C]-D-mannitol (black). Abbreviations: A=arabitol; M=mannitol. The
M1,6 peak is truncated in this figure. Note that the M2 of mannitol is split and the
M3 and M4 peaks have resolved separately under the influence of the 100% labelled
M1. The vertical scales have been adjusted to illustrate the situation where a large
accumulation of labelled mannitol can obscure the A1 and even the A5 when arabitol
is present at a lower abundance.
181
5.3.2 Identified Compounds
5.3.2.1 13C Natural Abundance Spectra
5.3.2.1.1 Replicates Inoculated from CZV8CS Agar Cultures
Representative spectra of SN15 and mpd1mdh1-107 are shown (Figures 5.3
and 5.4). An average of more than 80% of the total intensity for all NA spectra from
cultures started with inoculum from CZV8CS plates, was accounted for by nine
metabolites. The most abundant identified compounds in the strains, apart from the
growth substrate glucose, were glycerol, mannitol or trehalose (Figure 5.5A).
Mannitol was the principal metabolite found in SN15 and mdh1-71 with only traces of
glycerol and trehalose detected in these strains. In mpd1-1 and mpd1mdh1-107, the
principal metabolite was trehalose, followed by glycerol, and with about 10% of the
mannitol content of the wild type. These differences in the major metabolites were
statistically significant, with SN15 and mdh1-71 comprising one group, and mpd1-1
and mpd1mdh1-107 forming a second group.
Glucose was detected in all samples (Figure 5.5A) except for one SN15
replicate, and also accounted for less than 10% of total intensity in a second SN15
replicate and one of the mdh1-71 replicates. In all other samples it accounted for
greater than 10% of total intensity. The mean proportions of total intensity were not
found to be statistically different between the strains.
182
A
Gβ5
Gβ1
Gβ3
Gα1
T1
4E+ 05
B
M2,5
M3,4
4E+ 05
M1,6
4E+ 05
3E+ 05
2E+ 05
2E+ 05
Gβ5
Gα2/Gα5
Gβ3
Gβ2
Gβ4
Gα4
2E+ 05
A1
1E+ 05
A5
A4 A2/3
Gα3
Gly2
Gβ6
Gly1,3
Gα6
50000
0
76
75
74
73
72
71
70
69
68
f1 (ppm)
67
66
65
64
63
62
61
C
34000
32000
Gln3
30000
28000
Gln4
26000
Ala3
24000
22000
20000
Gln2
18000
16000
ArgαCH2
Ala2
14000
Glt3
AsnβCH
12000
Glt4
Glt2
10000
ArgδCH
8000
Arg Arg
6000
Orn5
4000
2000
0
60
55
50
45
40
35
30
25
20
15
f1 (ppm)
Figure 5.3: Natural abundance
13
C NMR spectrum of SN15 showing the regions
from (A) 75-100 ppm, (B) 60-76 ppm and (C) 15-60 ppm. Carbons have been
assigned using the compound abbreviations A=arabitol; Ala=alanine; Arg= arginine;
Asn=asparagine;
G=glucose;
Gln=glutamine;
Glt=glutamate;
Gly=glycerol;
M=mannitol; Orn=ornithine; T=trehalose. All chemical shifts were referenced to the
βC1 peak of glucose. The scales have been adjusted to the height of the most intense
peak in each section.
183
75.87
75.67
92.01
93.14
95.83
A
T1
Gβ5
Gβ3
Gβ1
Gα1
B
Gly1,3
T3
T5
T2
T4
T6
Gα2/α5 Gly2
Gβ5
Gα4/β4
A4
Gβ3
M2,5
M1,6
M3,4
Gβ2
A1
Gα3
Gα6
Gβ6
A5
A2/3
C
Gln3
Gln4
Ala3
Gln2
ArgαCH2
Glt2
Ala2
ArgδCH
MalβCH
Glt3
Glt4
Arg Arg
Figure 5.4: Natural abundance 13C NMR spectrum of mpd1mdh1-107 showing the regions
from (A) 75-100 ppm, (B) 60-76 ppm and (C) 15-60 ppm. Carbons have been assigned
using the compound abbreviations Ala=alanine; A=arabitol; Arg= arginine; G=glucose;
Gln=glutamine; Glt=glutamate; Gly=glycerol; M=mannitol; Mal=malate; T=trehalose. All
chemical shifts were referenced to the βC1 peak of glucose. The scales have been adjusted
to the height of the most intense peak in each section.
184
(A)
SN15
mdh1-71
mpd1-1
mpd1mdh1-107
80
Mean % of Total Intensity
A
A
B B
70
60
50
40
A A A A
30
B
B A
A
B B A A
20
10
0
Glucose
Glycerol
Mannitol
Trehalose
Compound
(B)
SN15
Mean % of Total Intensity
12
mdh1-71
mpd1-1
mpd1mdh1-107
A B AB AB
10
A A A A
8
6
A A A A
A A A A
4
A A A A
2
0
Alanine
Arabitol
Arginine
Glutamate
Glutamine
Compound
Figure 5.5: Mean relative abundance (±SE) of (A) major (>10%), and (B) minor
(<10%) soluble metabolites in extracts of strains of Stagonospora nodorum cultured
for 3 days in flasks with 40 mM glucose, as determined by
13
C NMR analysis.
Relative abundance was expressed in terms of the percentage of total
13
C intensity
for a spectrum above the sensitivity threshold, which accrued to the compound of
interest. Statistical significance was calculated for each group using the TukeyKramer HSD test, groups sharing a common letter were not significantly different.
N=3.
185
The five remaining identified metabolites, alanine, arabitol, arginine,
glutamate and glutamine, accounted for less than 10% each of total intensity (Figure
5.5B). The relative abundance of each was not significantly different between the
strains with the exception of arabitol, which was significantly more abundant in SN15
than in mdh1-71.
5.3.2.1.2 Replicates Inoculated from Minimal Medium Agar Cultures
The mpd1mdh1-107 cultures inoculated from minimal media agar plates
exhibited poor growth and their NA 13C-NMR spectra displayed a simpler metabolic
profile (Figure 5.6). The total intensity of one replicate was accounted for by glucose
(68.6%), glycerol (15.9%), alanine (6.1%), glutamate (1.6%) and unidentified peaks
(9.4%), while the second consisted of glucose (64.8%), glycerol (11.6%), alanine
(4.1%) glutamate (1.5%) and mannitol (1.1%) and unidentified peaks (16.9%). Apart
from glucose, glycerol and alanine, only a single peak was seen for the other assigned
compounds. All of the unidentified peaks were situated in the organic acids region of
the spectrum or in the aromatic/alkene region, and each accounted for less than 2% of
the total intensity.
5.3.2.2 [1-13C]-Glucose-Labelled Spectra
Based on the gross features of the spectra, the strains divided into two groups.
The first consisted of SN15 and mdh1-71, in which the most prominent peak was the
C1,6 of mannitol. A representative spectrum for SN15 is shown (Figure 5.7A). The
second group consisted of mpd1-1 and mpd1mdh1-107 in which the dominant peak
186
A
2000
1900
Gβ5
Gβ3
1800
Gβ1
1700
1600
1500
1400
1300
1200
1100
Gα1
1000
900
800
700
600
500
400
300
200
100
0
-100
100
99
98
97
96
95
94
93
92
91
90
89
88
87
f1 (ppm)
86
85
84
83
82
81
80
79
78
77
76
75
B
2000
1900
Gly1,3
Gβ5
Gβ3 Gβ2
1800
1700
Gβ4
Gβ6
1600
1500
1400
Gly2
1300
Gα2/5
Gα3
1200
Gα4
1100
Gα6
1000
900
800
700
600
500
400
300
200
100
0
-100
76
75
74
73
72
71
70
69
68
f1 (ppm)
67
66
65
64
63
62
61
C
1400
1300
1200
1100
1000
900
800
Ala3
700
600
Ala2
500
Glt3
400
300
200
100
0
-100
60
55
50
45
40
35
30
25
20
15
f1 (ppm)
Figure 5.6: Natural abundance
13C
NMR spectrum of mpd1mdh1-107 (inoculum sourced
from minimal medium agar plates) showing the regions from (A) 75-100 ppm, (B) 60-76 ppm
and (C) 15-60 ppm. Carbons have been assigned using the compound abbreviations
Ala=alanine; G=glucose; Glt=glutamate; Gly=glycerol. All chemical shifts were referenced
to the βC1 peak of glucose. The scales have been adjusted to the height of the most intense
peak in each section.
187
A
450
M1,6
400
350
300
250
200
150
Gβ1
100
M
Gα1
Gln4
?
50
Ala3
0
100
95
90
85
80
75
70
65
60
55
50
45
40
35
30
25
20
f 1 (ppm)
90000
B
85000
T1
80000
75000
70000
65000
60000
Gβ1
55000
50000
45000
40000
35000
Gα1
A1/5
Gly1,3 T6
30000
25000
20000
M
15000
Ala3
10000
Gln4
5000
0
100
95
90
85
80
75
70
65
60
55
50
45
40
35
30
25
20
15
f1 (ppm)
Figure 5.7:
13
C-NMR spectra of SN15 (A) and mpd1mdh1-107 (B) showing the
region from 15-100 ppm for cultures grown for 3 days on [1-13C]-glucose. Carbons
have been assigned using the compound abbreviations A=arabitol; Ala=alanine;
G=glucose;
Gln=glutamine;
Gly=glycerol;
M=mannitol;
T=trehalose;
?=unidentified. All chemical shifts were referenced to the βC1 peak of glucose. The
scales have been adjusted to the height of the most intense peak in each section.
188
was the C1 of trehalose. A representative spectrum for mpd1mdh1-107 is shown
(Figure 5.7B).
5.3.2.2.1 [1-13C]-Glucose
[1-13C]-labelled glucose was present in all samples. In SN15, only the labelled
anomeric C1 carbons were identified, and in one sample only a low amount of the β1
anomer was detected. The mean labelling of the C1 carbons in the other strains was
determined to be 91.1-fold ± 3.4 above NA (N=9). Spinning sidebands of the C1
carbons were present in most cases, and the β2, α2, β3, β6 and α6 carbon peaks were
split.
5.3.2.2.2 Mannitol
All three main mannitol peaks were assigned in all samples. In addition, split
C2 peaks were seen in all mdh1-71 samples as well as the SN15 sample in which
almost no glucose was detected. Assuming that labelling of only one terminal carbon
contributed to the C1,6 peak, the C1 had a mean fold-labelling above NA of 62.9 ±
13.1 (SN15), 68.6 ± 3.8 (mdh1-71), 40.8 ± 5.0 (mpd1-1), and was unlabelled in the
mpd1mdh1-107 samples (N≥ 3).
5.3.2.2.3 Trehalose
All trehalose peaks were assigned in all samples of the mpd1 strains. The C1
had a mean fold-labelling above NA of 32.7 ± 4.8 (mpd1-1) and 24.4 ± 1.1
189
(mpd1mdh1-107), while for the C6 this was 6.57 ± 0.9 and 5.02 ± 0.1 respectively
(N=3). One mdh1-71 replicate had a trehalose C1 peak, and one SN15 replicate had
C1 and C6 peaks.
5.3.2.2.4 Glycerol
Glycerol was detected in all samples except for one SN15 replicate. For SN15
and mdh1-71 only the C1,3 peak was assigned, which did not permit the amount of
labelling in this peak, if any, to be determined. Both glycerol peaks were assigned in
all samples of the other two strains, and the C1,3 peak had a mean fold-labelling
above NA of 7.4 ± 0.2 (mpd1-1) and 11.0 ± 0.5 (mpd1mdh1-107) (N=3).
5.3.2.2.5 Alanine
Alanine was present in all spectra except for the one SN15 sample in which
there was almost no glucose detected. In all other cases the C3 peak was present, and
for all remaining SN15 samples and one of the mdh1-71 samples, it was the only
alanine peak detected. The intensity of the peak and the absence of the sibling peaks
suggested that the C3 peak was labelled. In the remaining samples where sibling
peaks were present, it was possible to determine the degree of labelling in the C3
carbon. This was comparable between strains with a mean fold-increase above NA
labelling of 13.2 ± 0.8 (mdh1-71) (N=2), 16.5 ± 1.3 (mpd1-1) (N=3) and 16.0 ± 8.1
(mpd1mdh1-107) (N=3).
190
5.3.2.2.6 Glutamine
This compound was detected in all samples, but the full complement of peaks
was found in only one mdh1-71 spectrum. The C4 of SN15 and mdh1-71 had a mean
of 7.0 ± 1.4 fold labelling above NA (N=7), but this was only 1.0 ± 0.2 for the C4 of
mpd1-1 (N=3), and 1.3 for the only mpd1mhd1-107 sample for which this could be
determined. There was also labelling in the C2 carbon of SN15 and mdh1-71 of 1.9 ±
0.6 fold above NA (N=7), but no support for labelling of this carbon in the other
strains.
5.3.2.2.7 Glutamate
Glutamate was present in all samples apart from the SN15 sample in which
almost no glucose was detected. There was no indication of labelling in the compound
in any sample.
5.3.2.2.8 Arabitol
The mpd1mdh1-107 mutant was the only strain in which all carbons of arabitol
were detected in all samples. It was calculated that there was labelling in both the C5
(12.7 ± 0.5 fold above NA) and the C1 (8.6 ± 0.3 fold above NA) carbons (N=3). The
C5 peak in the other strains was obscured by a mannitol C1,6 sideband, apart from
one of the mpd1-1 samples. With the single exception of one SN15 sample in which
all peaks apart from C5 were identified, the only arabitol peak assigned in the samples
191
of the other strains was the C1. The C1 of the SN15 sample was labelled 1.5-fold
above NA.
5.3.2.2.9 Arginine
The full complement of arginine peaks was not present in any sample. No
arginine was detected in any mpd1mdh1-107 sample apart from one with a single C3
peak. There was good support in the mdh1-71 and mpd1-1 samples for mean foldlabelling of the C6 peak above NA of 2.3 ± 0.67 (N=3) and 3.7 ± 0.05 (N=2)
respectively. The data from SN15 was inconsistent with good match for C6 found in
only one replicate, with a calculated 11.8-fold labelling of the peak above NA.
5.3.2.3 [1-13C]-Mannitol-Labelled Spectra
5.3.2.3.1 Assay of mannitol uptake
The ability of strains to utilise [1-13C]-mannitol as a sole carbon substrate was
investigated. An assay of mannitol uptake by the strains was undertaken to optimise
usage of [1-13C]-mannitol. The mannitol standard curve demonstrated that there was a
linear relationship between mannitol concentration and net change in absorbance per
min at 340 nm due to mannitol oxidation, over the range of concentrations expected in
the medium (Figure 5.8A). The spent glucose media had no more activity than the
MM-C sample indicating that no mannitol was secreted by the strains (Figure 5.8B).
The 20 mM and 40 mM controls confirmed the accuracy of the assay and the
continued activity of the enzyme. After 24 h on 40 mM mannitol, the media aliquots
192
A: 0.00300
ΔA/min (340 nm)
0.00250
0.00200
0.00150
0.00100
y = 5E-05x
0.00050
2
R = 0.9927
0.00000
0
5
10
15
20
25
30
35
40
Mannitol (mM)
B:
50.0
Mannitol (mM)
40.0
30.0
20.0
10.0
0.0
1 DPI
2 DPI
20 mM
mannitol
40 mM
mannitol
MM-C
Spent MM
Enzyme reaction substrate
Figure 5.8:
A: Standard curve relating concentration of mannitol to net change in absorbance at 340 nm
due to the mannitol oxidation activity of mannitol dehydrogenase in a cell-free extract of
Stagonospora nodorum strain SN15. N≥ 3.
B: The concentration of mannitol in various samples and controls as determined by
observed mannitol oxidation activity in conjunction with the mannitol standard curve above.
1 DPI = mean mannitol concentration in aliquots of minimal media cultures (supplemented
with 40 mM mannitol and inoculated with 4 day old flask cultures of strains of S. nodorum)
taken at 24 hours post-inoculation; 2 DPI = as for 1 DPI but assay performed on aliquots
taken at 48 hours post inoculation; 20 mM & 40 mM mannitol = standard controls; MM-C
= minimal medium without carbon (medium control); Spent MM = aliquots taken from the
4 day old growth medium from which the fungal cultures were harvested prior to transfer to
the mannitol-supplemented media. N≥ 4.
193
contained a mean concentration of 20.0 ± 8.2 mM mannitol (N=12). This was further
reduced after 48 h incubation to a mean concentration of 11.0 ± 9.8 mM mannitol
(N=12). As a result, it was determined that 20 mM [1-13C]-mannitol would be
sufficient to meet the uptake requirements of all strains for the labelling experiment as
planned.
5.3.2.3.2 Gross features of spectra
The gross features of the spectra from the different strains were all similar
insofar as the C1,6 peak of mannitol was the most intense resonance peak. A
representative spectrum from each strain is shown (Figure 5.9). There were some
obvious differences between the strains in that the mpd1-1 and mpd1mdh1-107 strains
displayed few peaks other than mannitol, while the SN15 and mdh1-71 spectra had a
number of peaks in the sugar/polyol and organic acid regions.
5.3.2.3.3 Mannitol
All peaks of [1-13C]-mannitol were assigned in all samples, including C1,6
sidebands and split C2 peaks, and all with strong labelling above NA of the C1,6
peak. Assuming that only one terminal carbon was labelled, the mean fold-labelling
above NA of C1 was 80.0 ± 9.3 (SN15), and 53.84 ± 18.4 (mdh1-71) (N=3). The
other two strains additionally showed resolved C3 and C4 peaks, otherwise only seen
in the compound standard (Figure 5.2), and which were taken to be indicative of
100% labelling of the C1.
194
M1,6
Glt3
Gln4
M2,5 M3,4
T1
100
95
90
Figure 5.9:
85
13
80
75
70
65
Ala3
T6
60
f1 (ppm )
55
50
45
40
35
30
25
20
15
C-NMR spectra of Stagonospora nodorum strains SN15 (black),
mdh1-71 (red), mpd1-1 (green) and mpd1mdh1-107 (blue) grown for two days on
40 mM glucose followed by 24 h on 20 mM [1-13C]-mannitol. Each spectrum is
representative of three independent experiments. Peak heights have been
normalised to the intensities of the C1,6 peaks of mannitol. Assigned peaks in this
figure pertain to trehalose (T), mannitol (M), glutamine (Gln), glutamate (Glt) and
alanine (Ala) for the carbons as numbered.
195
5.3.2.3.4 Trehalose
Most trehalose peaks were detected in mdh1-71 samples, but in most other
instances only the C1 and C6 were assigned. In two of the mpd1mdh1-107 and one of
the mpd1-1 replicates, no trehalose was detected. In the mdh1-71 replicates, mean
fold-labelling of the C1 above NA was 8.0 ± 1.3, and of the C6 was 14.4 ± 5.3 (N=4).
5.3.2.3.5 Glucose
Glucose was only detected in one replicate each of SN15 and mdh1-71 and in
both cases it was only the C1 anomers and βC6 peaks which were seen.
5.3.2.3.6 Glycerol
The C1,3 peak of glycerol was assigned in all mdh1-71 and two SN15
replicates. No glycerol was detected in any of the mpd1 mutants.
5.3.2.3.7 Arabitol
The C5 of arabitol was obscured in all spectra. The mdh1-71 strain was the
only one in which arabitol was detected in all replicates. The C1 was not assigned in
any SN15 spectrum and labelling of other peaks was not suggested. The C1 of
arabitol in the mdh1-71 spectrum had a mean fold-labelling above NA of 2.6 ± 0.8
(N=4). Labelling of the C1 could only be determined in a single spectrum each of
mpd1-1 and mpd1mdh1-107 of 5.9-fold and 6.4-fold above NA respectively.
196
5.3.2.3.8 Amino acids
Alanine was detected in all SN15 and mdh1-71 spectra. Mean labelling of the
C3 in the mdh1-71spectra was 17.7 ± 3.6 fold above NA (N=3). It was only possible
to determine labelling in one SN15 replicate of 9.9-fold above NA. Alanine was
detected in one mdp1mdh1-107 and two mpd1-1 spectra. Only the C3 peak was
assigned, and its intensity was not suggestive of any labelling.
Most glutamate peaks were seen in all SN15 and mdh1-71 samples. There was
some circumstantial support for two- to three-fold labelling above NA of the C3 in
some spectra, but this was contradicted in other replicates. It was only spasmodically
detected in the other two strains.
Arginine peaks were seen in spectra of all strains, although they were absent in
several spectra. There was no indication of labelling of any peak in spectra where
such calculation was possible.
Glutamine was detected in all mdh1-71 replicates and most SN15 replicates
but spasmodic in the other two strains. There was no strong support for labelling
above NA in any carbon.
197
5.3.2.4 [1-13C]-Glucose Feed-Chase Spectra
The SN15 strain was investigated for the in vitro metabolic fate of labelled
carbon in a feed-chase experiment. The ‘feed’ phase involved growing cultures on [113
C]-glucose, with cultures transferred to unlabelled glucose during the ‘chase’ phase.
This technique is used in
13
C-NMR to identify the pathways and products of
metabolism, and to indicate which metabolite pools are active. Four replicates were
prepared for each time-point sampled. However, one of the 1h feed cultures was
found to be contaminated at harvest and discarded. A representative spectrum from
each time-point is shown (Figure 5.10).
The calculated mean fold-labelling above NA of carbons for which this could
be determined is shown (Figure 5.11).
5.3.2.4.1 Carbohydrates
Mannitol was present in all samples. The three main peaks were assigned in all
cases, and the split C2 peaks and C1,6 sidebands were assigned in the majority of
cases. Assuming that the increased intensity of the C1,6 peak was due to labelling in
one terminal carbon only, there was mean labelling above NA after 1 h on labelled
glucose substrate of 24.3-fold ± 2.6 (N=3), rising to 38.4-fold ± 2.9 (N=4) after four
hours. One hour after transfer to unlabelled glucose this fell to 19.6-fold ± 4.1 (N=4),
and 7.0-fold ± 0.1 after four hours (N=4).
198
M1,6
4 h chase
Gβ1
1 h chase
Ala3
4 h feed
M2,5 M3,4
1 h feed
100
95
90
85
80
75
70
65
60
f1 (ppm )
55
50
45
40
35
30
25
20
15
Figure 5.10: 13C-NMR spectra of SN15 cultures from a feed-chase experiment for
the range 15-100 ppm. Each spectrum is representative of at least three independent
experiments. Peak heights were been normalised to the natural abundance
intensities of the C2,5 and C3,4 peaks of mannitol. Cultures represent the 13C label
present in spectra after 1 h feed and 4 h feed on 40 mM [1-13C]-glucose and
followed by 1h and 4 h chase on 40 mM unlabelled glucose. Assigned peaks in this
figure pertain to glucose (G), mannitol (M) and alanine (Ala) for the carbons as
numbered.
199
Mean fold-increase of 13C-labelling
above natural abundance
1h Feed
4h Feed
1h Chase
4h Chase
100
90
80
70
60
50
40
30
20
10
0
Alanine C3
Glucose βC1
Glucose αC1
Glutamine C3
Glycerol C1,3
Mannitol C1
Trehalose C1
Trehalose C6
Compound and number of labelled 13C resonance peak
Figure 5.11: Changes in mean (±SE) fold labelling above
13
C natural abundance
for selected compounds over the course of a feed-chase experiment. Three day old
SN15 cultures were transferred to 40 mM [1-13C]-glucose. Half of the cultures
were harvested after 1 or 4 hours incubation on the labelled medium and the
remainder transferred to unlabelled 40 mM glucose and harvested after 1 or 4 hours
incubation on the unlabelled medium. N ≥ 3.
200
Glucose was present in all samples. Mean fold-labelling above NA of the βC1
and αC1 carbons was, respectively, 87.4 ± 6.4 and 89.8 ± 5.7 (N=3) after the 1h feed,
92.2 ± 5.1 and 93.4 ± 4.3 (N=4) (4h feed), dropping back to 1.8 ± 0.5 and 1.9 ± 0.6
(N=4) (1h chase) and 1.5 ± 0.6 and 1.6 ± 0.6 (N=4) (4h chase).
Glycerol was detected in all samples, although the lower-intensity C2 carbon
was not assigned in one spectrum each from each time period. There was no apparent
labelling for the 1h feed or 4h chase samples, but the data supported 2.2 ± 0.1 (N=3)fold labelling above NA of the C1,3 peak for the 4h feed, and 1.6 ± 1.0 (N=3)-fold
labelling for the 1h chase samples.
Trehalose peaks were assigned in all samples, but there was no instance where
all six peaks were assigned in a single sample. The C1 peak was the most intense peak
in the NA compound standard, and this was the only peak assigned in all samples. In
four spectra, the only trehalose peak assigned was the C1 peak. While this is good
prima facie evidence for its being heavily labelled, it was not possible to calculate its
RPI where it was the only peak detected. For the spectra where C1 labelling could be
calculated, the pattern was inconsistent. In the 1h feed samples it ranged from 0.5 to
4.8-fold above NA. In the 4h feed samples it ranged from a mean of 4.9-fold (±0.5) in
two samples to a mean of 28.2-fold (±7.5) in another two samples. In the 1h chase,
there were only two samples where the RPI of C1 could be calculated, and these had a
mean fold-increase of 22.3 ± 5.3 above NA. The 4h chase samples were the most
consistent with a mean fold-increase of 7.9 ±1.1 above NA for the three samples
where this could be determined. There was also the indication of labelling on the C6
of trehalose. This only occurred after the 1h feed and showed great variation after the
201
4h feed with a mean of 16.6 ±12.9 (N=3). It was more consistent during the chase
phase with a mean fold-labelling above NA of 4.7 ±1.0 (N=2) after 1h and 2.9 ±0.4
(N=3) after 4 h.
The C5 of arabitol could not be unambiguously identified as it was co-located
with one of the sidebands of the labelled mannitol C1,6 peak. All remaining arabitol
peaks were identified in all samples with the exception of the C1, which was not
assigned in two of the 1h feed samples, and one of the 4h chase samples. There was
no strong evidence for labelling above NA in any of the assigned arabitol peaks.
5.3.2.4.2 Amino acids
All peaks of alanine were assigned in all samples with the exception of the low
intensity C1 carbon, which was missing in one replicate each of the 1h feed and 4h
feed spectra. The mean fold-labelling above NA in the C3 carbon was 6.8 ± 1.4 after
the 1h feed, 10.3 ± 0.9 (4h feed), 3.4 ± 0.8 (1h chase), and 2.0 ± 0.3 (4h chase) (N≥
3).
Glutamine was detected in all samples, although the low-field low-RPI C1 and
C5 peaks were not assigned in all cases. The RPI of the C4 carbon was consistently
elevated in the feed samples relative to the chase samples and there was support for
mean fold-labelling above NA in this carbon of 1.20 ± 0.1 for the feed phase of the
experiment (N=7).
202
The full complement of glutamate peaks was found in all samples and there
was no indication of labelling.
The full complement of arginine peaks was not present in any sample, and the
compound was not detected in almost half of the samples. In those samples where it
was present, there was no indication of labelling.
5.3.3 Miscellaneous Peaks
There were 2,977 peaks across all spectra which were not assigned to any
compound with confidence. Of these, 139 represented more than 1% each of the total
intensity of the spectrum in which they were found. Forty-two of these peaks could be
grouped into 12 clusters of peaks which were within a range of 0.10 ppm (Table 5.3).
The majority of these clusters did not have a good match in the Compound Standard
Library. The clustering process was extended to include all peaks in the sugar/polyol
and organic acid regions of the spectrum (0-100 ppm). This resulted in 1766 peaks
being grouped into 146 clusters, each comprising peaks from 3-43 spectra within a
range of 0.12 ppm (Table 5.4). Twenty-eight of these clusters had good matches from
the Compound Standard Library. However, these were not always unambiguous, and
there was insufficient support from sibling peaks to justify identification of a
compound in any one spectrum.
Nine of the clusters were comprised of spectra from only one strain and were
all located in the organic acids region. Eight of these clusters were specific to SN15.
203
Table: 5.3: Peak clusters from 13C-NMR spectra of strains of Stagonospora nodorum
for peaks comprising >1% of total intensity. The range for each cluster, the number of
spectra comprising each cluster, the strains and treatments (including the number of
replicates), and the best match for the cluster from the ACNFP Compound Standard
Library are shown.
Cluster Range
(ppm)
85.01
No.
6
63.29-63.32
4
59.83-59.88
4
53.90-53.91
3
53.84-53.85
6
53.80
3
53.75
2
29.62-29.63
3
28.86-28.92
26.29
2
4
25.28-25.29
23.63-23.69
2
4
Strain/Treatment*
+ (no. of replicates)
SN15-13G (5);
mpd1-1-13G (1)
SN15-13G (1);
mpd1mdh1-107-13G (3)
SN15-13G (3);
mpd1-1-13G (1)
SN15-13G (1);
mdh1-1-71-NA (1);
mdh1-1-71-13M (1)
SN15-13G (2);
SN15-13M (1);
mdh1-1-71-13M (2);
mpd1mdh1-107-NA (1)
SN15-13G (1);
SN15-13M (1);
mpd1mdh1-107-NA (1)
SN15-13G (1);
SN15-13M (1)
SN15-13M (1);
mdh1-1-71-13M (2)
mdh1-1-71-13M (2)
SN15-13G (3);
SN15-13G (1)
mdh1-1-71-13M (2)
mdh1-1-71-13M (3);
mpd1mdh1-107-NA (1)
Best Library Match
None
Fructose βC1/
Mannitol 1-phosphate
None
Methionine αCH
None
None
None
Methionine βCH2
None
None
None
None
* NA = natural abundance spectrum from growth on unlabelled glucose;
spectrum from growth on [1-13C]-glucose;
13
G –
13
M – spectrum from growth on [1-13C]-
mannitol.
204
Table 5.4: Distribution of unidentified peaks from
13
C-NMR spectra of strains of
Stagonospora nodorum into clusters.
No. of Spectra per Cluster
No. of Clusters
> 40
4
30-39
6
20-29
10
10-19
59
3-9
67
Total
146
205
Three of the SN15 peaks came from natural abundance spectra while the remainder
were from labelled spectra. None of the SN15 clusters had a match in the compound
standard library. One cluster was specific to mdh1-71 and comprised 3 peaks from [113
C]-mannitol labelled spectra and was a good match for asparagine. No peak
accounted for more than 0.3% of total intensity. A further 21 clusters were only found
in strains with an intact Mpd1 gene. Seven of these were located in the sugar/polyol
region of the spectrum with one unsupported match for the αC1 of mannose. Of the
peaks in the organic acids region, there was one unsupported match for the C2 of
threonine, and another for the βCH3 of pyruvate. There were no clusters which were
specific to either or both of the mpd1 mutants.
There were a number of unidentified peaks which appeared to be labelled
based on their intensity. A peak at 85.01 ppm was only noted in spectra of cultures
grown on
13
C-labelled medium, and accounted for 1.6-6.4% of total intensity in 5
SN15 spectra and 1 mpd1-1 spectrum. It was present in a further 7 SN15 and 2 mdh171 spectra at lower intensities, and was not observed in any mpd1mdh1-107 spectrum.
A peak at 62.31 ppm appeared to be labelled in mpd1mdh1-107 cultures
grown on labelled glucose. This peak accounted for 2.4-4.4% of total intensity in all
replicates and was present at much lower intensity, or undetectable in natural
abundance spectra. It was also seen in several other SN15 and mdh1-71 spectra,
including labelled spectra, but accounted for at least an order of magnitude less of
total intensity than was the case for mpd1mdh1-107. The best match for this peak was
206
either the C1 of sorbitol or the αC6 of fructose, but neither match was supported by
associated peaks.
There were a number of peaks in the range 58.70-60.80 which accounted
greater than 1% of total intensity in spectra of SN15 and mdh1-71. They did not
justify grouping into clusters, and other strains were represented in the same region at
lower intensities. Some presented good matches to the C6 of galactose and mannose
or the C2 of threonine, but without good support for this identification.
The two clusters matching methionine from Table 5.3 had additional, lower
intensity peaks from all strains. Their identity was not supported by the absence of the
sulphonyl carbon in natural abundance spectra, and inconsistent RPI pattern of the
two tentatively identified peaks.
The cluster which appeared in most spectra (43) was located at 22.01 ppm and
had no match.
5.4 DISCUSSION
5.4.1 Disruption of Mpd1 Alters the Metabolite Profile
13
C natural abundance NMR was used to characterise differences in the
metabolite profiles of the Stagonospora nodorum wild type strain SN15, and three
mutant strains with disrupted mannitol metabolism genes. On average, over 80% of
207
the observable 13C in the spectra for all cultures started with inoculum from CZV8CS
agar plates, was accounted for by nine metabolites. These were divided into two
groups comprising the major metabolites (accounting for a mean percentage of total
intensity of >10% for the spectra of any strain) and minor metabolites (accounting for
a mean percentage of total intensity of no more than 10% for the spectra of any
strain).
The NA spectra for the mpd1mdh1-107 cultures started with inoculum from
minimal medium agar plates were designed to eliminate exogenously accumulated
mannitol as a factor in these spectra. While this was achieved, the growth was much
reduced compared to the same strain on the more complex medium. The presence of
glycerol but not trehalose in these spectra may have some implications regarding the
relative importance of the role(s) of mannitol for which these compounds appear to be
compensating. While this is not uninteresting, these spectra were not strictly
comparable with the other NA spectra. The poor growth and largely uninformative
metabolite profile of these cultures led to this avenue of investigation being
discontinued. All remaining discussion refers to the CZV8CS-derived cultures only.
5.4.1.1 Mannitol
The strains could be divided into two statistically significant groups on the
basis of the major metabolites present. Mannitol was the most abundant metabolite in
strains with an intact Mpd1 gene, encoding mannitol 1-phosphate dehydrogenase
(SN15 and mdh1-71), while this compound was present at an order of magnitude less
in the mpd1 mutant strains (mpd1-1 and mpd1mdh1-107). This is consistent with a
208
previous
13
C-NMR natural abundance (NA) study involving SN15 and mpd1-1 in S.
nodorum (Solomon et al., 2005a). It was demonstrated in Chapter 3 that the double
mutant strain is unable to synthesise or catabolise mannitol, but can accumulate it
from its environment and maintain a stable pool. The detection of low levels of
mannitol in all but one of the mpd1mdh1-107 NA samples implied that this was
carried over with the inoculum. The labelling component of this experiment further
confirmed this mannitol accumulation behaviour. In the [1-13C]-glucose cultures, the
mpd1mdh1-107 mannitol pool accounted for less than 1% of total intensity in all cases
and was unlabelled, while in the [1-13C]-mannitol cultures, this pool accounted for up
to 99% of total intensity and was essentially 100% labelled. This accumulation
behaviour is not universal for fungi. Germinating spores of the arbuscular mycorrhizal
fungus Glomus intraradices, did not take up exogenous mannitol under conditions of
asymbiotic growth (Bago et al., 1999).
In the case of the mpd1-1 strain, growth on [1-13C]-glucose indicated that
mannitol synthesis was still possible via the mannitol dehydrogenase (Mdh1)
pathway, since the C1,6 of mannitol in the strains was labelled 40-fold above NA
levels. This confirms the conclusion in Chapter 3 above, that this strain was capable
of synthesising mannitol, since it was able to sporulate in vitro when serially
subcultured onto minimal medium from which exogenous mannitol was absent. The
strain was also capable of taking up mannitol from the media as evidenced by growth
on [1-13C]-mannitol, with mannitol accounting for up to 99% of total intensity and
being essentially 100% labelled.
209
5.4.1.2 Trehalose and Glycerol
The main metabolites in the mpd1 mutants were trehalose and glycerol, with
both of these being either undetectable, or representing less than 2% of total intensity,
in the SN15 and mdh1-71 spectra. The virtual replacement of mannitol with trehalose
and glycerol suggests an alteration in metabolism to compensate for the loss of the
ability to accumulate mannitol to wild type levels. Evidently these compounds were
unable to repair the inability to sporulate in vitro of the double mutant. A number of
the postulated roles of these compounds overlap, but not all have been conclusively
proven, and they vary in distribution and abundance between species. Targeted gene
deletion mutants are elucidating these roles in fungi.
Trehalose 6-phosphate synthase (Tps1) has been knocked out in a number of
fungi including S. nodorum. Some mutants have been more susceptible to
environmental stresses including heat (Botrytis cinerea and S. nodorum (Doehlemann
et al., 2006; Lowe, 2006), but not Magnaporthe grisea (Foster et al., 2003)), and
oxidative stress (Aspergillus nidulans, Candida albicans and S. nodorum (Fillinger et
al., 2001; Gonzalez-Parraga et al., 2003; Lowe, 2006; Martinez-Esparza et al., 2007)).
Mutants have also been shown to be affected in pathogenicity (C. albicans,
Cryptococcus neoformans, M. grisea, and S. nodorum (Zaragoza et al., 1998; Foster
et al., 2003; Lowe, 2006; Petzold et al., 2006; Wilson et al., 2007), but not B. cinerea
(Doehlemann et al., 2006)). A more general role for trehalose, ascribed in a wide
range of fungal species, is as a storage carbohydrate (Thevelein, 1984; Bécard et al.,
1991).
210
Glycerol is the main compatible solute in the halophilic yeast Hortaea
werneckii and while erythritol, arabitol and mannitol are all present at optimal growth
salinities, only erythritol and glycerol are present during growth on 25% (w/v) NaCl
(Kogej et al., 2007). The filamentous fungus Cladosporium fulvum accumulated
glycerol and arabitol, but not mannitol, in response to osmotic stress (Clark et al.,
2003). Aspergillus nidulans strains with a disrupted NADP+-dependent glycerol
dehydrogenase gene, had strongly decreased levels of glycerol and elevated levels of
arabitol, erythritol and mannitol, and exhibited reduced growth on 1 M NaCl (de
Vries et al., 2003). Exogenous supplementation with any of these polyols corrected
for the deficiency, but uptake of all except glycerol was subject to glucose repression.
Osmotic stress resulted in significantly increased arabitol levels in a xylitol
dehydrogenase mutant (xhd1) of S. nodorum, while glycerol was significantly
increased in an L-arabitol dehydrogenase mutant (abd1), and a double mutant strain
(abd1xdh1) in which both of these genes had been disrupted (Lowe et al., 2008).
Glycerol has also been demonstrated to have roles in fungal phytopathogenicity as the
source of the tremendous turgor pressure in appressoria of M. grisea (Dixon et al.,
1999), and as a nutrient transferred from the host by Colletotrichum gloeosporioides
f.sp. malvae (Wei et al., 2004). Glycerol was shown to accumulate in sunflower
cotyledons when infected with Sclerotinia sclerotiorum, while uninfected cotyledons
and the in vitro-cultivated fungus had no detectable glycerol (Jobic et al., 2007).
It might be expected that fungal species in which no Mpd1 activity is present,
would show a similar metabolite profile to the S. nodorum mpd1 mutants.
Investigations into the enzymes of mannitol metabolism have demonstrated an
absence of Mpd1 activity in the majority of basidiomycetes for which this enzyme
211
was assayed, although mannitol-1-phosphate phosphatase activity was detected in
nearly half of these (Table 1.3). In the case of Agaricus bisporus, neither enzyme was
detected (Hult et al., 1980) which suggests that only the Mdh1 pathway is active in
this species. While only low levels of mannitol were detected in the mycelium of A.
bisporus, it was the main soluble metabolite in the sporophore and was accumulated
as the sporophore developed (Hammond and Nichols, 1976), but decreased postharvest (Donker and Braaksma, 1997). Trehalose was the second most abundant
metabolite, but levels decreased in both mycelium and sporophore as the latter
developed (Hammond and Nichols, 1976), and it could not be quantified reliably in
13
C-NMR NA spectra post-harvest as most peaks were below the detection threshold
(Donker and Braaksma, 1997). Glycerol was not noted as a significant metabolite in
these studies. It is therefore apparent that the increase in trehalose and glycerol seen in
the S. nodorum mpd1 mutants is not universally observed in organisms in which only
the Mdh1 pathway is active.
The fact that two compounds were accumulated in S. nodorum in response to
the abolition of mannitol synthesis, implies that there are at least two separate roles of
mannitol for which compensation is being made. Further, given that the defect in in
vitro sporulation of the mpd1mdh1-107 strain was not repaired by this altered
metabolome, there is circumstantial evidence for three roles of mannitol. This could
be considered an efficient use of a normally abundant resource. While it may be
tempting to consider that glycerol and trehalose are performing the roles of
compatible solute and carbohydrate store respectively, this is speculation without
specific manipulation of the experimental system to induce conditions which would
confirm or refute this. These roles have also not otherwise been conclusively
212
demonstrated for mannitol. The observation of these differences provides an
opportunity to explore the roles of these compounds further. The demonstrated ability
of S. nodorum gene disruption constructs to abolish either trehalose (Lowe, 2006) or
mannitol synthesis (this study), and the implication that trehalose replaces the
function of mannitol at least in part, suggests the possibility of combining these
constructs in a mutant strain. The production of a triple mutant in this fungus is not a
trivial matter, however, and attempts to produce a double mutant harbouring the mpd1
and tps1 constructs have thus far proven unsuccessful (Dr. P. Solomon, pers. comm.).
While this is suggestive of a lethal condition, this is speculative at this stage.
5.4.1.3 Glucose
This compound occupies an ambiguous position in the NA and [1-13C]glucose experiments as both the substrate, and as a metabolite. It has previously been
detected in 13C natural abundance NMR spectra (Dijkema and Visser, 1987; Solomon
et al., 2005a; Jobic et al., 2007), but there is little discussion as to whether it is the
result of poorly washed mycelium, substrate which has been taken up but not yet
metabolised, the result of de novo synthesis via gluconeogenesis, trehalose
degradation etc., or some combination of these. It is difficult to justify a claim for any
of these scenaria for growth on unlabelled glucose. The fact that one of the SN15
samples had no detectable glucose could be suggestive of variable efficiency in the
washing technique. However, this was also the sample with the heaviest extract dry
weight of 52.2 mg, more than twice that of all other samples bar one. It is more likely
that this sample represented a culture which had exhausted the glucose in the medium.
The argument is not trivial, since the degree of labelling of the C1 and C6 of glucose
213
following growth on [1-13C]-glucose, was used in several studies to estimate the flux
of carbon through pathways which cause such scrambling.
The observed pattern of
13
C labelling of glucose in the SN15 feed-chase
experiment offers some insight into the capacity of mycelium to take up carbon from
their environment. An extreme condition must first be considered where the mycelium
has virtually no capacity for glucose storage and that [1-13C]-glucose taken up is
metabolised immediately. In this instance one would expect to see no labelled glucose
in the spectrum of a properly-washed sample, while in an insufficiently washed
sample, any glucose present would have 100% labelling of the C1 carbons. However,
the mean fold-labelling above NA of the βC1 anomer of glucose in the spectra was
87.4 ± 6.4 after 1h on glucose and 92.2 ± 5.1 after 4 hours (N≥ 3). This is consistent
with the addition of [1-13C]-glucose to an initially unlabelled pool of accumulated
glucose and with the proportion of labelled substrate increasing with time of exposure
to media containing it. Similarly, in samples which were subsequently transferred to
medium containing unlabelled glucose, the mean fold-labelling above NA of the βC1
anomer of glucose in the spectra was 1.8 ± 0.5 after 1 h and 1.5 ± 0.6 after 4 h (N=4).
This is consistent with the ongoing dilution of label by uptake of unlabelled glucose.
If the majority of observed label had been due to poor washing, then it would be
expected that the differences in labelling between the 1 h and 4 h samples would be
reversed, since the fungus would have reduced the amount of carbon in the medium in
that time. The fact that the amount of labelling approached saturation on the labelled
medium, followed by its almost complete disappearance on the unlabelled medium,
provided evidence that the glucose pool is a transient one with a rapid turnover.
214
5.4.1.4 Arabitol and Amino Acids
The remaining minor metabolites of the spectra were largely unremarkable in
that the amino acids have been identified in 13C-NMR studies in other fungal species
(Donker and Braaksma, 1997; Ceccaroli et al., 2003; Jobic et al., 2007), and were not
present in significantly different amounts between the different S. nodorum strains.
The one exception to this was arabitol, where the NA spectra showed SN15 as
having significantly more of this compound than mdh1-71. The ability of all strains to
produce this compound, however, was confirmed by the labelled spectra in which the
C1 peak was routinely observed. The distribution of arabitol as determined by
13
C-
NMR studies of other filamentous fungal species indicated that this polyol was often
absent or below the threshold of detection (Martin et al., 1984; Yoshida et al., 1984;
Martin et al., 1985; Martin et al., 1988; Bécard et al., 1991; Donker and Braaksma,
1997; Peksel et al., 2002; Ceccaroli et al., 2003; Jobic et al., 2007). In species where
it has been detected, only the more intense peaks have been seen, suggesting that it is
of relatively low abundance (Dijkema et al., 1985; Martin et al., 1998; Rangel-Castro
et al., 2002; Clark et al., 2003).
The absence, in the labelled spectra of most replicates, of sibling peaks of
arabitol, suggests that the compound was not present in large amounts, and inferred
that the observed C1 peak was labelled. It has recently been shown that arabitol is
accumulated in response to osmotic stress in S. nodorum (Lowe et al., 2008). For
those spectra in which labelled mannitol was abundant, the C5 peak of arabitol could
not be unambiguously assigned. However, the spectra of mpd1mdh1-107 replicates
215
grown on [1-13C]-glucose demonstrated that both the C1 and C5 of arabitol were
labelled in a 1 (C1):1.5 (C5): ratio. The fact that this strain is unique in its inability to
synthesise labelled mannitol from labelled glucose has enabled this arabitol labelling
pattern to be observed for the first time in fungi. Previous studies have apparently
been unable to discriminate between the C1 and C5 of arabitol (Dijkema et al., 1985;
Martin et al., 1998; Rangel-Castro et al., 2002).The exception to this was the NA 13CNMR study of Cladosporium fulvum which did not involve a labelled substrate (Clark
et al., 2003). The mechanism and significance of this labelling pattern is discussed
further below (Section 5.4.3.4).
5.4.2 No Third Pathway of Mannitol Metabolism Detected in S. nodorum
The synthesis of [1-13C]-mannitol from [1-13C]-glucose by both of the single
mutants but not by the double mutant demonstrated that both metabolic pathways are
capable of mannitol synthesis, and also that there is no alternative anabolic pathway
for mannitol under these conditions.
The synthesis of [1-13C]-trehalose, [6-13C]-trehalose and [3-13C]-alanine from
[1-13C=6-13C]-mannitol by the mdh1-71 strain demonstrated firstly that this strain is
capable of mannitol catabolism as was concluded in Chapter 3. Secondly, it confirmed
that there is an as yet unidentified component of the Mpd1 pathway by which
mannitol can be phosphorylated to mannitol 1-phosphate. The inability of the mpd1-1
mutant to catabolise mannitol indicated that at least one of the enzymatic steps
catalysing the conversion of mannitol to fructose 6-phosphate by the Mdh1 pathway is
under tight physiological control. Since there is no evidence of a build-up of fructose,
216
whilst mannitol is observed to accumulate, it is most likely that this control is applied
to the step converting mannitol to fructose. It was demonstrated in the enzyme assays
in Chapter 3 that the Mdh1 enzyme has the ability to oxidise mannitol in a desalted
CFE. The inference is that the mannitol synthesised and/or accumulated by this strain,
is in some way compartmentalised such that the Mdh1 enzyme does not have access
to it, or that the catabolic reaction is subject to some form of inhibition. The inability
of the mpd1mdh1-107 mutant to catabolise mannitol indicated that there is no
alternative catabolic pathway for mannitol.
5.4.3 Scrambling of Label is not Proof of a Mannitol Cycle
Label scrambling has been previously seen in a number of studies and has
generally been explained by, and given as evidence of, an operational mannitol cycle
(Martin et al., 1988; Ramstedt et al., 1989; Pfeffer and Shachar-Hill, 1996). This has
not taken into account the possibility that a single pathway capable of both mannitol
synthesis and catabolism would be sufficient to contribute to label scrambling.
Alternative scrambling mechanisms have been suggested including the pentose
phosphate pathway (in both forward and reverse flux), and the aldolase/
triosephosphate isomerase triangle (den Hollander and Shulman, 1983; Portais and
Delort, 2002).
In the mannitol scrambling model [1-13C]-glucose is metabolised to [1-13C=613
C]-mannitol. Subsequent catabolism of mannitol, and passage to glucose 6-
phosphate via gluconeogenesis, results in the synthesis of trehalose and glucose which
are labelled on both the C1 and C6 carbons. The simplest version of this model would
217
result in equal distribution of label on the C1 and C6 of trehalose. Studies have
invariably shown that this is not the case, with reported ratios of 13C6 to 13C1 of 0.7
(Martin et al., 1985), 0.8 (Martin et al., 1988) or lower (Ramstedt et al., 1989). This
has been explained as being due to a proportion of trehalose being synthesised
directly from [1-13C]-glucose, resulting in enrichment of the label in the C1 of
trehalose (Peksel et al., 2002). In light of the conclusion of Chapter 3 that the
mannitol cycle does not exist as proposed in S. nodorum, this model requires some
revision.
5.4.3.1 The Mdh1 Pathway does not Contribute to Label Scrambling
The first observation to consider is that the double mutant strain mpd1mdh1107, when grown on [1-13C]-glucose, produced trehalose which was labelled on both
the C1 and C6 carbons and with a
13
C6/13C1 ratio of 0.21 ± 0.01 (N=3). This
demonstrated that label scrambling occurred in this strain. However, there was no
labelling of mannitol in these samples, and when cultured on [1-13C]-mannitol, there
was no labelling of trehalose or any other compound. It is apparent, therefore, that
scrambling is not solely due to cycling of carbon via mannitol, and that alternative
scrambling pathways are operational in S. nodorum.
Secondly, the mpd1-1 strain, in which the Mdh1 pathway was operational,
exhibited a similar labelling pattern to the double mutant. On [1-13C]-glucose it
synthesised trehalose with a 13C6/13C1 ratio of 0.20 ± 0.00, and mannitol with a foldlabelling of 40.8 ± 4.9 above NA (N=3). On [1-13C]-mannitol, there was no labelling
of trehalose or any other compound. The almost identical labelling pattern of the two
218
mpd1 strains, and the demonstrated inability of the mpd1-1 mutant to catabolise
mannitol, indicates that the Mdh1 metabolic spur does not contribute to the
scrambling observed in the mpd1-1 strain.
Thirdly, while there was virtually no trehalose detected in the mdh1-71 strain
when grown on [1-13C]-glucose, the [1-13C]-mannitol grown samples all had
significant amounts of labelled trehalose with a 13C6/13C1 ratio of 3.47 ± 1.74 (N=4).
This suggests that the pathway(s) by which mannitol is metabolised in this strain,
result in a trehalose scrambling pattern which is the inverse of that experimentally
observed in other species, and in the mpd1 mutants of S. nodorum.
The two alternative scrambling pathways proposed by den Hollander and
Shulman (1983) would therefore appear to be of greater importance in the mechanism
of scrambling than has previously been supposed.
5.4.3.2 The Aldose/Triosephosphate Isomerase Triangle
The aldolase/triosephosphate isomerase (TPI) triangle follows the metabolism
of [1-13C]-glucose to [1-13C]-fructose-1,6-bisphosphate (FBP) via glycolysis (Figure
5.12). FBP aldolase reversibly cleaves FBP to [1-13C]-dihydroxy acetone phosphate
(DHAP) and glyceraldehyde 3-phosphate. These two intermediates of glycolysis are
readily interconvertible via TPI, resulting in the production of [3-13C]-glyceraldehyde
3-phosphate. Under conditions of high FBP aldolase activity, a population of FBP
molecules can be generated which may in theory be labelled on the C1, C6, both
terminal carbons or neither (den Hollander and Shulman, 1983). These can then be
219
1 2 3 4 5 6
1 2 3 4 5 6
1 2 3 4 5 6
1 2 3 4 5 6
Trehalose
1 2 3 4 5 6
[1-13C]-Glucose
Hexokinase
1 2 3 4 5 6
Glucose 6-phosphate
Tps1/Tpp1
1 2 3 4 5 6
1 2 3 4 5 6
1 2 3 4 5 6
1 2 3 4 5 6
Glucose 6-phosphate
Phosphoglucose
isomerase
Phosphoglucose
isomerase
Fructose 6-phosphate
1 2 3 4 5 6
Fructose 6-phosphate
Phosphofructokinase
1 2 3 4 5 6
Fructose 1,6-bisphosphate
Fructose 1,6bisphosphatase
1 2 3 4 5 6
1 2 3 4 5 6
1 2 3 4 5 6
1 2 3 4 5 6
Fructose 1,6-bisphosphate
Aldolase
1 2 3
DHAP
Aldolase
1 2 3
G3P
1 2 3
1 2 3
DHAP
1 2 3
1 2 3
G3P
Triosephosphate
isomerase
Glycolysis
Figure 5.12: Aldolase/triosephosphate isomerase triangle mechanism for
scrambling from [1-13C]-glucose to [1-13C]/[6-13C] trehalose.
13
13
C label
C-labelled carbons
are shown in red. Metabolites are shown in boxes. Enzymes are shown in italics.
Abbreviations: DHAP: dihydroxy acetone phosphate; G3P: glyceraldehyde 3phosphate; Tps1: trehalose 6-phosphate synthase; Tpp1: trehalose 6-phosphate
phosphatase.
220
converted via the gluconeogenic pathway back to glucose 6-phosphate and thence to
trehalose. The 13C6/13C1 labelling ratio of trehalose would be expected to reflect that
of the glucose 6-phosphate pool from which it was formed. No glucose 6-phosphate
was positively detected in any of the labelled or NA spectra. This was not surprising
since intermediates of active metabolism are not often seen, and this has been
attributed to their low abundance and short lifetime (Yoshida et al 1984). However,
the aldolase/TPI triangle would be expected to contribute equimolar amounts of
13
C1/13C6-labelled glucose 6-phosphate, while the passage of the labelled glucose
substrate via glucose 6-phosphate would contribute to the
13
C1-labelled pool only.
This is consistent with the trehalose labelling pattern observed for the mpd1 mutants,
but does not account for the trehalose labelling pattern of the mdh1-71 strain grown
on [1-13C]-mannitol.
5.4.3.3 The Pentose Phosphate Pathway (Forward Flux)
The observed formation of trehalose in the mdh1-71 strain grown on [1-13C]mannitol, requires the conversion of mannitol to glucose 6-phosphate (G6P) via
gluconeogenesis (Figure 5.13). In the absence of any other contributory pathway, the
symmetry of the mannitol molecule would result in trehalose, and all intermediates,
with equimolar
13
C1- and
13
C6-labelled carbon atoms. The
13
C6/13C1 ratio of
trehalose in these samples indicated that a scrambling mechanism must exist whereby
labelling of the C6 is enriched to a greater extent than that of the C1. G6P can enter a
number of metabolic pathways including that of trehalose biosynthesis, the pentose
phosphate pathway (PPP), gluconeogenesis, glycolysis, and glycogen synthesis
(Stryer, 1997). For the labelling pattern observed in trehalose to have occurred,
221
Figure 5.13 (overleaf): Pentose phosphate pathway mechanism for
13C
label
scrambling from [1-13C]-mannitol to [1-13C]/[6-13C] trehalose. 13C-labelled carbons
are shown in red. Metabolites are shown in boxes. Enzymes are shown in italics.
Abbreviations: MPP: mannitol 1-phosphate phosphatase; MPD: mannitol 1phosphate
h h t dehydrogenase;
d h d
PGI phosphoglucose
PGI:
h h l
i
isomerase;
G6PDH glucose
G6PDH:
l
6
6phosphate dehydrogenase; 6PGDH: 6-phosphogluconate dehydrogenase; TPS:
trehalose 6-phosphate synthase; TPP: trehalose 6-phosphate phosphatase.
Enzymatic cleavage sites are indicated by a dotted line. Boxes of metabolites
occurring more than once are given a matching colour. Co-factors are not shown.
NB: For simplicity, the pathway from mannitol to fructose 6-phosphate via
mannitol 1-phosphate only has been depicted in this figure. This was demonstrated
to be the major pathway contributing to the labelling patterns observed. The
alternative pathway from mannitol to fructose 6-phosphate via fructose, mediated
by mannitol 2-dehydrogenase and hexokinase, could be depicted in addition to, or
instead of, the depicted pathway.
222
1 2 3 4 5 6
1 2 3 4 5 6
11
6PGDH
1 2 3 4 5
1 2 3 4 5
CO2
6-phosphogluconate
Ribulose 5-phosphate
Lactonase
1 2 3 4 5 6
1 2 3 4 5 6
6-phosphogluconoδ-lactone
G6PDH
1 2 3 4 5 6
1 2 3 4 5 6
Glucose 6-phosphate
Phosphopentose epimerase
1 2
1 2
Ribose 5-phosphate
Transketolase
1 2 3 4 5 6 7
2 3 4 5 6 7
1
Glyceraldehyde
Transaldolase
1 2 3 4 5 6
1 2 3 4 5 6
Fructose 6-phosphate
Fructose 6-phosphate
Mannitol 1-phosphate
?
MPP
223
1 2 3 4 5 6
1 2 3 4 5 6
[1-13C=6-13C]-Mannitol
Sedoheptulose 7-phosphate
3-phosphate
1 2 3 4 5 6
1 2 3 4 5 6
1 2 3 4 5 6
1 2 3 4 5 6
Xylulose 5-phosphate
1 2 3
1 2 3
PGI
MPD
1 2 3 4 5
1 2 3 4 5
3 4 5
3 4 5
1 2
1 2
1 2 3 4
1 2 3 4
Erythrose 4-phosphate
PGI
Xylulose 5-phosphate
Transketolase
1 2 3 4 5 6
1 2 3 4 5 6 PGI
1 2 3 4 5 6
1 2 3 4 5 6
Glucose 6-phosphate
TPS/TPP
1 2 3 4 5 6
1 2 3 4 5 6
Fructose 6-phosphate
1 2 3 4 5 6
1 2 3 4 5 6
1 2 3 4 5 6
1 2 3 4 5 6
Trehalose
3 4 5
3 4 5
1 2 3
1 2 3
Glyceraldehyde
3-phosphate
Glycolysis
carbon must have been cycled through one or more of these alternative pathways. The
PPP has previously been suggested as a means by which label scrambling could occur
(den Hollander and Shulman, 1983).
During the PPP step involving the oxidation of 6-phosphogluconate, the C1 is
lost to CO2. Thus, in a population of molecules with
only the
13
C-labelling of the C1 or C6,
13
C6 carbons would be retained. The action of the PPP transketolase and
transaldolase enzymes would ultimately result in the synthesis of fructose 6-phosphate
which was labelled on the C6 only (Figure 5.13). Conversion of this to G6P and
trehalose would result in an increase in the 13C6/13C1 ratio. This is consistent with the
observation for this strain. Action of the PPP would also yield [3-13C]-glyceraldehyde
3-phosphate. This could lead to further labelling of trehalose, but only with equimolar
contributions of 13C6 and 13C1 as outlined above (Section 5.4.3.1).
5.4.3.4 The Pentose Phosphate Pathway (Reverse Flux)
It has been previously assumed that the reactions of the non-oxidative portion
of the PPP are fully reversible, and that this could contribute to label scrambling
(Portais and Delort, 2002). Given a population of F6P molecules variously labelled on
one, both or neither terminal carbon, this would give rise to a number of PPP
intermediates reflecting this labelling (Figure 5.14). This may very well explain some
of the unidentified peaks in the 13C-labelled spectra acquired during this investigation,
since few of these intermediates are present in the Compound Standard Library. Good
evidence for this mechanism of scrambling is provided by the labelling pattern
observed for arabitol in the mpd1mdh1-107 strains. The passage of label via the
224
1 2 3 4 5
1 2 3 4 5
1 2 3 4 5
1 2 3 4 5
1 2 3 4 5
1 2 3 4 5
1 2 3 4 5
1 2 3 4 5
L-arabitol
D-ribulose 5-phosphate
Phosphopentose
Ldh1
epimerase
L-xylulose
1 2 3 4 5
1 2 3 4 5
1 2 3 4 5
1 2 3 4 5
1 2 3 4 5
1 2 3 4 5
1 2 3 4 5
1 2 3 4 5
D-ribose 5-phosphate
D-xylulose 5-phosphate
Lxr1
Pi
Xylitol
Xdh1
Transketolase
1 2 3 4 5 6 7
1 2 3 4 5 6 7
1 2 3 4 5 6 7
1 2 3 4 5 6 7
1 2 3
1 2 3
Glyceraldehyde
3-phosphate
D-xylulose
Pi
Sedoheptulose 7-phosphate
?
Xylulokinase
Transaldolase
1 2 3
4 5 6
4
5 6
1 2 3
1 2 3
4 5 6
1 2 3
4 5 6
D-fructose 6-phosphate
PGI
1 2 3 4 5 6
1 2 3 4 5 6 PGI
1 2 3 4 5 6
1 2 3 4 5 6
D-glucose 6-phosphate
1 2 3 4 5
1 2 3 4 5
1 2 3 4 5
1 2 3 4 5
1 2 3 4
1 2 3 4
D-erythrose 4-phosphate
D-xylulose 5-phosphate
Transketolase
1 2
3 4 5 6
1 2
3 4 5 6
1 2
3 4 5 6
1 2
3 4 5 6
D-fructose 6-phosphate
1 2 3
1 2 3
Glyceraldehyde
3-phosphate
Figure 5.14: Pentose phosphate pathway mechanism for 13C label scrambling from [1-13C]/
[6-13C]-glucose 6-phosphate to [1-13C]/[5-13C] L-arabitol. 13C-labelled carbons are shown in
red. Metabolites are shown in boxes. Enzymes are shown in italics. Abbreviations: PGI:
phosphoglucose isomerase; Xdh1: xylitol dehydrogenase; Lxr1: L-xylulose reductase;
Ldh1: L-arabitol dehydrogenase. Enzymatic cleavage sites are indicated by a dotted line.
Boxes of metabolites occurring more than once are given a matching colour. Co-factors are
not shown.
225
reverse of the PPP is the only obvious means by which arabitol could be labelled on
the C1. If the F6P pool had equal portions of label on the C1 and C6, then arabitol,
ceteris paribus, would also have equimolar labelling of its termini. However, as seen
in the spectra of the mpd1mdh1-107 replicates grown on [1-13C]-glucose, the labelling
of the C5 was 1.5-fold that of the labelling on the C1. This imbalance was explained
by the forward flux of the PPP which, as outlined above, lead to the preferential
labelling of the C5 of arabitol.
5.4.4 Mannitol Metabolism does not Contribute to NADPH Regeneration
The main role proposed for the mannitol cycle was for the regeneration of
NADPH at the expense of NADH (Hult and Gatenbeck, 1978). The evidence from the
investigation of mannitol mutants in S. nodorum indicates that while mannitol can be
synthesised and catabolised via the Mpd1 metabolic spur, and that this is the
dominant pathway, only a limited amount of mannitol can be synthesised via the
Mdh1 metabolic spur, and, under the experimental conditions employed in this study,
this pathway is essentially irreversible. While a de facto cycling of mannitol would, in
theory, be possible in the wild type strain, provided that both enzymes were active and
had access to a common pool of mannitol, the direction of operation would be the
reverse of that required for NADPH regeneration and would lead instead to net
NADPH consumption. For the mdh1-71 mutant, synthesis and catabolism of mannitol
via the Mpd1 pathway effectively disassociates mannitol metabolism from NADPH
regeneration. The fact that the phenotype of this mutant is unaffected by this, is
evidence that NADPH regeneration is not a major role of mannitol in S. nodorum.
Furthermore, in the mpd1-1 mutant grown on mannitol, NADPH would be a major
226
product of the catabolism of mannitol via the Mdh1 pathway. However, this strain
was unable to grow on mannitol as a sole carbon source. Furthermore, the enzyme
assays in Chapter 3 demonstrated that the activity of the catabolic reaction was poor.
It would appear, therefore, that the main pathway of NADPH regeneration is the
pentose phosphate pathway.
5.4.5 Experimental Considerations
5.4.5.1 Co-located Peaks in Biological Samples Obscure Labelling
The biological extracts represented a more complex sample than the pure
standards. Carbons from different compounds which had an equivalent chemical
environment could not be separated on the spectrometer used. In natural abundance
samples, this was apparent where a peak for a compound was more intense than it
should be according to the standard spectrum. In some cases it was found that a peak
from another standard compound was co-located, in other cases it was presumed that
an unidentified compound was involved. In the case of the
13
C-labelled spectra this
created a problem, since labelling might be clearly present in such a peak, but some
ambiguity would arise in terms of whether the labelling was due to one or both (or
more) compounds, and the percentage labelling which would be due to each. A
particular case is the C5 of arabitol. In the mpd1mdh1-107 samples grown on labelled
glucose, this peak had a intensity equivalent to being labelled 12-fold above natural
abundance. The ability of all other strains to metabolise glucose to mannitol, however,
resulted in the presence of a mannitol C1,6 sideband which was co-located with the
C5 of arabitol and prevented the determination of the amount of label in this carbon.
227
5.4.5.2 Low Sample Weights Affect Detection of Low Abundance Metabolites
The amount of mycelium available for analysis was quite variable between the
strains. SN15 and mdh1-71 gave consistently better yields (mean dry weight per
sample of 122.7 ± 21.7 mg and 159.4 ± 30.7mg respectively) than mpd1-1 and
mpd1mdh1-107 (66.5 ± 13.2 mg and 45.0 ± 7.0 mg respectively) when grown on 40
mM glucose. Following the extraction of polar metabolites, the mean dry weights of
extracts available for analysis were 28.2 ± 8.1 mg (SN15), 21.0 ± 3.3 mg (mdh1-71),
12.9 ± 1.1 mg (mpd1-1) and 12.8 ± 3.2 mg (mpd1mdh1-107). Since
13
C-NMR is a
relatively insensitive technique, low abundance metabolites were either not observed
in samples with a lower dry weight, or only their most intense peaks were seen. This
problem could be alleviated in future by increasing the number of cultures for the
slower-growing strains and combining the harvested mycelium to bulk up the sample
size.
5.4.5.3 Spectrometer Artefacts/Variation
There were a number of peaks which were observed regularly in some samples
including the standards and which appeared to be artefacts. These were discounted
from the analysis. It was also noted that samples which were run consecutively
produced spectra which varied less from each other in terms of chemical shift, than
spectra which were not consecutively run. There was some minor variation in the
operating temperature of the spectrometer and this may account for some of the
variation in chemical shifts between samples. The use of an internal reference
compound corrected for this latter variation.
228
5.4.5.4 Quantification of 13C Labelling
The quantification of the distribution of
13
C-label in NMR spectra has been
approached by a number of methods. This includes comparison of the labelled peak(s)
of a compound with unlabelled sibling peak(s) (Martin et al., 1998; Bago et al., 1999;
Peksel et al., 2002), and the addition of a standard of known concentration, such as
EDTA (Ramstedt et al., 1989; Ceccaroli et al., 2003) and maleate (Jobic et al., 2007).
This latter approach has also been used to determine the efficiency of the extraction
process (Aubert et al., 1996b) and the concentration of metabolites in a spectrum.
The use of natural abundance peaks for quantification of label in labelled
peaks, depends on the assumption that the natural abundance peaks themselves have
not been labelled. However, the ratios of the NA peaks in the 6-carbon compounds
were equivalent to those of their compound standards. This was considered sufficient
evidence that there was no uniform labelling of these peaks above NA. This also
constituted good evidence that NA peaks of lower-carbon-number compounds were
not uniformly labelled, since cycling and scrambling of label would be revealed as
perturbations in the ratios of the “NA” peaks of other compounds.
5.4.5.5 Internal Referencing of Chemical Shifts
There have been a number of internal and external standards used in fungal
13
C-NMR studies in order to act as a point of reference for the chemical shifts of
resonance peaks. These include acetone (de Koker et al., 2004), acetate (Donker and
Braaksma, 1997), EDTA (Martin et al., 1988; Ramstedt et al., 1989; Ceccaroli et al.,
229
2003),
dioxane
(Yoshida
et
al.,
1984;
Thomas
and
Baxter,
1987),
hexamethyldisiloxane (Jobic et al., 2007), TMS (Martin et al., 1984; Martin et al.,
1985; Martin et al., 1988; Shachar-Hill et al., 1995; Martin et al., 1998; Bago et al.,
1999) and trimethylsilyl propionate (TMSP) (Forgue et al., 2006). Deficiencies have
been noted with each of these and the recommendation has been made that 2,2dimethyl-2-silapentane-5-sulfonic acid (DSS) be used as the universal standard, due
to its being insensitive to variations in temperature and pH, and the fact that it is
chemically inert and has a single, sharp, unambiguous highfield resonance peak
(Wishart and Sykes, 1994). This recommendation was not noted until after this
portion of the study had commenced. The practise of using the solvent to lock the
signal, and using a compound known to be present and abundant in the samples as an
internal reference, such as mannitol, glucose or trehalose, was adopted as per Dijkema
et al. (1985). Although glycerol was the most abundant peak in some spectra, the fact
that it only has two peaks, and that the more intense C1,3 peak could not always be
unambiguously assigned, rendered it unsuitable as an internal reference compound.
For the purposes of this study, this practise gave good correspondence between the
chemical shifts of compounds in the one sample, along with the chemical shifts of
their compound standards, and the same compounds in different samples. Typically,
the sibling peaks of the reference compound had a variation in chemical shift of no
more than 0.02 ppm from the compound standard, and the peaks of other major
metabolites in the same sample had a variation of no more than 0.03 ppm. It is
entirely likely that most of this variation would be explained by the compound
standards having been acquired with slight variations in temperature. The use of an
internal standard with the compound standards would have allowed correction for any
such variation.
230
5.4.5.6 Limitations of Published Chemical Shifts
The identification of unknown peaks appearing in the spectra of biological
samples was hindered by the lack of a more comprehensive public database of
naturally occurring metabolites. While the SDBS database contains some 12,500 13CNMR spectra (AIST, 2008), it does not at present include some compounds of interest
to this study, including fructose 6-phosphate and mannitol 1-phosphate. Whilst it was
a simple matter to produce standard spectra for these compounds, the identification of
unknown peaks was constrained by a lack of certainty that the unknown compound
was contained in the SDBS database. Furthermore, the searching function on the
SDBS database does not allow for searches delimited to a particular carbon atom of a
compound, but returns any carbon atom matching a submitted chemical shift
parameter. When searches were conducted, they were required to take into account
the observation above (Table 5.2) that the published chemical shift for a given peak
could vary by as much as 2.8 ppm (after excluding the extreme chemical shifts
reported in Fan (1996)), depending on the source of the data. A variation of this
magnitude, even when limiting the search to 6-carbon compounds, could return well
in excess of 100 possible compounds. A search function which returned records
matching two or more chemical shift parameters would be of value in potential
identification of unknown peaks in a spectrum.
While most published spectra agreed on the assignment of carbon atoms to
particular peaks, there were a couple of exceptions to this. In the case of trehalose,
there is some disagreement in the published spectra regarding the assignment of the
C2 and C5 carbons. The SDBS database and some fungal 13C NMR studies place the
231
C5 downfield of the C2 (Dijkema et al., 1985; Thomas and Baxter, 1987; Bago et al.,
1999; Rangel-Castro et al., 2002; AIST, 2008). In Fan’s Chemical Tables and a
smaller number of studies, the assignment of these peaks reversed (Martin et al.,
1985; Fan, 1996; Deveau et al., 2008). While this was not of great consequence to the
findings of this study in terms of the pathways by which labelling occurs, it was noted
in several spectra, where the C1 of trehalose had 23-fold labelling or more above NA,
that the more downfield of these two peaks exhibited spin-spin splitting. This was
consistent with its being the immediate neighbouring carbon atom to the C1. It was
therefore decided to adopt the less frequently applied assignment pattern in this study
i.e. C2 at 72.08 ppm and C5 at 70.97 ppm.
5.4.5.7 Necessity for a Local Library of Compound Standards
Given that spectra acquired on the same instrument under the same conditions
agree well with each other, it was apparent that generating a local library of
compound standards conferred an advantage in the process of identifying unknown
peaks. This library was designed to contain not only compounds anticipated or
predicted to be involved in mannitol metabolism, but also those compounds generally
involved in glycolysis, gluconeogenesis, amino acid metabolism, etc. Elimination of
peaks assigned to any these compounds would better allow the identification of
significantly abundant or labelled unknown peaks. In the light of the relevance of the
PPP to metabolism in the strains under investigation, it would be of value to acquire
standards for intermediates in this pathway.
232
5.4.5.8 Assumption of Labelling of Mannitol on One Terminal Carbon
It was assumed throughout the study, for the purposes of determining the
labelling above NA of the C1 of mannitol, that this compound was labelled on only
one terminal carbon. Since the molecule is symmetrical, it is of no consequence for
the calculation whether it is the “C1” or “C6” which is labelled. Given the labelscrambling observed via the PPP and the TPI triangle, it is likely, that there would be
a population of mannitol molecules produced which were labelled on both or neither
of the terminal carbons. As long as these populations were of equal proportions, there
should be no impact on the validity of the labelling calculation. The TPI triangle
would not be expected to favour production of one population over the other. A large
flux through the PPP would result in the production of unlabelled fructose 6phosphate, but not in fructose 6-phosphate labelled on both termini. This could
translate into a dilution of signal in mannitol. While this limitation of the calculation
is acknowledged, it does not seem to have seriously impacted on the levels of
labelling observed.
5.5 CONCLUSION
A 13C-NMR investigation of carbon metabolism in the Stagonospora nodorum
wild type strain SN15, and mutant strains with disrupted mannitol metabolism genes,
revealed the operation of a number of primary metabolic pathways (Figure 5.15). It
confirmed that the postulated mannitol cycle is not a necessary feature, and most
likely does not exist in this species. It was demonstrated that there are two separable
pathways of mannitol metabolism. The Mdh1 pathway contributed to mannitol
233
trehalose
TREHALOSE
METABOLISM
trehalose 6P
glucose
PENTOSE PHOSPATE
PATHWAY
ribulose 5P
glucose 6P
mannitol
fructose
MANNITOL
fructose 6P
METABOLISM
mannitol 1P
mannitol
erythrose
th
4P
D-xylulose 5P
fructose 1,6-bisP
D-xylulose
glycerol 3P
DHAP
glyceraldehyde 3P
xylitol
glycerol
DHA
GLYCEROL CYCLE
1,3-bisPG
L-xylulose
3-PG
L-arabitol
ARABITOL
GLUCONEOGENESIS/ 2-PG
GLYCOLYSIS
METABOLISM
PEP
L-asparagine
pyruvate
acetyl
y CoA
L-aspartate
citrate
oxaloacetate
arginosuccinate
UREA
citrulline
L-arginine
CYCLE
ornithine
L-alanine
malate
fumarate
TCA
isocitrate
L-glutamine
CYCLE
a-ketoglutarate
L l t
L-glutamate
t
succinate
succinyl CoA
Figure 5.15: Summary of metabolic pathways demonstrated to be active in Stagonospora
nodorum, based on the detection of metabolic intermediates in 13C-NMR spectra. Detected
metabolites are shown in bold and inferred metabolites in italics. Not all possible pathways
for metabolism of compounds are shown. Abbreviations: P = phosphate; DHAP =
dihydroxyacetone; PG = phosphoglycerate; PEP = phosphoenol pyruvate.
234
synthesis, but was unable to catabolise mannitol under the conditions of the
experiment. The Mpd1 pathway demonstrated the capacity for mannitol synthesis and
catabolism and was the dominant pathway by which this occurred. There was no third
pathway of mannitol metabolism detected. It could not be determined how the Mpd1
pathway mediated the conversion of mannitol to mannitol 1-phosphate. The simplest
mechanism for this conversion would require a mannitol kinase, but activity
indicating the presence of such an enzyme has not been detected in this species. A
number of unidentified peaks were observed in the spectra, but none which suggested
the existence of an intermediate compound between mannitol and mannitol 1phosphate.
The pattern of
13
C-label scrambling observed in the mutant strains was
consistent with the cycling of labelled substrate via the pentose phosphate pathway
and the aldose/triosephosphate isomerase triangle. Scrambling occurred in the absence
of either or both of the mannitol metabolic spurs, and would not, therefore,
necessarily constitute evidence of a mannitol cycle in other species. It would appear
that previous studies, which have attributed the bulk of observed scrambling to a
mannitol cycle, and used the degree of scrambling as an measure of the cycling
between mannitol and hexose, will have resulted in an overestimate of the rapidity of
this cycling (Pfeffer et al., 2001).
The role proposed for mannitol of NADPH regeneration, which formed the
basis of the mannitol cycle theory, is not supported by this investigation. The
13
C-
label scrambling pattern of trehalose in the mutant strains indicated that NADPH
requirements are met by the passage of carbon via the pentose phosphate pathway.
235
The operation of the Mpd1 pathway in isolation would make no contribution to
NADPH generation, and the apparent unidirectional operation of the Mdh1 pathway
in conjunction with the Mpd1 pathway would result in a de facto mannitol cycle
operating in reverse, and resulting in NADPH consumption. There is no evidence that
such a cycle was in operation.
236
CHAPTER 6: GENERAL CONCLUSIONS
237
6.1 Overview
The relationship between metabolism and pathogenicity in the wheat pathogen
Stagonospora nodorum was investigated with specific reference to the metabolism of
mannitol. This compound is one of the most abundant metabolites found in the
mycelium of fungi and while a number of roles have been attributed to it, there has
been little experimental evidence to support them. The metabolism of mannitol has
been described as occurring in an enzymatic cycle, the principle roles of which have
been for NADPH regeneration, or for dissipation of energy. It has also been suggested
that mannitol has a role in phytopathogenicity. A strain of S. nodorum in which the
mannitol 1-phosphate dehydrogenase (Mpd1) gene was disrupted, was unable to
sporulate in planta, but was still able to synthesise mannitol at about 10% of wild type
levels. The unidirectional nature of the theoretical mannitol cycle suggested that
disruption of a mannitol 2-dehydrogenase (Mdh1) gene would abolish mannitol
utilisation. However, a strain of S. nodorum in which this gene was disrupted was
phenotypically identical to the wild type, retaining full pathogenicity and wild type
production of mannitol. This study aimed to create a mutant harbouring both
disruption constructs in order to establish the relationship between these two genes, to
identify any alternative pathways of mannitol metabolism, and to attempt to abolish
the metabolism of mannitol and by doing so elucidate its role in pathogenicity.
6.2 Key Findings
Disruption of both Mdh1 and Mpd1 genes resulted in a strain which was
unable to synthesise or catabolise mannitol, although it was readily able to accumulate
238
mannitol from complex medium. Evidence of the inability of the strain to catabolise
exogenously accumulated mannitol was provided by the lack of a demonstrated
ability to grow on mannitol as a sole carbon source. Accumulated mannitol was
retained as a stable pool and it required serial plating of the strain on minimal medium
in which no mannitol was present, for mannitol levels to be reduced to the extent
where in vitro sporulation was abolished. The strain was unable to sporulate in planta,
although the addition of exogenous mannitol partially corrected this defect. This is the
first time a role has been conclusively demonstrated for this compound.
A 13C-NMR investigation of wild type strains and mutant strains with one or
both of the Mdh1 and Mpd1 genes disrupted was undertaken to further characterise
mannitol metabolism. Strains (including the wild type) with an intact Mpd1 gene were
characterised by having mannitol as their principal soluble carbohydrate. The mpd1
mutants were characterised by having trehalose and glycerol as their principal soluble
carbohydrates. There was no evidence to support a mannitol cycle in S. nodorum. It
was demonstrated that cycling of labelled carbon can be explained by the
triosephosphate isomerase triangle and the pentose phosphate pathway in the absence
of mannitol metabolism. The importance of mannitol metabolism to carbon cycling is
likely to have been overestimated in previous studies. There was no evidence to
suggest that mannitol metabolism is critical to NADPH regeneration.
A metabolomics investigation of diseased versus healthy tissue from leaves
inoculated with S. nodorum compared to mock-inoculated and uninoculated leaves
did not detect any compounds which could be characterised as phytoalexins. PCA
showed that the fungal specific metabolites mannitol and trehalose were associated
239
with diseased leaves, while the plant-specific metabolite sucrose was associated with
healthy leaves.
6.3 Future Directions
It is still not apparent how the catabolism of mannitol is accomplished in the
Mpd1-mediated pathway. There was no obvious candidate intermediate metabolite
suggested by the
13
C-NMR investigation. If such a intermediate exists, it must be
rapidly metabolised. This is not uncommon for metabolic intermediates in other
pathways. While mannitol kinase activity has not previously been found in this
species, the presence of such an enzyme would represent the simplest explanation for
the phosphorylation of mannitol to mannitol 1-phosphate.
There were a large number of unidentified peaks observed in the
13
C-NMR
spectra including a number which appeared to be labelled following growth upon 13Clabelled substrate. These may be worth further investigation and the simplest way to
progress this would be through the use of alternatively labelled substrates. The use of
[2-13C] or [1,2-13C] would help to identify peaks belonging to the same compound.
Given the flux observed through the pentose phosphate pathway, it is likely that
adding intermediates of this pathway to the compound standard library would result in
further identification of unknown peaks.
The fact that the abolition of synthesis of a fungal-specific metabolite leads to
the inability to sporulate in planta is of significance for the control of this pathogen. A
more complete understanding of the mechanism by which mannitol impacts on
240
sporulation would potentially lead to anti-fungal strategies. It would be interesting to
see whether abolition of mannitol had a similar effect in other phytopathogens. It has
also been demonstrated in the previous studies that the expression of mannitol
catabolic genes in plants can increase their resistance to fungal pathogens. A recent
study described the effects on salt and water tolerance of wheat transformed with an
E. coli mannitol dehydrogenase gene. It would be interesting to see whether the
presence of this gene conferred any improved resistance to a fungal pathogen such as
S. nodorum.
241
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296
CHAPTER 8: APPENDICES
297
Table 8.1: Plant species other than Triticum aestivum L. subsp. aestivum (excluding hybrids) which have been reported as hosts of Stagonospora nodorum.
The fungus was regarded as being pathogenic where studies have reported the plant as exhibiting disease or necrosis, or where the pathogen had caused
infection/lesion formation and/or undergone pycnidiation. Instances where the literature was in conflict are noted, as are instances where the pathogen was
described as causing disease, but was not reisolated from the plant. Also noted are studies where a plant was described as being a host for S. nodorum, but
where infection was not explicitly reported.
Scientific Name
Aegilops bicornis (Forsk.) Jaub. & Spach.
Common Name
goatgrass
References
(Frauenstein and Hammer, 1985; Hammer, 1985)
Aegilops columnaris Zhuk.
goatgrass
(Frauenstein and Hammer, 1985; Hammer, 1985)
goatgrass
(Frauenstein and Hammer, 1985; Hammer, 1985)
Aegilops crassa subsp. crassa Boiss.
Persian goatgrass
(Frauenstein and Hammer, 1985; Hammer, 1985)
Aegilops cylindrica Host
jointed goatgrass
(Frauenstein and Hammer, 1985; Hammer, 1985; Khokhar and Pacumbaba, 1987)
Aegilops geniculata Roth
Ovate goatgrass
(Frauenstein and Hammer, 1985; Hammer, 1985)
Aegilops juvenalis (Thell.) Eig.
goatgrass
(Frauenstein and Hammer, 1985; Hammer, 1985)
Aegilops kotschyi Boiss.
ovate goatgrass
(Frauenstein and Hammer, 1985; Hammer, 1985)
Aegilops longissima Schweinf. & Muschl.
goatgrass
(Frauenstein and Hammer, 1985; Hammer, 1985; Ecker et al., 1990b)
Aegilops lorentii Hochst. syn. biuncialis
Vis.
Aegilops markgrafii (Greuter) K.
Hammer
Aegilops mutica Boiss.
Lorent’s goatgrass
(Frauenstein and Hammer, 1985; Hammer, 1985)
goatgrass
(Frauenstein and Hammer, 1985; Hammer, 1985)
goatgrass
(Frauenstein and Hammer, 1985; Hammer, 1985)
Aegilops neglecta subsp. neglecta
three-awned goatgrass
(Frauenstein and Hammer, 1985; Hammer, 1985)
Aegilops neglecta subsp. recta
goatgrass
(Frauenstein and Hammer, 1985)
Aegilops peregrina (Hack.) Maire &
Weiller
goatgrass
(Frauenstein and Hammer, 1985; Hammer, 1985)
Aegilops comosa Sibth. & Smith
1
Continued on the following page
Table 8.1: (continued)
Scientific Name
Aegilops searsii Feldman & Kislev
Common Name
Sears’ goatgrass
References
(Hammer, 1985)
Aegilops speltoides Tausch.
goatgrass
(Frauenstein and Hammer, 1985; Hammer, 1985; Ecker et al., 1990a)
Aegilops tauschii Cross syn. squarrosa
L. syn. Triticum tauschii 2
Aegilops triuncialis L.
Tausch’s goatgrass
barbed goatgrass
(Jahier and Trottet, 1980; Frauenstein and Hammer, 1985; Hammer, 1985; Doussinault et al., 1992; Ma
and Hughes, 1993; Murphy, 1997; Murphy et al., 2000; Loughman et al., 2001; Murphy et al., 2001)
(Frauenstein and Hammer, 1985; Hammer, 1985)
Aegilops umbellulata Zhuk.
goatgrass
(Frauenstein and Hammer, 1985; Hammer, 1985; Maksimov et al., 2006)
goatgrass
(Frauenstein and Hammer, 1985; Hammer, 1985)
Aegilops ventricosa Tausch.
barbed goatgrass
(Trottet et al., 1975; Frauenstein and Hammer, 1985; Hammer, 1985)
Agropyron cristatum (L.) Gaernt.
fairway crested wheatgrass
(Krupinsky, 1982; Krupinsky, 1997a)
Agropyron desertorum (Fisch. ex Link)
Schultes
desert wheatgrass/ standard
crested wheatgrass/clustered
wheatgrass
Siberian wheatgrass
(Krupinsky, 1982; Krupinsky, 1997a)
couchgrass/dog grass/
quackgrass
(Becker, 1957; Derevyankin, 1969; Williams and Jones, 1973; Ao and Griffiths, 1976; Rufty et al., 1981b)
Aegilops uniaristata Vis.
1
Agropyron fragile subsp. Sibiricum
(Willd.) Melderis
Agropyron repens (L.) Beauv. 3
Agropyron spp.
(Krupinsky, 1997a)
(Krupinsky, 1983)
Agrostis capillaris syn. tenuis L.
Colonial bent
(Williams and Jones, 1973; Ao and Griffiths, 1976)
common wild oat
(Williams and Jones, 1973; Ao and Griffiths, 1976)
Avena ludoviciana Dur.
oats
(Williams and Jones, 1973; Ao and Griffiths, 1976)
Avena sativa L.
oats
(Clark and Zillinsky, 1960; Arseniuk et al., 1997)
Bromus diandrus Roth. syn. gussonii
Parl. 4
Bromus hordeaceus syn. mollis L.
brome grass/great brome
(Brown and Rosielle, 1980)
soft brome
(Ao and Griffiths, 1976)
awnless brome/smooth
brome/smooth bromegrass
(Dorokhova, 1967; Krupinsky, 1986b; Krupinsky, 1986a; Khokhar and Pacumbaba, 1987; Krupinsky,
1994; Krupinsky, 1997b; Krupinsky, 1997a)
Avena fatua L.
3
Bromus inermis Leyss.
3
Continued on the following page
Table 8.1: (continued)
Scientific Name
Bromus sterilis L.
Common Name
poverty brome/barren
brome/sterile brome
downy brome/early chess/
military grass/thatch
bromegrass
wild barley/foxtail barley
(Fernandes, 1985)
Bermuda grass
(Khokhar and Pacumbaba, 1987)
(Baker, 1969)
Elymus canadensis L.
cocksfoot/orchard grass/
cocksfoot grass
Canadian wildrye
Elymus histrix syn. Hystrix patula L.
eastern bottlebrush grass
(Rufty et al., 1981b)
Bromus tectorum L.
Critesion syn. Hordeum jubatum L.
Cynodon dactylon (L.) Pers.
Dactylis glomerata L.
5
6
Elymus virginicus L.
References
(Williams and Jones, 1973; Ao and Griffiths, 1976; Harrower, 1977)
(Krupinsky, 1997a)
(Krupinsky, 1997a)
(Rufty et al., 1981b)
Elytrigia repens L.
couch grass/quackgrass
(Shearer and Zadoks, 1972; Khokhar and Pacumbaba, 1987)
Festuca elatior syn. arundunacea L.
tall fescue
(Ao and Griffiths, 1976; Rufty et al., 1981b)
Festuca pratensis Huds.
meadow fescue
(Rufty et al., 1981b)
Holcus lanatus L.
(Williams and Jones, 1973; Ao and Griffiths, 1976)
Hordeum brachyantherum Nevski 4
Yorkshire fog/common
velvetgrass
meadow barley
Hordeum bulbosum L.
bulbous barley
(Rufty et al., 1981b)
(Brown and Rosielle, 1980)
Hordeum marinum L.
barley grass/mediterranean
barley grass
sea barley
Hordeum murinum L. subsp leporinum
(Link) Arcang.
Hordeum pusillum Nutt.
barley grass/mouse barley/
wild barley
little barley
(Brown and Rosielle, 1980)
Hordeum hystrix Roth.
4
(Sprague, 1956)
(Rufty et al., 1981b)
(Rufty et al., 1981b; Cunfer and Youmans, 1983; Khokhar and Pacumbaba, 1987; Ueng et al., 1995)
Continued on the following page
Table 8.1: (continued)
Scientific Name
Hordeum vulgare L.
Common Name
barley/cereal
barley/common barley
Hordeum vulgare L. pallidum Ser. [syn.
vulgare]
Leymus syn. Elymus angustus (Trin.)
Pilger
Leymus syn. Elymus cinereus (Scribn. &
Merr.) A. Löve
Leymus racemosis (Lam.) Tzvelev subsp.
vacemosus [syn. Elymus giganteus Vahl.]
Lolium multiflorum Lam.
barley
References
(Hansen and Magnus, 1969; Richardson and Noble, 1970; Richardson, 1972; Shearer and Zadoks, 1972; Holmes
and Colhoun, 1973; Hewett, 1975; Jones, 1975; Ao and Griffiths, 1976; King, 1977; Rufty et al., 1981b; Sharma
et al., 1982; Sharma and Brown, 1983; Cunfer, 1984; Osbourn et al., 1987; Berecket et al., 1990; Mansfield et al.,
1991; Newton and Caten, 1991; Cunfer et al., 1992; Polley et al., 1993; Krupinsky, 1994; Ueng et al., 1995;
Arseniuk et al., 1997; Bousquet and Kollmann, 1998; Duczek et al., 1999; Turkington et al., 2002)
(Weber, 1922b)
Altai wildrye
(Krupinsky, 1994; Krupinsky, 1997b; Krupinsky, 1997a)
basin wildrye
(Krupinsky, 1994; Krupinsky, 1997b; Krupinsky, 1997a)
mammoth wildrye,
Volga wildrye
Italian ryegrass
(Krupinsky, 1997a)
perennial ryegrass
annual ryegrass
(Shearer and Zadoks, 1972; Ao and Griffiths, 1976; Rufty et al., 1981b; Khokhar and Pacumbaba, 1987; Jenkyn
and King, 1988; Ueng et al., 1995)
(Brown and Rosielle, 1980)
Smith’s melicgrass
(Sprague, 1955)
Alaska oniongrass
(Sprague, 1955)
Pascopyrum syn. Agropyron smithii
(Rydb.) A. Löve
Phleum pratense L.
pubescent wheatgrass/
western wheatgrass
Timothy grass
(Krupinsky, 1982; Krupinsky, 1994; Krupinsky, 1997b)
(Ao and Griffiths, 1976; Harrower, 1977)
Poa annua L.
annual meadow-grass
(Becker, 1957; Shearer and Zadoks, 1972; Ao and Griffiths, 1976)
flattened meadow-grass
(Rufty et al., 1981b)
Lolium perrene L.
Lolium rigidum Gaudin 4
4
Melica smithii (Porter ex A. Gray) Vasey
Melica subulata (Griseb.) Scribn.
Poa compressa L.
4
7
Poa diversifolia (Boiss. & Ball.) Hack. ex
Boiss.
Poa pratensis L.
Poa trivialis L.
(Harrower, 1977; Rufty et al., 1981b)
(Rufty et al., 1981b)
smooth meadow-grass or
Kentucky bluegrass
rough bluegrass
(Weber, 1922b; Becker, 1957; Williams and Jones, 1973; Rufty et al., 1981b)
(Williams and Jones, 1973)
Continued on the following page
Table 8.1: (continued)
Scientific Name
Psathyrostachys juncea (Fisch.) Nevski
Common Name
Russian wildrye
References
(Krupinsky, 1997a)
Secale cereale L.
rye
(Weber, 1922b; Becker, 1957; Derevyankin, 1969; Arseniuk et al., 1997; Joshi and Miedaner, 2003)
Thinopyrum syn. Agropyron intermedium
(Host) Barkworth & Dewey
Triticum aestivum L. subsp. compactum
(Host) Mackey
Triticum aestivum L subsp. macha
(Dekapr. & A. M. Menabde) Mackey
Triticum aestivum L subsp. spelta (L.)
Thell.
Triticum aestivum L. subsp.
sphaerococcum (Perc.) Mackey
Triticum monococcum L. subsp.
monococcum
Triticum monococcum L. subsp.
aegilopoides (Link) Thell.
Triticum timopheevii (Zhuk.) Zhuk.
subsp. timopheevii
Triticum turgidum L. subsp. carthlicum
(Nevski) A. Löve & D. Löve
Triticum turgidum L. subsp. dicoccoides
(Körn. ex Ascb. & Graebn.) Thell.
Triticum turgidum L. subsp. dicoccon
(Schrank.) Thell. syn. T. dicoccum
Shrank.
Triticum turgidum L. subsp. durum
(Desf.) Husn.
intermediate wheatgrass
(Krupinsky, 1982; Krupinsky, 1994; Krupinsky, 1997b)
club wheat/cluster wheat/
dwarf wheat/hedgehog wheat
(Weber, 1922b; Krupinsky et al., 1977; Mielke, 1989)
Triticum turgidum L. subsp. polonicum
(L.) Thell.
Triticum turgidum L. subsp. turgidum
(Mielke, 1989)
dinkel wheat/spelt wheat
(Weber, 1922b; Krupinsky et al., 1977; Mielke, 1989; Aguilar et al., 2005)
Indian dwarf wheat/shot
wheat
einkorn wheat
(Tomerlin et al., 1984)
(Weber, 1922b; Tomerlin et al., 1984; Mielke, 1989; Ma and Hughes, 1993; Singh et al., 2006)
wild einkorn
(Tomerlin et al., 1984)
Sanduri wheat
(Krupinsky et al., 1977; Scharen and Eyal, 1980; Tomerlin et al., 1984; Mielke, 1989; Ma and Hughes,
1993; Ma and Hughes, 1995; Singh et al., 2006)
(Tomerlin et al., 1984; Mielke, 1989)
Persian black wheat/Persian
wheat
wild emmer wheat
emmer wheat, Samba wheat
durum wheat, macaroni
wheat
(Krupinsky et al., 1977; Scharen and Eyal, 1980; Tomerlin et al., 1984; Mielke, 1989; Singh et al., 2006;
Chu et al., 2008)
(Weber, 1922b; Chona and Munjal, 1954; Krupinsky et al., 1977; Tomerlin et al., 1984; Mielke, 1989;
Ma and Hughes, 1993; Singh et al., 2006)
Polish wheat
(Weber, 1922b; Krupinsky et al., 1977; Scharen and Eyal, 1980; Mullaney et al., 1983; Tomerlin et al.,
1984; Mielke, 1989; Gilbert and Tekauz, 1992b; Ma and Hughes, 1993; Fernandez et al., 1996; Cao et
al., 2001; Xu et al., 2004a; Singh et al., 2006; Singh et al., 2007)
(Weber, 1922b; Mielke, 1989)
rivet wheat/cone wheat
(Mielke, 1989)
Continued on the following page
Table 8.1: (continued)
Scientific Name
Vulpia bromoides (L.) Gray 4
NA
Common Name
squirrel tail fescue/brome
fescue
Triticale
References
(Brown and Rosielle, 1980)
(Khokhar and Pacumbaba, 1987; Valuevich et al., 1992; Góral et al., 1994; Ueng et al., 1995; Abreu et al.,
1996; Arseniuk et al., 1997; Arseniuk et al., 1998; Oettler and Schmid, 2000; Tian et al., 2005)
Notes:
1
Reaction described by Hammer (1985) as resistant to moderately resistant
2
Most studies report resistance due to a single locus. Ma and Hughes (1993) found S. nodorum caused necrosis, while Jahier and Trottet (1980) found it to be
only weakly pathogenic
3
Weber (1922b) found no disease was caused by S. nodorum on this species
4
Reported as a host from which the fungus was isolated, but no disease symptoms were described
5
Very weakly pathogenic.
6
Slight infection only with no pycnidia produced
7
Fungus was reported as causing infection, but was not reisolated
NA = not applicable
Table 8.2: Names (in English) which have been used to describe the disease caused by Stagonospora nodorum on wheat.
Disease Name
References
(Salmon and Throckmorton, 1930)
basal glume rot
dry blight (caused by three Septoria spp. – from the description one of them was certainly (Sutton, 1920)
S. nodorum)
(Weber, 1922a; Rosen, 1947; Doling, 1961; Scharen, 1963; Scharen and Krupinsky, 1970; Melville
and Jemmett, 1971; Kees and Obst, 1972; Harrower, 1974; van der Wal and Cowan, 1974; Hampton,
1975; Kent and Strobel, 1976; Harris and Grossbard, 1978; Scharen and Bryan, 1979; Straley and
Scharen, 1979; Brown and Paddick, 1980; Nelson, 1980; Allingham and Jackson, 1981; Cunfer and
Johnson, 1981; Rufty et al., 1981a; Babadoost and Hebert, 1982; Luke et al., 1983; Karjalainen and
Salovaara, 1988; Peltonen and Karjalainen, 1992; Cunfer, 1993; Aris, 1999; Halama et al., 1999;
glume blotch
Pazzagli et al., 1999)
(Howard
et al., 1994)
glume blotch and leaf spot
(Mehta, 1975; Kleijer et al., 1977; Wainshilbaum and Lipps, 1991; Peltonen, 1997; Fraaije et al.,
Leaf and glume blotch
2001; Agrios, 2005; Tan, 2007)
(Cunfer and Youmans, 1983; Cooley et al., 1999)
leaf spot and glume blotch
(Scott, 1988)
leaf spot disease of wheat (in concert with S. tritici and Pyrenophora tritici-repentis)
(McFadden and Harding, 1989)
leafspotting complex (in concert with Pyrenophora tritici-repentis)
(Bockus and Shroyer, 1998)
Nodorum leaf and glume blotch
(Tyldesley
and Thompson, 1980; Rosielle and Brown, 1981)
Septoria (including in concert with Septoria tritici)
(Broscious et al., 1985; Lupei et al., 2000; Yusupova et al., 2006)
septoria blotch
(Hart et al., 1984; Mundt et al., 1995)
septoria blotch disease (in concert with Septoria tritici)
(Pedersen and Hughes, 1992; Sundin et al., 1999; Bockus et al., 2001)
septoria (disease) complex (in concert with Septoria tritici)
(Rosen, 1921; Cunfer et al., 1980; Scharen and Eyal, 1980; Eyal, 1981; Negassa, 1987; Ecker et al.,
septoria glume blotch
1989; Bruno and Nelson, 1990; Bostwick et al., 1993; Hu et al., 1996; Tyryshkin and Ershova, 2004)
(Watson et al., 1982; Leath and Papke, 1989; Caten and Newton, 2000)
septoria leaf and glume blotch
(Nass and Johnston, 1985; Peltonen, 1993)
septoria leaf blotch
(Gilbert
and Tekauz, 1992a)
septoria leaf blotch (in concert with S. avenae f. sp. triticea)
(Gilbert et al., 1993; Sooväli et al., 2006)
septoria leaf blotch complex (in concert with S. tritici and S. avenae f. sp. triticea)
(Shipton, 1966)
septoria leaf spot and glume blotch
Continued on the following page
Table 8.2 (Cont.): Names (in English) which have been used to describe the disease caused by Stagonospora nodorum on wheat.
Disease Name
septoria nodorum blotch
septoria nodorum leaf and glume blotch
septoria nodorum leaf blotch
Septoria nodorum spot
Septoria (caused by three Septoria spp. – from the description one of them was
certainly S. nodorum)
Stagonospora blotch
Stagonospora glume blotch
Stagonospora leaf blotch
stagonospora nodorum blotch (SNB)
Stagonospora nodorum leaf and glume blotch
Stagonospora nodorum leaf blotch
wheat glume blotch
wheat leaf and glume blotch
wheat leaf blotch
Reference
(Luke et al., 1985; Spadafora et al., 1987; Stooksbury et al., 1987; Nelson and Marshall, 1990;
Scharen et al., 1991; Shah and Bergstrom, 1991; Leath et al., 1993; Ma and Hughes, 1993; Pedersen
and Hughes, 1993; Keller et al., 1994; Orth and Grybauskas, 1994; Azam Parsa and Hughes, 1995;
Shah et al., 1995; Dubin and Rajaram, 1996; Lemerle et al., 1996; Wicki, 1997; Bhathal and
Loughman, 2001; Cao et al., 2001; Murphy et al., 2001)
(Oettler and Schmid, 2000)
(Spadafora and Cole Jr., 1987; Mergoum et al., 2006)
(da Luz and Bergstrom, 1986)
(Sutton, 1920)
(Milus and Chalkley, 1997; Sundin et al., 1999; De Wolf and Francl, 2000; Kim and Bockus, 2003)
(Paillard et al., 2003; Schnurbusch et al., 2003; Tommasini et al., 2007)
(Gaurilčikienė and Ronis, 2006)
(Du et al., 1999; Eyal, 1999; Shah et al., 2000; Cunfer et al., 2001; Krupinsky and Tanaka, 2001;
Mebrate and Cooke, 2001; Shah et al., 2001; Czembor et al., 2003; Fraser et al., 2003; Arseniuk et
al., 2004; Feng et al., 2004; Kim et al., 2004; Xu et al., 2004a; Aguilar et al., 2005; Bennett et al.,
2005; Liu et al., 2005; Liu et al., 2006; Oliver et al., 2006; Singh et al., 2006; Solomon et al., 2006c;
Cowger and Murphy, 2007; Krupinsky et al., 2007; Singh et al., 2007; Ali et al., 2008; Friesen et al.,
2008a; Friesen et al., 2008b; Oliver et al., 2008b; Shankar et al., 2008)
(Engle et al., 2006)
(Liu et al., 2004b)
(Jordan, 1981; Makkar et al., 1995; Huber et al., 1996; Hou and Forman III, 2000)
(Liu et al., 2004a; Oliver et al., 2008a)
(Kimpinski et al., 1987; Kimpinski et al., 1989; Troshina et al., 2007)
Table 8.3: ACNFP library of 13C chemical shifts, carbon assignments, peak
intensities, and calculated ideal natural abundance relative peak intensities (RPI) for
compound standards.
Compound and
Carbon*
Concentration
Alanine 250 mM
Alanine 250 mM
Alanine 250 mM
Arabinose 200 mM
Arabinose 200 mM
Arabinose 200 mM
Arabinose 200 mM
Arabinose 200 mM
Arabinose 200 mM
Arabinose 200 mM
Arabinose 200 mM
Arabinose 200 mM
Arabinose 200 mM
Arabitol 250 mM
Arabitol 250 mM
Arabitol 250 mM
Arabitol 250 mM
Arabitol 250 mM
Arginine 200 mM
Arginine 200 mM
Arginine 200 mM
Arginine 200 mM
Arginine 200 mM
Arginine 200 mM
Asparagine 200 mM
Asparagine 200 mM
Asparagine 200 mM
Asparagine 200 mM
Aspartate 37.6 mM
Aspartate 37.6 mM
Aspartate 37.6 mM
Aspartate 37.6 mM
Citrate 200 mM
Citrate 200 mM
Citrate 200 mM
Citrate 200 mM
Meso-Erythritol 200 mM
Chemical
Shift (ppm)
C1
C2
C3
βC1
αC1
C4
C2
C3
C1
C5
C1
C6
C2
C5
C3
C4
C4
C1
C2
C3
C4
C1
C2
C3
C6OOC1,5OOβC
α,γCH
C2,3
Intensity RPI
(%)
175.70
13364.1
43.62
50.37
23275
75.97
16.02
30637.6
100.00
96.7
7616.3
97.45
92.5
3937.3
50.38
72.38
7442.3
95.23
71.77
7135.5
91.30
68.63
3778
48.34
68.52
4038.2
51.67
68.41
7815.3
100.00
68.36
3833.1
49.05
66.31
6964.8
89.12
62.38
3593.2
45.98
70.83
27632.1
95.41
70.36
27377.8
94.53
70.17
28961.3
100.00
62.95
27804.2
96.00
62.86
28080.6
96.96
174.35
6600.1
69.82
156.76
1990.1
21.05
54.30
8795
93.04
40.49
8133.3
86.04
27.52
9452.5
100.00
23.87
9143.7
96.73
174.39
1739.6
38.22
173.23
2598.2
57.08
51.24
4552
100.00
34.42
4100.8
90.09
174.24
1493.3
66.22
172.83
1172
51.97
50.62
1851.3
82.09
34.52
2255.2
100.00
176.67
3217.8
29.62
173.32
9785.3
90.09
73.2
6467.6
59.54
43.2
10862.1
100.00
71.88
23633.4
100.00
Continued on the following page
306
Table 8.3: (Contd)
Compound and
Carbon*
Concentration
Meso-Erythritol 200 mM
Fructose 250 mM
Fructose 250 mM
Fructose 250 mM
Fructose 250 mM
Fructose 250 mM
Fructose 250 mM
Fructose 250 mM
Fructose 250 mM
Fructose 250 mM
Fructose 250 mM
Fructose 250 mM
Fructose 250 mM
Fructose 6-phosphate 100 mM
Fructose 6-phosphate 100 mM
Fructose 6-phosphate 100 mM
Fructose 6-phosphate 100 mM
Fructose 6-phosphate 100 mM
Fructose 6-phosphate 100 mM
Fructose 6-phosphate 100 mM
Fructose 6-phosphate 100 mM
Fructose 6-phosphate 100 mM
Fructose 6-phosphate 100 mM
Fructose 6-phosphate 100 mM
Fructose 6-phosphate 100 mM
Galactose 200 mM
Galactose 200 mM
Galactose 200 mM
Galactose 200 mM
Galactose 200 mM
Galactose 200 mM
Galactose 200 mM
Galactose 200 mM
Galactose 200 mM
Galactose 200 mM
Galactose 200 mM
Galactose 200 mM
Gluconate 200 mM
Gluconate 200 mM
Gluconate 200 mM
Gluconate 200 mM
Gluconate 200 mM
Chemical
Shift (ppm)
C1,4
αC2
βC2
αC3
βC5
αC5
αC4
βC3
βC4
αC1
βC1
βC6
αC6
βC1
αC1
βC5
βC3
βC2
αC5
αC4
αC3
βC4
αC2
αC6
βC6
-
Intensity RPI
(%)
62.57
22642.3
95.81
101.41
2290.5
28.63
97.99
7586.6
94.83
80.58
2495.6
31.19
75.24
2356.4
29.45
74.33
2400.1
30.00
69.58
7865.1
98.31
69.11
8000.2
100.00
67.44
7339.1
91.74
63.77
6532.2
81.65
63.27
6789.6
84.87
62.52
1968.8
24.61
62.29
2292.1
28.65
104.40
391.9
18.62
101.44
1708.5
81.19
81.75
550.8
26.18
80.01
903.2
42.92
79.90
1004.5
47.74
75.87
357.3
16.98
75.21
2104.2
100.00
74.40
1787.1
84.93
64.27
751.7
35.72
64.21
1035
49.19
62.86
405.7
19.28
62.66
1767.9
84.02
96.39
3143.2
53.77
92.22
5841.5
99.92
75.09
3325.1
56.88
72.74
3001.3
51.34
71.81
3216
55.01
70.42
5846.1
100.00
69.25
5760.6
98.54
69.1
5758.2
98.50
68.68
3136
53.64
68.28
5605
95.88
61.13
5107.7
87.37
60.92
2924.6
50.03
178.59
4887
56.04
74.02
8705.4
99.83
72.51
8578.5
98.38
71.13
8692.7
99.69
70.90
8720
100.00
Continued on the following page
307
Table 8.3: (Contd)
Compound and
Carbon*
Concentration
Gluconate 200 mM
Glucose 200 mM
Glucose 200 mM
Glucose 200 mM
Glucose 200 mM
Glucose 200 mM
Glucose 200 mM
Glucose 200 mM
Glucose 200 mM
Glucose 200 mM
Glucose 200 mM
Glucose 200 mM
Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
[1-13C] Glucose 200 mM
Chemical
Shift (ppm)
βC1
αC1
βC5
βC3
βC2
αC3
αC2
αC5
αC4
βC4
βC6
αC6
βC1 sideband
βC1 sideband
βC1 sideband
βC1 sideband
βC1 sideband
βC1
βC1 sideband
αC1 sideband
αC1
αC1 sideband
βC5
βC3 (split)
βC3 (split)
βC2 (split)
βC2 (split)
αC3
αC2 (split)
αC5
αC2 (split)
αC4
βC4
βC6 (split)
βC6 (split)
αC6 (split)
αC6 (split)
Intensity RPI
(%)
62.56
7845.4
89.97
95.83
8932.9
95.71
92.02
5271.3
56.48
75.87
9187.6
98.44
75.68
8901.2
95.37
74.06
9010.7
96.54
72.69
5402.5
57.88
71.40
5799
62.13
71.36
5467.5
58.58
69.57
5723.9
61.33
69.52
9333.4
100.00
60.67
7436.7
79.68
60.51
4530.2
48.54
108.56
326
15.23
102.3
385.5
18.01
96.76
487.6
22.78
96.36
590.1
27.56
96.13
1578.8
73.75
95.83
190060.8 8878.03
95.52
1201.4
56.12
92.32
856.7
40.02
92.02
114200.3 5334.47
91.71
696.4
32.53
75.87
2140.8
100.00
75.71
1056.5
49.35
75.65
1031.5
48.18
74.35
967.5
45.19
73.74
825.6
38.57
72.68
1183.1
55.26
71.7
558.6
26.09
71.34
783.5
36.60
71.08
523
24.43
69.56
1366.4
63.83
69.52
2091.9
97.72
60.7
948.7
44.32
60.64
951.5
44.45
60.53
610.4
28.51
60.49
602.5
28.14
Continued on the following page
308
Table 8.3: (Contd)
Compound and
Carbon*
Concentration
Glucose 6-phosphate 75 mM
Glucose 6-phosphate 75 mM
Glucose 6-phosphate 75 mM
Glucose 6-phosphate 75 mM
Glucose 6-phosphate 75 mM
Glucose 6-phosphate 75 mM
Glucose 6-phosphate 75 mM
Glucose 6-phosphate 75 mM
Glucose 6-phosphate 75 mM
Glucose 6-phosphate 75 mM
Glucose 6-phosphate 75 mM
Glucose 6-phosphate 75 mM
Glutamate 200 mM
Glutamate 200 mM
Glutamate 200 mM
Glutamate 200 mM
Glutamate 200 mM
Glutamine 200 mM
Glutamine 200 mM
Glutamine 200 mM
Glutamine 200 mM
Glutamine 200 mM
Glycerol 200 mM
Glycerol 200 mM
Histidine 200 mM
Histidine 200 mM
Histidine 200 mM
Histidine 200 mM
Histidine 200 mM
Histidine 200 mM
Inosine 60 mM
Inosine 60 mM
Inosine 60 mM
Inosine 60 mM
Inosine 60 mM
Inosine 60 mM
Inosine 60 mM
Inosine 60 mM
Inosine 60 mM
Inosine 60 mM
Malate 200 mM
Chemical
Shift (ppm)
βC1
αC1
βC5
βC3
αC2
αC3
βC2
αC5
αC4
βC4
βC6
αC6
C5
C1
C2
C4
C3
C5
C1
C2
C4
C3
C2
C1,3
COO(H)
C2H, ring
C5H, ring
C4H, ring
αCH
βCH
COO-
Intensity RPI
(%)
95.93
1773.8
84.68
92.11
1266.9
60.48
75.43
1781.6
85.05
74.78
905.2
43.21
74.68
852.1
40.68
74.00
1840.1
87.85
72.49
1079.5
51.53
71.35
1141.4
54.49
70.35
513.3
24.50
69.05
2094.7
100.00
63.94
754.2
36.01
63.88
1119.1
53.43
181.24
5417.3
60.83
174.49
4948.2
55.56
54.53
7836
87.99
33.38
8888
99.80
26.87
8905.8
100.00
177.55
3507.3
32.79
173.9
3828.3
35.79
54.03
8920.7
83.41
30.72
8536.6
79.82
26.09
10695.2
100.00
72.01
7551.3
52.01
62.42
14519.2
100.00
172.47
4145.8
64.18
133.92
2808.7
43.48
127.31
3574.5
55.34
117.63
5168.2
80.01
53.47
6122.8
94.79
25.68
6459.3
100.00
158.40
1035
29.48
148.36
1396.9
39.79
146.03
2985.5
85.04
140.19
2963
84.40
124.14
1197.8
34.12
88.37
3326.8
94.76
85.52
3307.3
94.21
74.05
3510.6
100.00
70.34
3383.6
96.38
61.28
2945.1
83.89
179.10
4947.7
51.76
Continued on the following page
309
Table 8.3: (Contd)
Compound and
Carbon*
Concentration
Malate 200 mM
Malate 200 mM
Malate 200 mM
Mannitol 1-phosphate 100 mM
Mannitol 1-phosphate 100 mM
Mannitol 1-phosphate 100 mM
Mannitol 1-phosphate 100 mM
Mannitol 1-phosphate 100 mM
Mannitol 1-phosphate 100 mM
Mannitol 1-phosphate 100 mM
Mannitol 1-phosphate 100 mM
Mannitol 200 mM
Mannitol 200 mM
Mannitol 200 mM
[1-13C] Mannitol
[1-13C] Mannitol
[1-13C] Mannitol
[1-13C] Mannitol
[1-13C] Mannitol
[1-13C] Mannitol
[1-13C] Mannitol
[1-13C] Mannitol
[1-13C] Mannitol
Mannose 200 mM
Mannose 200 mM
Mannose 200 mM
Mannose 200 mM
Mannose 200 mM
Mannose 200 mM
Mannose 200 mM
Mannose 200 mM
Mannose 200 mM
Mannose 200 mM
Mannose 200 mM
Methionine 200 mM
Methionine 200 mM
Methionine 200 mM
Methionine 200 mM
Methionine 200 mM
Ornithine 200 mM
Chemical
Shift (ppm)
αCH
βCH
M2
M5
M5
M3 or M4
M3 or M4
M1
M1
M6
C2,5
C3,4
C1,6
C2 (split)
C5
C2 (split)
C3,4 (split)
C3,4 (split)
C1,6 sideband
C1,6 sideband
C1,6
C1,6 sideband
αC1
βC1
βC5
βC3
αC5
βC2
αC2
αC3
αC4
βC4
αC6,βC6
COO(H)
αCH
βCH2
γCH2
S-CH3
C1
Intensity RPI
(%)
176.31
5532.3
57.87
68.48
9559.8
100.00
39.96
9419.1
98.53
71.02
2810
100.00
70.34
1438.1
51.18
70.26
1456.6
51.84
69.32
2804.1
99.79
68.64
2572.4
91.54
65.42
1150.7
40.95
65.36
1191.5
42.40
63.26
2550.3
90.76
70.74
20807.4
100.00
69.18
20545.4
98.74
63.16
18510.7
88.96
71.02
3004.2
40.06
70.75
7498.5
100.00
70.47
3481.6
46.43
69.19
7757.3
103.45
69.17
8035.6
107.16
63.69
1258.6
16.78
63.42
4656.7
62.10
63.16
634706.5 4538.71
62.87
4077.8
54.38
93.97
5273.5
94.40
93.60
2515.2
45.02
76.10
2674.3
47.87
72.98
2543.2
45.52
72.33
5241.6
93.83
71.15
2673.9
47.86
70.61
5233.5
93.68
70.16
5270.3
94.34
66.78
5249.5
93.97
66.54
2683.3
48.03
60.90
5586.4
100.00
174.16
6762.8
58.73
53.82
9733.1
84.52
29.59
10808.4
93.86
28.75
11339.4
98.47
13.86
11515.2
100.00
174.05
2944.3
40.30
Continued on the following page
310
Table 8.3: (Contd)
Compound and
Carbon*
Concentration
Ornithine 200 mM
Ornithine 200 mM
Ornithine 200 mM
Ornithine 200 mM
Phenylalanine 200 mM
Phenylalanine 200 mM
Phenylalanine 200 mM
Phenylalanine 200 mM
Phenylalanine 200 mM
Phenylalanine 200 mM
Phenylalanine 200 mM
Pyruvate 200 mM
Pyruvate 200 mM
Pyruvate 200 mM
Serine 200 mM
Serine 200 mM
Serine 200 mM
Sorbitol 200 MM
Sorbitol 200 MM
Sorbitol 200 MM
Sorbitol 200 MM
Sorbitol 200 MM
Sorbitol 200 MM
Sucrose 200 mM
Sucrose 200 mM
Sucrose 200 mM
Sucrose 200 mM
Sucrose 200 mM
Sucrose 200 mM
Sucrose 200 mM
Sucrose 200 mM
Sucrose 200 mM
Sucrose 200 mM
Sucrose 200 mM
Sucrose 200 mM
Threonine 200 mM
Threonine 200 mM
Threonine 200 mM
Threonine 200 mM
Trehalose 200 mM
Trehalose 200 mM
Trehalose 200 mM
Chemical
Shift (ppm)
C2
C5
C3
C4
COOH
C1, ring
C2,6, ring
C3,5, ring
C4, ring
αCH
βCH
αC=O
COOβCH3
C1
C3
C2
C5
C2
C4
C3
C6
C1
F2
G1
F5
F3
F4
G3
G5
G2
G4
F6
F1
G6
C1
C3
C2
C4
C1
C3
C2
Intensity RPI
(%)
54.08
6298
86.21
38.85
5987.9
81.96
27.37
7305.6
100.00
22.71
7289.4
99.78
173.88
2880.5
29.32
135.03
3040.1
30.95
129.31
9756.2
99.31
129.06
9824.1
100.00
127.64
4728.3
48.13
55.98
4153.6
42.28
36.29
4484.7
45.65
205.09
1841.6
28.28
170.22
1407.6
21.61
26.43
6512.5
100.00
171.87
1038
38.19
59.67
2718.3
100.00
55.9
2133.9
78.50
72.88
11689.1
99.21
71.05
11781.7
100.00
70.93
11691.6
99.24
69.61
11612.2
98.56
62.76
10577.8
89.78
62.37
10662.7
90.50
103.57
9033.7
100.00
92.06
6902.1
76.40
81.25
6679.8
73.94
76.23
6707.2
74.25
73.85
6791.6
75.18
72.44
6834.6
75.66
72.28
7162.7
79.29
70.95
6958.5
77.03
69.08
7040.3
77.93
62.25
5777.4
63.95
61.18
5424.4
60.05
59.97
5541.8
61.35
172.74
16899.4
51.42
65.83
32865.6
100.00
60.37
29513.6
89.80
19.39
32329.4
98.37
93.16
20671
100.00
72.46
20285.4
98.13
72.09
20500.5
99.18
Continued on the following page
311
Table 8.3: (Contd)
Compound and
Carbon*
Concentration
Trehalose 200 mM
C5
Trehalose 200 mM
C4
Trehalose 200 mM
C6
Tryptophan 200 mM
COO(H)
Tryptophan 200 mM
C8, ring
Tryptophan 200 mM
C9, ring
Tryptophan 200 mM
C2H, ring
Tryptophan 200 mM
C5H, ring
Tryptophan 200 mM
C4H, ring
Tryptophan 200 mM
C6H, ring
Tryptophan 200 mM
C7H, ring
Tryptophan 200 mM
C3, ring
Tryptophan 200 mM
αCH
Tryptophan 200 mM
βCH
Xylitol 200 mM
C2,4
Xylitol 200 mM
C3
Xylitol 200 mM
C1,5
* Carbon assignments given as published.
Chemical
Intensity RPI
Shift (ppm)
70.98
69.64
60.47
174.43
136.25
126.56
124.95
122.05
119.38
118.37
111.86
107.39
54.97
26.3
71.86
70.72
62.56
(%)
20201.4
20461.4
16228.3
1699.7
1154.4
1277.7
2152.7
2465
2440.9
2423.1
2388.3
1554.6
2212
2314.3
17475.9
8684.5
15821.8
97.73
98.99
78.51
68.95
46.83
51.83
87.33
100.00
99.02
98.30
96.89
63.07
89.74
93.89
100.00
49.69
90.53
312
Ormonde Waters