The genus Cladosporium and similar dematiaceous ... - CBS - KNAW

Studies in Mycology 58 (2007) 

The genus Cladosporium and similar 

dematiaceous hyphomycetes 

Pedro W. Crous, Uwe Braun, Konstanze Schubert 

and Johannes Z. Groenewald 

Centraalbureau voor Schimmelcultures, 

Utrecht, The Netherlands 

An institute of the Royal Netherlands Academy of Arts and Sciences

The genus Cladosporium and similar dematiaceous hyphomycetes 

Studies in Mycology 58, 2007

Studies in Mycology 

The Studies in Mycology is an international journal which publishes systematic monographs of filamentous fungi and yeasts, and in rare 

occasions the proceedings of special meetings related to all fields of mycology, biotechnology, ecology, molecular biology, pathology and 

systematics. For instructions for authors see www.cbs.knaw.nl. 

Executive Editor 

Prof. dr Robert A. Samson, CBS Fungal Biodiversity Centre, P.O. Box 85167, 3508 AD Utrecht, The Netherlands. 

E-mail: r.samson@cbs.knaw.nl 

Layout Editor 

Manon van den Hoeven-Verweij, CBS Fungal Biodiversity Centre, P.O. Box 85167, 3508 AD Utrecht, The Netherlands. 

E-mail: m.verweij@cbs.knaw.nl 

Scientific Editors 

Prof. dr Uwe Braun, Martin-Luther-Universität, Institut für Biologie, Geobotanik und Botanischer Garten, Neuwerk 21, 

D-06099 Halle, Germany. 

E-mail: uwe.braun@botanik.uni-halle.de 

Prof. dr Pedro W. Crous, CBS Fungal Biodiversity Centre, P.O. Box 85167, 3508 AD Utrecht, The Netherlands. 

E-mail: p.crous@cbs.knaw.nl 

Prof. dr David M. Geiser, Department of Plant Pathology, 121 Buckhout Laboratory, Pennsylvania State University, University Park, PA, U.S.A. 16802. 

E-mail: dgeiser@psu.edu 

Dr Lorelei L. Norvell, Pacific Northwest Mycology Service, 6720 NW Skyline Blvd, Portland, OR, U.S.A. 97229-1309. 

E-mail: llnorvell@pnw-ms.com 

Dr Erast Parmasto, Institute of Zoology & Botany, 181 Riia Street, Tartu, Estonia EE-51014. 

E-mail: e.parmasto@zbi.ee 

Prof. dr Alan J.L. Philips, Faculdade de Ciências e Tecnologia, Universidade Nova de Lisboa, Quinta de Torre, 2829-516 Caparica, Portugal. 

E-mail: alp@mail.fct.unl.pt 

Dr Amy Y. Rossman, Rm 304, Bldg 011A, Systematic Botany & Mycology Laboratory, Beltsville, MD, U.S.A. 20705. 

E-mail: amy@nt.ars-grin.gov 

Dr Keith A. Seifert, Research Scientist / Biodiversity (Mycology and Botany), Agriculture & Agri-Food Canada, KW Neatby Bldg, 960 Carling Avenue, 

Ottawa, ON, Canada K1A OC6. 

E-mail: seifertk@agr.gc.ca 

Prof. dr Jeffrey K. Stone, Department of Botany & Plant Pathology, Cordley 2082, Oregon State University, Corvallis, OR, U.S.A. 97331-2902. 

E-mail: stonej@bcc.orst.edu 

Dr Richard C. Summerbell, 27 Hillcrest Park, Toronto, Ont. M4X 1E8, Canada. 

E-mail: summerbell@aol.com 

Copyright 2007 Centraalbureau voor Schimmelcultures, Fungal Biodiversity Centre, P.O. Box 85167, 3508 AD Utrecht, The Netherlands. 

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Publication date: 31 August 2007 

Published and distributed by CBS Fungal Biodiversity Centre, P.O. Box 85167, 3508 AD Utrecht, The Netherlands. Internet: www.cbs.knaw.nl. 

E-mail: info@cbs.knaw.nl. 

ISBN/EAN : 978-90-70351-61-0 

Online ISSN : 1872-9797 

Print ISSN : 0166-0616 

Cover: Top: page 137, Fig. 30C Cladosporium pseudiridis (CBS 116463), conidiophores and conidia; page 40, Fig. 4D Toxicocladosporium irritans (type 

material), macroconidiophores; page 70, Fig. 11C Ramichloridium musae (CBS 365.36), sympodially proliferating conidiogenous cells, resulting in a long 

conidium-bearing rachis; Bottom: page 19, Fig. 8F Penidiella columbiana (type material), conidiophores with chains of disarticulating conidia; page 120, Fig. 

12C Cladosporium bruhnei (CPC 12211), conidial chains; page 126, Fig. 18L Davidiella tassiana (CPC 12181), asci in culture.

The genus Cladosporium and similar 

dematiaceous hyphomycetes 

edited by 

Pedro W. Crous 

CBS Fungal Biodiversity Centre, P.O. Box 85167, 3508 AD Utrecht, The Netherlands 

Uwe Braun 

Martin-Luther-Universität Institut für Biologie, Geobotanik und Botanischer Garten, Neuwerk 21, D-06099 Halle (Saale), Germany 

Konstanze Schubert 

Botanische Staatssammlung München, Menzinger Strasse 67, D-80638 München, Germany 

and 

Johannes Z. Groenewald 

CBS Fungal Biodiversity Centre, P.O. Box 85167, 3508 AD Utrecht, The Netherlands 

Centraalbureau voor Schimmelcultures, 

Utrecht, The Netherlands 

An institute of the Royal Netherlands Academy of Arts and Sciences

CONTENTS 

P.W. Crous, U. Braun and J.Z. Groenewald: Mycosphaerella is polyphyletic ...................................................................................................1 

P.W. Crous, U. Braun, K. Schubert and J.Z. Groenewald: Delimiting Cladosporium from morphologically similar genera ............................33 

M. Arzanlou, J.Z. Groenewald, W. Gams, U. Braun, H.-D Shin and P.W. Crous: Phylogenetic and morphotaxonomic revision of Ramichloridium 

and allied genera............................................................................................................................................................................................. 57 

K. Schubert, U. Braun, J.Z. Groenewald and P.W. Crous: Cladosporium leaf-blotch and stem rot of Paeonia spp. caused by Dichocladosporium 

chlorocephalum gen. nov.................................................................................................................................................................................95 

K. Schubert, J.Z. Groenewald, U. Braun, J. Dijksterhuis, M. Starink, C.F. Hill, P. Zalar, G.S. de Hoog and P.W. Crous: Biodiversity in 

the Cladosporium herbarum complex (Davidiellaceae, Capnodiales), with standardisation of methods for Cladosporium taxonomy 

and diagnostics............................................................................................................................................................................................. 105 

P. Zalar, G.S. de Hoog, H.-J. Schroers, P.W. Crous, J.Z. Groenewald and N. Gunde-Cimerman: Phylogeny and ecology of the 

ubiquitous saprobe Cladosporium sphaerospermum, with descriptions of seven new species from hypersaline environments 

...................................................................................................................................................................................................................... 157 

P.W. Crous, K. Schubert, U. Braun, G.S. de Hoog, A.D. Hocking, H.-D. Shin and J.Z. Groenewald: Opportunistic, humanpathogenic 

species in the Herpotrichiellaceae are phenotypically similar to saprobic or phytopathogenic species in the 

Venturiaceae ..............................................................................................................................................................................................185 

G.S. de Hoog, A.S. Nishikaku, G. Fernandez Zeppenfeldt, C. Padín-González, E. Burger, H. Badali and A.H.G. Gerrits van den Ende: 

Molecular analysis and pathogenicity of the Cladophialophora carrionii complex, with the description of a novel species.......................... 219 

K.A. Seifert, S.J. Hughes, H. Boulay and G. Louis-Seize: Taxonomy, nomenclature and phylogeny of three cladosporiumlike 

hyphomycetes, Sorocybe resinae, Seifertia azalea and the Hormoconis anamorph of Amorphotheca resinae 

...................................................................................................................................................................................................................... 235

PREFACE 

DEDICATION 

This volume of the Studies in Mycology is dedicated to the memory 

of Gerardus Albertus de Vries (1919–2005), who spent his scientific 

career as mycologist at the CBS, where he was appointed as 

medical mycologist. 

of loyal friends and fellow mycologists dating back to the “Baarnera” 

of CBS. To the very end the CBS received a yearly Christmas 

card, which was always a water colour painting depicting some 

fascinating birds that he happened to be studying at the time. The 

Cladosporium notebooks, annotations and live cultures are still at 

the CBS. It is thus with great joy that we dedicate this volume to 

Gerard de Vries, and build on his Cladosporium legacy, most of 

which is still sporulating, and will be available for scientific debate 

for generations to come. 

The genus Cladosporium and similar dematiaceous 

hyphomycetes 

INTRODUCTION 

Fig. 1. Gerard de Vries studying Cladosporium spp. on different cultural growth 

media. 

On the 10 th of July 1952, Gerard de Vries graduated from the 

University of Utrecht, where he completed his Ph.D. on the topic 

“Contribution to the knowledge of the genus Cladosporium Link 

ex Fr.” under the guidance of the then director of the CBS, Prof. 

dr Johanna Westerdijk. At this time, his thesis (de Vries 1952) 

represented a benchmark and synthesis of our knowledge and 

understanding of Cladosporium spp. studied in culture (Fig. 1). 

Gerard de Vries learned the basics of mycology by working during 

the 1930’s with mushroom taxonomy under the guidance of 

Abraham van Luyk. Via van Luyk he was also introduced to the 

Dutch Mycological Society, who organised excursions, also around 

Baarn, which is where the de Vries family lived. For the rest of his 

career, Gerard would retain his love for studying and collecting 

mushrooms. In 1948 he was employed at CBS under Johanna 

Westerdijk, and given the task of establishing and heading a 

new division called Medical Mycology. This he did, right up to his 

retirement in 1984. For several years de Vries played an important 

role in this discipline, and attended numerous medical congresses 

and workshops, and also published extensively on the topic. De 

Vries loved the outdoors and traveling, and combined this with his 

other passion, which was ornithology (van der Aa 2005). In 1952 

he graduated from the University of Utrecht, producing a revision 

of major species in the genus Cladosporium of importance to the 

medical, industrial and plant pathology disciplines. His book soon 

became highly popular, and with a strong demand for additional 

copies, resulting in it being reprinted by J. Cramer. His contribution 

to Cladosporium was also recently acknowledged with the 

introduction of the cladosporium-like heat-tolerant genus, Devriesia 

by Seifert et al. (2004). Additional books published by de Vries dealt 

with mushrooms for amateur mycologists (1955) and a treatment of 

Hypogaea and truffels published in 1971 as part 3 of the “Fungi of 

the Netherlands” (van der Aa 2005). 

Gerard de Vries was a well tempered, softly spoken man, 

who avoided conflicts, except when it dealt with scientific issues. 

He never married, and died a bachelor, surrounded by a circle 

Species of Cladosporium are common and widespread, and interact 

with humans in every phase of life, from growing behind your bed 

or bedroom cupboard and producing allergens, or growing on the 

bathroom ceiling, to the fruit decay happening in the fruit basket in 

the kitchen, to colonising the debris lying outside your house, and 

even the plant diseases observed on some of the shrubs, trees, or 

flowers cultivated in your garden. However, scientists generally shy 

away from trying to identify these similar looking organisms, and 

therefore the main aims of this volume were to: 

1) Establish standardised conditions and protocols for studying 

cladosporioid species and their teleomorphs in culture; 

2) Determine how they can morphologically be distinguished in 

culture, and highlight important diagnostic features; 

3) Circumscribe the genus, and delineate it from morphologically 

similar dematiaceous hyphomycetes; 

4) Determine which DNA gene loci are informative to accurately 

distinguish species of Cladosporium, and initiate a database of 

Cladosporium sequences that can in future be used to set up an 

online polyphasic identification key. 

Although Cladosporium is one of the largest and most 

heterogeneous genera of hyphomycetes, currently comprising 

more than 772 names (Dugan et al. 2004), only a mere fraction of 

these species are known from culture, and thus the real number 

of taxa that exist remains unknown. Species of Cladosporium are 

commonly encountered on plant and other kinds of debris, frequently 

colonising lesions of plant pathogenic fungi, and are also isolated 

from air, soil, food, paint, textiles and other organic matters (Ellis 

1971, 1976; Schubert 2005a), they are also common endophytes 

(Brown et al. 1998, El-Morsy 2000) as well as phylloplane fungi 

(Islam & Hasin 2000, de Jager et al. 2001, Inacio et al. 2002, 

Stohr & Dighton 2004, Levetin & Dorsey 2006). Some species of 

Cladosporium have a medical relevance in clinical laboratories, 

and also cause allergic lung mycoses (de Hoog et al. 2000). In 

spite of its obvious importance, species of Cladosporium are still 

poorly understood. 

Taxonomy of the anamorph 

The first binominal introduced for this group of fungi was that of 

Dematium herbarum Pers. (Persoon 1794) (Fig. 2). Cladosporium 

herbarum (Pers.) Link was subsequently selected to serve as

lectotype for the genus by Clements & Shear (1931), a proposal 

which was accepted by de Vries (1952), Hughes (1958), and others 

(Prasil & de Hoog 1988). In subsequent years the number of taxa 

described in the genus grew rapidly, though the generic concept 

was rather vague. As a consequence, numerous morphologically 

similar dematiaceous hyphomycetes with catenulate conidia were 

incorrectly assigned to Cladosporium, making this one of the largest 

genera of hyphomycetes. 

In an attempt to circumscribe some of the more well-known taxa, 

de Vries (1952) published a revision of nine Cladosporium species in 

vivo and in vitro, and 13 additional taxa in an appendix. Ellis (1971, 

1976) described and illustrated 43 species, while Morgan-Jones 

and McKemy dealt with selected species in the series “Studies in 

the genus Cladosporium s. lat.” (Morgan-Jones & McKemy 1990, 

McKemy & Morgan-Jones 1990, 1991a–c). Other significant works 

that also treated Cladosporium species include Ho et al. (1999), 

and Zhang et al. (2003), though these authors still followed a 

wider generic concept. David (1997) followed the taxonomy of de 

Vries (1952), who first considered Heterosporium as a synonym 

of Cladosporium, and introduced the combination Cladosporium 

subgen. Heterosporium. Subsequent to this publication, further 

monographic studies on the genus Cladosporium s. lat. were 

initiated by Braun and co-workers (Braun et al. 2003, 2006, Dugan 

et al. 2004, Schubert & Braun 2004, 2005a, b, 2006, 2007, Schubert 

2005a, b, Heuchert et al. 2005). 

Treatments of human pathogenic Cladosporium species 

(Masclaux et al. 1995, Untereiner 1997, Gerrits van den Ende & de 

Hoog 1999, Untereiner & Naveau 1999, Untereiner et al. 1999; de 

Hoog et al. 2000), concluded that they represent species belonging 

to the Herpotrichiellaceae (Capronia Sacc./Cladophialophora 

Borelli). Saprobic species, which appear morphologically similar, 

were found to belong to the Venturiaceae (Caproventuria U. Braun/ 

Pseudocladosporium U. Braun; Braun et al. 2003, Schubert et al. 

2003, Beck et al. 2005) (see Crous et al. 2007 – this volume). Further 

genera that were separated from Cladosporium include Sorocybe 

resinae (Fr.) Fr. [≡ Cladosporium resinae (Lindau) G.A. de Vries, 

teleomorph: Amorphotheca resinae Parbery; Partridge & Morgan- 

Jones 2002] (see Seifert et al. 2007 – this volume), Devriesia Seifert 

& N.L. Nickerson, erected for heat tolerant species (Seifert et al. 

2004), Cladoriella Crous, erected for saprobic species (Crous et 

al. 2006b), Metulocladosporiella Crous, Schroers, Groenewald, U. 

Braun & K. Schub., erected for the causal agent of banana speckle 

disease (Crous et al. 2006a), Digitopodium U. Braun, Heuchert & 

K. Schub. and Parapericoniella U. Braun, Heuchert & K. Schub., 

Fig. 2. Type specimen of Dematium herbarum Pers. (1794), preserved in the 

National Herbarium of the Netherlands in Leiden. 

representing two genera of hyperparasitic hyphomycetes (Heuchert 

et al. 2005). 

Taxonomy of the teleomorph 

Teleomorphs of Cladosporium have traditionally been described in 

Mycosphaerella Johanson. The first indication that this may not be 

the case was the rDNA ITS sequence data presented by Crous et 

al. (2001), which revealed cladosporium-like taxa to cluster basal 

to Mycosphaerella s. str. This finding was further strengthened by 

adding 18S rDNA data, which clearly distinguished the Cladosporium 

clade from Mycosphaerella (Braun et al. 2003). These results 

lead to the erection of the genus Davidiella Crous & U. Braun for 

teleomorphs of Cladosporium, though it was largely established 

based on its unique anamorphs, rather than distinct teleomorph 

features. In a revision of the genus Mycosphaerella, Aptroot (2006) 

provided the first clear morphological characteristics to distinguish 

Davidiella from Mycosphaerella, referring to their sole-shaped 

ascospores, and angular lumina that are to be seen in Davidiella 

ascospores. Further phylogenetic evidence for the distinction was 

found by Schoch et al. (2006), which led to the erection of the 

family Davidiellaceae (Capnodiales). Detailed cultural studies of 

Davidiella teleomorphs, however, were still lacking (see Schubert 

et al. 2007 – this volume). 

What is Cladosporium? 

David (1997) provided the first modern concept of Cladosporium by 

conducting comprehensive scanning electron microscopic (SEM) 

examinations of the scar and hilum structure in Cladosporium and 

Heterosporium, thereby confirming the observations of Roquebert 

(1981). He introduced the term “coronate” for the Cladosporium 

scar type, which is characterised by having a central convex part 

(dome), surrounded by a raised periclinal rim (Fig. 3), and proved 

that these anamorphs are linked to teleomorphs now placed in 

Davidiella (see David 1997, fig. 12). 

This new concept of Cladosporium s. str. and Davidiella 

(David 1997, Braun et al. 2003, Aptroot 2006), supported by 

morphological and molecular data, rendered it possible to initiate 

a comprehensive revision of the genus. The first step was the 

preparation of a general, annotated check-list of Cladosporium 

names (Dugan et al. 2004), followed by revisions of fungicolous 

(Heuchert et al. 2005) and foliicolous species of Cladosporium s. 

lat. (Schubert 2005b, Braun et al. 2006, Schubert & Braun 2004, 

2005a, b, 2006, 2007). The present study is the first to integrate 

these concepts on cladosporioid species in culture, in an attempt to 

further elucidate species of Cladosporium, and delineate the genus 

from other, morphologically similar dematiaceous genera that have 

traditionally been confused with Cladosporium s. str. 

How natural should anamorph genera be? 

Article 59 of the International Code of Botanical Nomenclature 

was introduced to enable mycologists to name the asexual states 

of fungi that they encountered, and for which no teleomorph 

association was known. It was and remains a completely artificial 

system, complicated further by the evolution of the same anamorph 

morphology in different families, and even orders. In 1995 Gams 

discussed “How natural should anamorph genera be”, concluding 

that paraphyletic genera should be an acceptable option, and that 

anamorphs cannot reflect natural relationships. In a special volume 

dedicated at integrating molecular data and morphology, Seifert et 

al. (2000) proposed using anamorph names as adjectives, e.g., 

acremonium-like, when they clustered in different clades, or were 

linked to different teleomorphs than the type species of the genus

Fig. 3. Coronate scar structure of Cladosporium herbaroides, visible by means of 

Scanning Electron Microscopy. Scale bar = 5 µm (Photo: Jan Dijksterhuis). 

Acremonium. In a phylogenetic study of the Herpotrichiellaceae, 

Haase et al. (1999) proposed to accept anamorphs as poly- and 

paraphyletic within the order Chaetothyriales, as their taxonomy 

was unsupported by phylogeny, and Cook et al. (1997) as well as 

Braun et al. (2002) followed this methodology in naming anamorphs 

of the Erysiphaceae (Erysiphales). After much debate, we have 

chosen to use the same approach in this volume, and will refrain 

from introducing different anamorph genera for the same phenotype 

clustering within different clades of the same order. It is hoped that 

this approach will stop the unnecessary proliferation of names, until 

we can move to a single nomenclature for ascomycetous fungi. 

Anamorphs are just form taxa, established for the sole purpose 

of enabling mycologists to name asexual states that occur in 

the absence of their teleomorphs. Anamorph genera are simply 

phenotypic concepts that lack phylogenetic relevance within the 

order. 

REFERENCES 

Aa HA van der (2005). In memoriam Gerard de Vries (1919–2005). Coolia 48(4): 

173–175. 

Aptroot A (2006). Mycosphaerella and its anamorphs: 2. Conspectus of 

Mycosphaerella. CBS Biodiversity Series 5: 1–231. 

Beck A, Ritschel A, Schubert K, Braun U, Triebel D (2005). Phylogenetic relationship 

of the anamorphic genus Fusicladium s. lat. as inferred by ITS data. Mycological 

Progress 4: 111–116. 

Braun U, Cook RTA, Inman AJ, Shin HD (2002). The taxonomy of the powdery 

mildew fungi. In: The powdery mildews, a comprehensive treatise (Bélanger 

RR, Bushnell WR, Dik AJ, Carver TLW, eds). APS Press, St. Paul, U.S.A.: 

13–55. 

Braun U, Crous PW, Dugan FM, Groenewald JZ, Hoog GS de (2003). Phylogeny 

and taxonomy of cladosporium-like hyphomycetes, including Davidiella gen. 

nov., the teleomorph of Cladosporium s. str. Mycological Progress 2: 3–18. 

Braun U, Hill CF, Schubert K (2006). New species and new records of biotrophic 

micromycetes from Australia, Fiji, New Zealand and Thailand. Fungal Diversity 

22: 13–35. 

Brown KB, Hyde KD, Guest DI (1998). Preliminary studies on endophytic fungal 

communities of Musa acuminata species complex in Hong Kong and Australia. 

Fungal Diversity 1: 27–51. 

Clements FE, Shear CL (1931). The Genera of Fungi. H.W. Wilson, New York. 

Cook RTA, Inman AJ, Billings C (1997). Identification and classification of powdery 

mildew anamorphs using light and scanning electron microscopy and host 

range data. Mycological Research 101: 975–1002. 

Crous PW, Kang JC, Braun U (2001). A phylogenetic redefinition of anamorph 

genera in Mycosphaerella based on ITS rDNA sequences. Mycologia 93: 

1081–1101. 

Crous PW, Schroers HJ, Groenewald JZ, Braun U, Schubert K (2006a). 

Metulocladosporiella gen. nov. for the causal organism of Cladosporium 

speckle disease of banana. Mycological Research 110: 264–275. 

Crous PW, Schubert K, Braun U, de Hoog GS, Hocking AD, Shin H-D, Groenewald 

JZ (2007). Opportunistic, human-pathogenic species in the Herpotrichiellaceae 

are phenotypically similar to saprobic or phytopathogenic species in the 

Venturiaceae. Studies in Mycology 58: 185–217. 

Crous PW, Verkley GJM, Groenewald JZ (2006b). Eucalyptus microfungi known 

from culture. 1. Cladoriella and Fulvoflamma genera nova, with notes on some 

other poorly known taxa. Studies in Mycology 55: 53–63. 

David JC (1997). A contribution to the systematics of Cladosporium. Revision of the 

fungi previously referred to Heterosporium. Mycological Papers 172: 1–157. 

Dugan FM, Schubert K, Braun U (2004). Check-list of Cladosporium names. 

Schlechtendalia 11: 1–103. 

El-Morsy EM (2000). Fungi isolated from the endorhizosphere of halophytic plants 

from the Red Sea Coast of Egypt. Fungal Diversity 5: 43–54. 

Ellis MB (1971). Dematiaceous Hyphomycetes. Commonwealth Mycological 

Institute, Kew. 

Ellis MB (1976). More Dematiaceous Hyphomycetes. Commonwealth Mycological 


Gams W (1995). How natural should anamorph genera be? Canadian Journal of 

Botany 73: S747–S753. 

Gerrits van den Ende AHG, Hoog GS de (1999). Variability and molecular diagnostics 

of the neurotrophic species Cladophialophora bantiana. Studies in Mycology 

43: 151–162. 

Haase G, Sonntag L, Melzer-Krick B, Hoog GS de (1999). Phylogenetic inference 

by SSU-gene analysis of members of the Herpotrichiellaceae with special 

reference to human pathogenic species. Studies in Mycology 43: 80–97. 

Heuchert B, Braun U, Schubert K (2005). Morphotaxonomic revision of fungicolous 

Cladosporium species (hyphomycetes). Schlechtendalia 13: 1–78. 

Ho MH-M, Castañeda RF, Dugan FM, Jong, SC (1999). Cladosporium and 

Cladophialophora in culture: descriptions and an expanded key. Mycotaxon 

72: 115–157. 

Hoog GS de, Guarro, J, Gené J, Figueras MJ (2000). Atlas of clinical fungi. 2 nd ed. 

Centralbureau voor Schimmelcultures, Utrecht and Universitat rovira I virgili, 

Reus. 

Hughes SJ (1958). Revisiones hyphomycetum aliquot cum appendice de nominibus 

rejiciendis. Canadian Journal of Botany 36: 727–836. 

Inacio J, Pereira P, de Cavalho M, Fonseca A, Amaral-Collaco MT, Spencer-Martins 

I (2002). Estimation and diversity of phylloplane mycobiota on selected plants in 

a Mediterranean-type ecosystem in Portugal. Microbial Ecology 44: 344–353. 

Islam M, Hasin F (2000). Studies on phylloplane mycoflora of Amaranthus viridis L. 

National Academy Science Letters, India 23: 121–123. 

Jager ES de, Wehner FC, Karsten L (2001). Microbial ecology of the mango 

phylloplane. Microbial Ecology 42: 201–207. 

Levetin E, Dorsey K (2006). Contribution to leaf surface fungi to the air spora. 

Aerobiologia 22: 3–12. 

Masclaux F, Guého E, Hoog GS de, Christen R (1995). Phylogenetic relationship of 

human-pathogenic Cladosporium (Xylohypha) species. Journal of Medical and 

Veterinary Mycology 33: 327–338. 

McKemy JM, Morgan-Jones G (1990). Studies in the genus Cladosporium sensu 

lato II. Concerning Heterosporium gracile, the causal organism of leaf spot 

disease of Iris species. Mycotaxon 39: 425–440. 

McKemy JM, Morgan-Jones G (1991a). Studies in the genus Cladosporium 

sensu lato III. Concerning Cladosporium chlorocephalum and its synonym 

Cladosporium paeoniae, the causal organism of leaf-blotch of peony. 

Mycotaxon 41: 135–146. 

McKemy JM, Morgan-Jones G (1991b). Studies in the genus Cladosporium sensu 

lato IV. Concerning Cladosporium oxysporum, a plurivorous predominantly 

saprophytic species in warm climates. Mycotaxon 41: 397–405. 

McKemy JM, Morgan-Jones G (1991c). Studies in the genus Cladosporium sensu 

lato V. Concerning the type species Cladosporium herbarum. Mycotaxon 42: 

307–317. 

Morgan-Jones G, McKemy JM (1990). Studies in the genus Cladosporium sensu 

lato I. Concerning Cladosporium uredinicola, occurring on telial columns of 

Cronartium quercuum and other rusts. Mycotaxon 39: 185–202. 

Partridge EC, Morgan-Jones G (2002). Notes on hyphomycetes, LXXXVIII. New 

genera in which to classify Alysidium resinae and Pycnostysanus azaleae, with 

a consideration of Sorocybe. Mycotaxon 83: 335–352. 

Persoon CH (1794). Neuer Versuch einer systematischen Eintheilung der 

Schwämme. Neues Magazin fur die Botanik 1: 63–128. 

Prasil K, Hoog GS de (1988). Variability in Cladosporium herbarum. Transactions of 

the British Mycological Society 90: 49–54. 

Roquebert MF (1981). Analyse des phénoménes pariétaux au cours de la 

conidiogenése chez quelques champignon microscopiques. Memoires du 

Museum d’Histoire Naturelle, Sér B, Botanique 28: 3–79. 

Schoch C, Shoemaker RA, Seifert KA, Hambleton S, Spatafora JW, Crous PW 

(2006). A multigene phylogeny of the Dothideomycetes using four nuclear loci. 

Mycologia 98: 1041–1052.

Schubert K (2005a). Morphotaxonomic revision of foliicolous Cladosporium species 

(hyphomycetes). PhD thesis, Martin-Luther-University, Halle. 

Schubert K (2005b). Taxonomic revision of the genus Cladosporium s. lat. 3. A 

revision of Cladosporium species described by J.J. Davis and H.C. Greene 

(WIS). Mycotaxon 92: 55–76. 

Schubert K, Braun U (2004). Taxonomic revision of the genus Cladosporium s. lat. 2. 

Morphotaxonomic examination of Cladosporium species occurring on hosts of 

the families Bignoniaceae and Orchidaceae. Sydowia 56: 76–97. 

Schubert K, Braun U (2005a). Taxonomic revision of the genus Cladosporium 

s. lat. 1. Species reallocated to Fusicladium, Parastenella, Passalora, 

Pseudocercospora and Stenella. Mycological Progress 4: 101–109. 

Schubert K, Braun U (2005b). Taxonomic revision of the genus Cladosporium s. 

lat. 4. Species reallocated to Asperisporium, Dischloridium, Fusicladium, 

Passalora, Pseudasperisporium and Stenella. Fungal Diversity 20: 187–208. 


Validation and description of new species. Schlechtendalia 14: 55–83. 

Schubert K, Braun U (2007). Taxonomic revision of the genus Cladosporium s. lat. 

6. New species, reallocations to and synonyms of Cercospora, Fusicladium, 

Passalora, Septonema and Stenella. Nova Hedwigia 84(1–2): 189–208. 

Schubert K, Groenewald JZ, Braun U, Dijksterhuis J, Starink M, Hill CF, Zalar P, 

Hoog GS de, Crous PW (2007). Biodiversity in the Cladosporium herbarum 

complex (Davidiellaceae, Capnodiales), with standardisation of methods for 

Cladosporium taxonomy and diagnostics. Studies in Mycology 58: 105–156. 

Schubert K, Ritschel A, Braun U (2003). A monograph of Fusicladium s. lat. 

(hyphomycetes). Schlechtendalia 9: 1–132. 

Seifert KA, Gams W, Crous PW, Samuels GJ (2000). Molecules, morphology and 

classification: Towards monophyletic genera in the Ascomycetes. Studies in 

Mycology 45: 1–230. 

Seifert KA, Hughes SJ, Boulay H, Louis-Seize G (2007). Taxonomy, nomenclature 

and phylogeny of three cladosporium-like hyphomycetes, Sorocybe resinae, 

Seifertia azalea and the Hormoconis anamorph of Amorphotheca resinae. 

Studies in Mycology 58: 235–245. 

Seifert KA, Nickerson NL, Corlett M, Jackson ED, Lois-Seize G, Davies RJ (2004). 

Devriesia, a new hyphomycete genus to accommodate heat-resistant, 

cladosporium-like fungi. Canadian Journal of Botany 82: 914–926. 

Stohr SN, Dighton J (2004). Effects of species diversity on establishment and 

coexistence: A phylloplane fungal community model system. Microbial Ecology 

48: 431–438. 

Untereiner WA (1997). Taxonomy of selected members of the ascomycete genus 

Capronia with notes on anamorph-teleomorph connection. Mycologia 89: 

120–131. 

Untereiner WA, Gerrits van den Ende AHG, Hoog GA de (1999). Nutritional 

physiology of species of Capronia. Studies in Mycology 43: 98–106. 

Untereiner WA, Naveau FA (1999). Molecular systematic of the Herpotrichiellaceae 

with an assessment of the phylogenetic position of Exophiala dermatitidis and 

Phialophora americana. Mycologia 91: 67–83. 

Vries GA de (1952). Contribution to the Knowledge of the Genus Cladosporium Link 

ex Fr. Centraalbureau voor Schimmelcultures, Baarn. 

Vries GA de (1955). Paddestoelen (en schimmels). WJ Thieme & Cie. Zutputen. 

Vries GA de (1971). De Fungi van Nederland. 3. Hypogaea. Wetenschappelijke 

mededelingen van de Koninklijke Nederlandse Natuurhistorische Vereniging. 

Hoogwoud. 

Zhang ZY, Liu YL, Zhang T, Li TF, Wang G, Zhang H, He YH, Peng HH (2003). 

Cladosporium, Fusicladium, Pyricularia. Flora Fungorum Sinicorum 14: 1– 

297. 

The Editors 1 July 2007 

Propositions for this volume: 

“Evolution gives rise to lineages, which we try to recognise as genera and species” 

“The most interesting fungi are those isolated by accident”

available online at www.studiesinmycology.org 

doi:10.3114/sim.2007.58.01 

Studies in Mycology 58: 1–32. 2007. 

Mycosphaerella is polyphyletic 

P.W. Crous 1* , U. Braun 2 and J.Z. Groenewald 1 

1 

CBS Fungal Biodiversity Centre, P.O. Box 85167, 3508 AD, Utrecht, The Netherlands; 2 Martin-Luther-Universität, Institut für Biologie, Geobotanik und Botanischer Garten, 

Herbarium, Neuwerk 21, D-06099 Halle, Germany 

*Correspondence: Pedro W. Crous, p.crous@cbs.knaw.nl 

Abstract: Mycosphaerella, one of the largest genera of ascomycetes, encompasses several thousand species and has anamorphs residing in more than 30 form genera. 

Although previous phylogenetic studies based on the ITS rDNA locus supported the monophyly of the genus, DNA sequence data derived from the LSU gene distinguish several 

clades and families in what has hitherto been considered to represent the Mycosphaerellaceae. Several important leaf spotting and extremotolerant species need to be disposed 

to the genus Teratosphaeria, for which a new family, the Teratosphaeriaceae, is introduced. Other distinct clades represent the Schizothyriaceae, Davidiellaceae, Capnodiaceae, 

and the Mycosphaerellaceae. Within the two major clades, namely Teratosphaeriaceae and Mycosphaerellaceae, most anamorph genera are polyphyletic, and new anamorph 

concepts need to be derived to cope with dual nomenclature within the Mycosphaerella complex. 

Taxonomic novelties: Batcheloromyces eucalypti (Alcorn) Crous & U. Braun, comb. nov., Catenulostroma Crous & U. Braun, gen. nov., Catenulostroma abietis (Butin & 

Pehl) Crous & U. Braun, comb. nov., Catenulostroma chromoblastomycosum Crous & U. Braun, sp. nov., Catenulostroma elginense (Joanne E. Taylor & Crous) Crous & U. 

Braun, comb. nov., Catenulostroma excentricum (B. Sutton & Ganap.) Crous & U. Braun, comb. nov., Catenulostroma germanicum Crous & U. Braun, sp. nov., Catenulostroma 

macowanii (Sacc.) Crous & U. Braun, comb. nov., Catenulostroma microsporum (Joanne E. Taylor & Crous) Crous & U. Braun, comb. nov., Catenulostroma protearum (Crous 

& M.E. Palm) Crous & U. Braun, comb. nov., Penidiella Crous & U. Braun, gen. nov., Penidiella columbiana Crous & U. Braun, sp. nov., Penidiella cubensis (R.F. Castañeda) U. 

Braun, Crous & R.F. Castañeda, comb. nov., Penidiella nectandrae Crous, U. Braun & R.F. Castañeda, nom. nov., Penidiella rigidophora Crous, R.F. Castañeda & U. Braun, sp. 

nov., Penidiella strumelloidea (Milko & Dunaev) Crous & U. Braun, comb. nov., Penidiella venezuelensis Crous & U. Braun, sp. nov., Readeriella blakelyi (Crous & Summerell) 

Crous & U. Braun, comb. nov., Readeriella brunneotingens Crous & Summerell, sp. nov., Readeriella considenianae (Crous & Summerell) Crous & U. Braun, comb. nov., 

Readeriella destructans (M.J. Wingf. & Crous) Crous & U. Braun, comb. nov., Readeriella dimorpha (Crous & Carnegie) Crous & U. Braun, comb. nov., Readeriella epicoccoides 

(Cooke & Massee) Crous & U. Braun, comb. nov., Readeriella gauchensis (M.-N. Cortinas, Crous & M.J. Wingf.) Crous & U. Braun, comb. nov., Readeriella molleriana (Crous 

& M.J. Wingf.) Crous & U. Braun, comb. nov., Readeriella nubilosa (Ganap. & Corbin) Crous & U. Braun, comb. nov., Readeriella pulcherrima (Gadgil & M. Dick) Crous & 

U. Braun, comb. nov., Readeriella stellenboschiana (Crous) Crous & U. Braun, comb. nov., Readeriella toledana (Crous & Bills) Crous & U. Braun, comb. nov., Readeriella 

zuluensis (M.J. Wingf., Crous & T.A. Cout.) Crous & U. Braun, comb. nov., Teratosphaeria africana (Crous & M.J. Wingf.) Crous & U. Braun, comb. nov., Teratosphaeria alistairii 

(Crous) Crous & U. Braun, comb. nov., Teratosphaeria associata (Crous & Carnegie) Crous & U. Braun, comb. nov., Teratosphaeria bellula (Crous & M.J. Wingf.) Crous & U. 

Braun, comb. nov., Teratosphaeria cryptica (Cooke) Crous & U. Braun, comb. nov., Teratosphaeria dentritica (Crous & Summerell) Crous & U. Braun, comb. nov., Teratosphaeria 

excentrica (Crous & Carnegie) Crous & U. Braun, comb. nov., Teratosphaeria fimbriata (Crous & Summerell) Crous & U. Braun, comb. nov., Teratosphaeria flexuosa (Crous 

& M.J. Wingf.) Crous & U. Braun, comb. nov., Teratosphaeria gamsii (Crous) Crous & U. Braun, comb. nov., Teratosphaeria jonkershoekensis (P.S. van Wyk, Marasas & 

Knox-Dav.) Crous & U. Braun, comb. nov., Teratosphaeria maxii (Crous) Crous & U. Braun, comb. nov., Teratosphaeria mexicana (Crous) Crous & U. Braun, comb. nov., 

Teratosphaeria molleriana (Thüm.) Crous & U. Braun, comb. nov., Teratosphaeria nubilosa (Cooke) Crous & U. Braun, comb. nov., Teratosphaeria ohnowa (Crous & M.J. Wingf.) 

Crous & U. Braun, comb. nov., Teratosphaeria parkiiaffinis (Crous & M.J. Wingf.) Crous & U. Braun, comb. nov., Teratosphaeria parva (R.F. Park & Keane) Crous & U. Braun, 

comb. nov., Teratosphaeria perpendicularis (Crous & M.J. Wingf.) Crous & U. Braun, comb. nov., Teratosphaeria pluritubularis (Crous & Mansilla) Crous & U. Braun, comb. 

nov., Teratosphaeria pseudafricana (Crous & T.A. Cout.) Crous & U. Braun, comb. nov., Teratosphaeria pseudocryptica (Crous) Crous & U. Braun, comb. nov., Teratosphaeria 

pseudosuberosa (Crous & M.J. Wingf.) Crous & U. Braun, comb. nov., Teratosphaeria quasicercospora (Crous & T.A. Cout.) Crous & U. Braun, comb. nov., Teratosphaeria 

readeriellophora (Crous & Mansilla) Crous & U. Braun, comb. nov., Teratosphaeria secundaria (Crous & Alfenas) Crous & U. Braun, comb. nov., Teratosphaeria stramenticola 

(Crous & Alfenas) Crous & U. Braun, comb. nov., Teratosphaeria suberosa (Crous, F.A. Ferreira, Alfenas & M.J. Wingf.) Crous & U. Braun, comb. nov., Teratosphaeria suttonii 

(Crous & M.J. Wingf.) Crous & U. Braun, comb. nov., Teratosphaeria toledana (Crous & Bills) Crous & U. Braun, comb. nov., Teratosphaeriaceae Crous & U. Braun, fam. nov. 

Key words: Ascomycetes, Batcheloromyces, Colletogloeopsis, Readeriella, Teratosphaeria, Trimmatostroma, DNA sequence comparisons, systematics. 

Introduction 

The genus Mycosphaerella Johanson as presently circumscribed 

contains close to 3 000 species (Aptroot 2006), excluding its 

anamorphs, which represent thousands of additional species 

(Crous et al. 2000, 2001, 2004a, b, 2006a, b, 2007b, Crous & 

Braun 2003). Crous (1998) predicted that Mycosphaerella would 

eventually be split according to its anamorph genera, and Crous 

et al. (2000) recognised six sections, as originally defined by Barr 

(1972). This was followed by a set of papers (Crous et al. 2001, 

Goodwin et al. 2001), where it was concluded, based on ITS DNA 

sequence data, that Mycosphaerella was monophyletic. A revision 

of the various coelomycete and hyphomycete anamorph concepts 

led Crous & Braun (2003) to propose a system whereby the asexual 

morphs could be allocated to various form genera affiliated with 

Mycosphaerella holomorphs. 

In a recent study that formed part of the US “Assembling 

the Fungal Tree of Life” project, Schoch et al. (2006) were able 

to show that the Mycosphaerellaceae represents a family within 

Capnodiales. Furthermore, some variation was also depicted within 

the family, which supported similar findings in other recent papers 

employing LSU sequence data, such as Hunter et al. (2006), and 

Batzer et al. (2007). To further elucidate the phylogenetic variation 

observed within the Mycosphaerellaceae in these studies, a 

subset of isolates was selected for the present study, representing 

the various species recognised as morphologically distinct from 

Mycosphaerella s. str. 

The genus Mycosphaerella has in recent years been linked 

to approximately 30 anamorph genera (Crous & Braun 2003, 

Crous et al. 2007b). Many of these anamorph genera resulted 

from a reassessment of cercosporoid forms. Chupp (1954) was 

of the opinion that they all represented species of the genus 

Cercospora Fresen., although he clearly recognised differences in 

their morphology. In a series of papers by Deighton, as well as 

others such as Sutton, Braun and Crous, the genus Cercospora 

was delimited based on its type species, Cercospora penicillata 

(Ces.) Fresen., while taxa formerly included in the genus by Chupp 

(1954) but differing in conidiophore arrangement, conidiogenesis, 

pigmentation, conidial catenulation, septation, and scar/hilum 

structure were allocated to other genera. Similar studies in which 

the type species were recollected and subjected to DNA sequence

Crous et al. 

Table 1. Isolates for which new sequences were generated. 

Anamorph Teleomorph Accession number 1 Host Country Collector GenBank Accession number 

Batcheloromyces eucalypti CBS 313.76; CPC 3632 Eucalyptus tessellaris Australia J.L. Alcorn EU019245 

Batcheloromyces leucadendri CBS 110892; CPC 1837 Leucadendron sp. South Africa L. Swart EU019246 

Batcheloromyces proteae CBS 110696; CPC 1518 Protea cynaroides South Africa L. Viljoen EU019247 

Capnobotryella renispora CBS 214.90*; CBS 176.88; IAM 13014; JCM 6932 Capnobotrys neessii Japan J. Sugiyama EU019248 

Catenulostroma abietis CBS 290.90 Man, skin lesion Netherlands R.G.F. Wintermans EU019249 

Catenulostroma castellanii CBS 105.75*; ATCC 24788 Man, tinea nigra Venezuela — EU019250 

Catenulostroma chromoblastomycosum CBS 597.97 Man, chromoblastomycosis Zaire V. de Brouwere EU019251 

Catenulostroma elginense CBS 111030; CPC 1958 Protea grandiceps South Africa J.E. Taylor EU019252 

Catenulostroma germanicum CBS 539.88 Stone Germany — EU019253 

Catenulostroma macowanii CBS 110756; CPC 1872 Protea nitida South Africa J.E. Taylor EU019254 

Catenulostroma microsporum Teratosphaeria microspora CBS 110890; CPC 1832 Protea cynaroides South Africa L. Swart EU019255 

Catenulostroma sp. Teratosphaeria 

CBS 118911; CPC 12085 Eucalyptus sp. Uruguay M.J. Wingfield EU019256 

pseudosuberosa 

Cercosporella centaureicola CBS 120253 Centaurea solstitiales Greece D. Berner EU019257 

Cibiessia dimorphospora CBS 120034; CPC 12636 Eucalyptus nitens Australia — EU019258 

Cibiessia minutispora CPC 13071* Eucalyptus henryii Australia A.J. Carnegie EU019259 

Cibiessia nontingens Teratosphaeria sp. CBS 120725*; CPC 13217 Eucalyptus tereticornis Australia B. Summerell EU019260 

Cladosporium bruhnei Davidiella allicina CBS 115683; ATCC 66670; CPC 5101 CCA-treated Douglas-fire pole U.S.A., New York C.J. Wang EU019261 

Cladosporium cladosporioides CBS 109.21; ATCC 11277; ATCC 200940; IFO 6368; IMI 049625 Sooty mould on Hedera helix U.K. — EU019262 

Cladosporium sphaerospermum CBS 188.54; ATCC 11290; IMI 049638 — — — EU019263 

Cladosporium uredinicola ATCC 46649 Hyperparasite on Cronartium 

fusiforme f. sp. quercum 

U.S.A., Alabama — EU019264 

Coccodinium bartschii CBS 121708; CPC 13861–13863 Sooty mould on unidentified tree Canada K.A. Seifert EU019265 

Dissoconium aciculare CBS 342.82*; CPC 1534 Erysiphe, on Medicago lupulina Germany T. Hijwegen EU019266 

Dissoconium commune “Mycosphaerella” communis CBS 114238*; CPC 10440 Eucalyptus globulus Spain J.P.M. Vazquez EU019267 

Dissoconium dekkeri “Mycosphaerella” lateralis CBS 567.89*; CPC 1535 Juniperus chinensis Netherlands T. Hijwegen EU019268 

Fumagospora capnodioides Capnodium salicinum CBS 131.34 Sooty mould on Bursaria spinosa Indonesia — EU019269 

Hortaea werneckii CBS 107.67* Man, tinea nigra Portugal — EU019270 

Nothostrasseria dendritica Teratosphaeria dendritica CPC 12820 Eucalyptus nitens Australia A.J. Carnegie EU019271 

“Passalora” zambiae CBS 112970*; CPC 1228 Eucalyptus globulus Zambia T. Coutinho EU019272 

CBS 112971*; CMW 14782; CPC 1227 Eucalyptus globulus Zambia T. Coutinho EU019273 

Penidiella columbiana CBS 486.80 Paepalanthus columbianus Colombia W. Gams EU019274

Phylogenetic lineages in the Capnodiales 


Penidiella nectandrae CBS 734.87*; ATCC 200932; INIFAT 87/45 Nectandra coriacea Cuba R.F. Castañeda & EU019275 

G. Arnold 

Penidiella rigidophora CBS 314.95* Leaf litter of Smilax sp. Cuba R.F. Castañeda EU019276 

Penidiella strumelloidea CBS 114484*; VKM F-2534 Carex leaf, from stagnant water Russia S. Ozerskaya EU019277 

Penidiella venezuelensis CBS 106.75* Man, tinea nigra Venezuela D. Borelli EU019278 

Phaeotheca triangularis CBS 471.90* Wet surface of humidifier of 

airconditioning 

Belgium H. Beguin EU019279 

Phaeothecoidea eucalypti CPC 13010 Corymbia henryii Australia B. Summerell EU019280 

CPC 12918* Eucalyptus botryoides Australia B. Summerell EU019281 

Pleurophoma sp. Teratosphaeria fibrillosa CPC 1876 Protea nitida South Africa J.E. Taylor EU019282 

Pseudotaeniolina globosa CBS 109889* Rock Italy C. Urzi EU019283 

Ramularia pratensis var. pratensis CPC 11294 Rumex crispus Korea H.D. Shin EU019284 

Ramularia sp. CBS 324.87 On Mycosphaerella sp., leaf spot 

on Brassica sp. 

Netherlands — EU019285 

Readeriella brunneotingens CPC 13303 Eucalyptus tereticornis Australia P.W. Crous EU019286 

Readeriella destructans CBS 111369*; CPC 1366 Eucalyptus grandis Indonesia M.J. Wingfield EU019287 

Readeriella epicoccoides Teratosphaeria suttonii CPC 12352 Eucalyptus sp. U.S.A.,Hawaii W. Gams EU019288 

Readeriella eucalypti CPC 11186 Eucalyptus globulus Spain M.J. Wingfield EU019289 

Readeriella gauchensis CBS 120303*; CMW 17331 Eucalyptus grandis Uruguay M.J. Wingfield EU019290 

Readeriella mirabilis CBS 116293; CPC 10506 Eucalyptus fastigata New Zealand W. Gams EU019291 

Readeriella molleriana Teratosphaeria molleriana CBS 111164*; CMW 4940; CPC 1214 Eucalyptus globulus Portugal M.J. Wingfield EU019292 

Readeriella ovata complex CPC 18 Eucalyptus cladocalyx South Africa P.W. Crous EU019293 

CBS 111149; CPC 23 Eucalyptus cladocalyx South Africa P.W. Crous EU019294 

Readeriella stellenboschiana CBS 116428; CPC 10886 Eucalyptus sp. South Africa P.W. Crous EU019295 

Readeriella zuluensis CBS 120301*; CMW 17321 Eucalyptus grandis South Africa M.J. Wingfield EU019296 

Septoria tritici Mycosphaerella graminicola CBS 100335; IPO 69001.61 Triticum aestivum — G.H.J. Kema EU019297 

CBS 110744; CPC 658 Triticum sp. South Africa P.W. Crous EU019298 

Trimmatostroma betulinum CBS 282.74 Betula verrucosa Netherlands W.M. Loerakker EU019299 

Trimmatostroma salicis CPC 13571 Salix alba Germany U. Braun EU019300 

Teratosphaeria bellula CBS 111700; CPC 1821 Protea eximia South Africa J.E. Taylor EU019301 

Teratosphaeria mexicana CPC 12349 Eucalyptus sp. U.S.A.,Hawaii W. Gams EU019302 

Teratosphaeria nubilosa CBS 114419; CPC 10497 Eucalyptus globulus New Zealand — EU019303 

CBS 116005*; CMW 3282; CPC 937 Eucalyptus globulus Australia A. Carnegie EU019304 

www.studiesinmycology.org

Crous et al. 

Table 1. (Continued). 


Teratosphaeria ohnowa CBS 112896*; CMW 4937; CPC 1004 Eucalyptus grandis South Africa M.J. Wingfield EU019305 

Teratosphaeria secundaria CBS 115608; CPC 504 Eucalyptus grandis Brazil A.C. Alfenas EU019306 

Teratosphaeria sp. CBS 208.94; CPC 727 Eucalyptus grandis Indonesia A.C. Alfenas EU019307 

1 ATCC: American Type Culture Collection, Virginia, U.S.A.; CBS: Centraalbureau voor Schimmelcultures, Utrecht, The Netherlands; CPC: Culture collection of Pedro Crous, housed at CBS; CMW: Culture collection of Mike Wingfield, housed at FABI, 

Pretoria, South Africa; IAM: Institute of Applied Microbiology, University of Tokyo, Institute of molecular and cellular bioscience, Tokyo, Japan; IFO: Institute For Fermentation, Osaka, Japan; IMI: International Mycological Institute, CABI-Bioscience, 

Egham, Bakeham Lane, U.K.; INIFAT: Alexander Humboldt Institute for Basic Research in Tropical Agriculture, Ciudad de La Habana, Cuba; JCM: Japan Collection Of Microorganisms, RIKEN BioResource Center, Japan; VKM: All-Russian Collection of 

Microorganisms, Institute of Biochemistry and Physiology of Microorganisms, Russian Academy of Sciences, Pushchino, Russia. 

*Ex-type cultures. 

analysis were undertaken to characterise Mycosphaerella (Verkley 

et al. 2004), and anamorph genera such as Pseudocercospora 

Speg., Stigmina Sacc., Phaeoisariopsis Ferraris (Crous et al. 

2006a), Ramulispora Miura (Crous et al. 2003), Batcheloromyces 

Marasas, P.S. van Wyk & Knox-Dav. (Taylor et al. 2003), 

Phaeophleospora Rangel and Dothistroma Hulbary (Crous et al. 

2000, 2001, Barnes et al. 2004). 

To assess the phylogeny of the species selected for the present 

study, DNA sequences were generated of the 28S rRNA (LSU) 

gene. In a further attempt to address monophyletic groups within 

this complex, these data were integrated with their morphological 

characteristics. To further resolve pleomorphism among the species 

studied, isolates were examined on a range of cultural media to 

induce possible synanamorphs. 

Materials and methods 

Isolates 

Chosen isolates represent various species previously observed to 

be morphologically distinct from Mycosphaerella s. str. (Crous 1998, 

Crous et al. 2004a, b, 2006a, b, 2007b). In a few cases, specifically 

Teratosphaeria fibrillosa Syd. & P. Syd. and Coccodinium bartschii 

A. Massal., fresh material had to be collected from South Africa 

and Canada, respectively. Excised tissue pieces bearing ascomata 

were soaked in water for approximately 2 h, after which they were 

placed in the bottom of Petri dish lids, with the top half of the 

dish containing 2 % malt extract agar (MEA) (Gams et al. 2007). 

Ascospore germination patterns were examined after 24 h, and 

single-ascospore and conidial cultures established as described by 

Crous (1998). Colonies were sub-cultured onto synthetic nutrientpoor 

agar (SNA), potato-dextrose agar (PDA), oatmeal agar (OA), 

MEA (Gams et al. 2007), and incubated at 25 °C under continuous 

near-ultraviolet light to promote sporulation. 

DNA phylogeny 

Fungal colonies were established on agar plates, and genomic 

DNA was isolated following the CTAB-based protocol described 

in Gams et al. (2007). The primers V9G (de Hoog & Gerrits van 

den Ende 1998) and LR5 (Vilgalys & Hester 1990) were used to 

amplify part of the nuclear rDNA operon spanning the 3’ end of 

the 18S rRNA gene (SSU), the first internal transcribed spacer 

(ITS1), the 5.8S rRNA gene, the second ITS region (ITS2) and the 

5’ end of the 28S rRNA gene (LSU). The primers ITS4 (White et 

al. 1990), LR0R (Rehner & Samuels 1994), LR3R (www.biology. 

duke.edu/fungi/mycolab/primers.htm), and LR16 (Moncalvo et al. 

1993), were used as internal sequence primers to ensure good 

quality sequences over the entire length of the amplicon. The 

ITS1, ITS2 and 5.8S rRNA gene (ITS) were only sequenced for 

isolates of which these data were not available. The ITS data 

were not included in the analyses but deposited in GenBank 

where applicable. The PCR conditions, sequence alignment and 

subsequent phylogenetic analysis using parsimony, distance and 

Bayesian analyses followed the methods of Crous et al. (2006c). 

Gaps longer than 10 bases were coded as single events for the 

phylogenetic analyses; the remaining gaps were treated as new 

character states. Sequence data were deposited in GenBank 

(Table 1) and alignments in TreeBASE (www.treebase.org). 

Taxonomy 

Wherever possible, 30 measurements (× 1 000 magnification)


were made of structures mounted in lactic acid, with the extremes 

of spore measurements given in parentheses. Ascospores were 

frequently also mounted in water to observe mucoid appendages 

and sheaths. Colony colours (surface and reverse) were assessed 

after 1–2 mo on MEA at 25 °C in the dark, using the colour charts of 

Rayner (1970). All cultures obtained in this study are maintained in 

the culture collection of the Centraalbureau voor Schimmelcultures 

(CBS) in Utrecht, the Netherlands (Table 1). Nomenclatural 

novelties and descriptions were deposited in MycoBank (www. 

MycoBank.org). 

Results 

DNA phylogeny 

Amplification products of approximately 1 700 bases were obtained 

for the isolates listed in Table 1. The LSU region of the sequences 

was used to obtain additional sequences from GenBank which 

were added to the alignment. The manually adjusted alignment 

contained 97 sequences (including the two outgroup sequences) 

and 844 characters including alignment gaps. Of the 844 characters 

used in the phylogenetic analysis, 308 were parsimony-informative, 

105 were variable and parsimony-uninformative, and 431 were 

constant. 

The parsimony analysis of the LSU region yielded 1 135 equally 

most parsimonious trees (TL = 1 502 steps; CI = 0.446; RI = 0.787; 

RC = 0.351), one of which is shown in Fig. 1. Three orders are 

represented by the ingroup isolates, namely Chaetothyriales (100 

% bootstrap support), Helotiales (100 % bootstrap support) and 

Capnodiales (100 % bootstrap support). These are discussed in 

detail in the Taxonomy and Discussion sections. A new collection of 

Coccodinium bartschii A. Massal clusters (100 % bootstrap support) 

with members of the Herpotrichiellaceae (Chaetothyriales), whereas 

the type species of the genus Trimmatostroma Corda, namely 

T. salicis Corda, as well as T. betulinum (Corda) S. Hughes, are 

allied (99 % bootstrap support) with the Dermateaceae (Helotiales). 

The Capnodiales encompasses members of the Capnodiaceae, 

Trichosphaeriaceae, Davidiellaceae, Schizothyriaceae and taxa 

traditionally placed in the Mycosphaerellaceae, which is divided 

here into the Teratosphaeriaceae, (65 % bootstrap support), and 

the Mycosphaerellaceae (76 % bootstrap support), which contains 

several subclades. Also included in the Capnodiales are Devriesia 

staurophora (W.B. Kendr.) Seifert & N.L. Nick., Staninwardia 

suttonii Crous & Summerell and Capnobotryella renispora Sugiy. 

as sister taxa to Teratosphaeriaceae s. str. Neighbour-joining 

analysis using three substitution models on the sequence data 

yielded trees supporting the same topologies, but differed from 

the parsimony tree presented with regard to the order of the 

families and orders at the deeper nodes, e.g., the Helotiales and 

Chaetothyriales are swapped around, as are the Capnodiaceae and 

the Trichosphaeriaceae / Davidiellaceae (data not shown). Using 

neighbour-joining analyses, the Mycosphaerellaceae s. str. clade 

obtained 71 %, 70 % and 70 % bootstrap support respectively with 

the uncorrected “p”, Kimura 2-parameter and HKY85 substitution 

models wherease the Teratosphaeriaceae clade obtained 74 %, 79 

% and 78 % bootstrap support respectively with the same models. 

The Schizothyriaceae clade appeared basal in the Capnodiales, 

irrespective of which substitution model was used. 

Bayesian analysis was conducted on the same aligned LSU 

dataset using a general time-reversible (GTR) substitution model 

with inverse gamma rates and dirichlet base frequencies. The 

Markov Chain Monte Carlo (MCMC) analysis of 4 chains started 

from a random tree topology and lasted 23 881 500 generations. 

Trees were saved each 100 generations, resulting in 238 815 saved 

trees. Burn-in was set at 22 000 000 generations after which the 

likelihood values were stationary, leaving 18 815 trees from which 

the consensus tree (Fig. 2) and posterior probabilities (PP’s) were 

calculated. The average standard deviation of split frequencies 

was 0.011508 at the end of the run. The same overall topology as 

that observed using parsimony was obtained, with the exception of 

the inclusion of Staninwardia suttonii in the Mycosphaerellaceae 

(PP value of 0.74) and not in the Teratosphaeriaceae. The 

Mycosphaerellaceae s. str. clade, as well as the Teratosphaeriaceae 

clade, obtained a PP value of 1.00. 

Taxonomy 

Based on the dataset generated in this study, several well-supported 

genera could be distinguished in the Mycosphaerella complex (Figs 

1–2), for which we have identified morphological characters. These 

genera, and a selection of their species, are treated below. 

Key to Mycosphaerella, and Mycosphaerella-like genera treated 

1. Ascomata thyrothecial; anamorph Zygosporium ................................................................................................................. Schizothyrium 

1. Ascomata pseudothecial ............................................................................................................................................................................ 2 

2. Ascospores with irregular, angular lumens typical of Davidiella; anamorph Cladosporium s. str. .............................................. Davidiella 

2. Ascospores guttulate or not, lacking angular lumens; anamorph other than Cladosporium ...................................................................... 3 

3. Ascomata frequently linked by superficial stroma; hamathecial tissue, ascospore sheath, multi-layered endotunica, prominent 

periphysoids, and ascospores turning brown in asci frequently observed ......................................................................... Teratosphaeria 

3. Ascomata not linked by superficial stroma; hamathecial tissue, ascospore sheath, multi-layered endotunica, prominent periphysoids, 

ascospores turning brown in asci not observed ......................................................................................................................................... 4 

4. Conidiophores solitary, pale brown, giving rise to primary and secondary, actively discharged conidia; anamorph Dissoconium 

.................................................................................................................................................................... teleomorph Mycosphaerella-like 

4. Conidiomata variable from solitary conidiophores to sporodochia, fascicles to pycnidia, but conidia not actively discharged ..................... 

.................................................................................................................................................................................. Mycosphaerella s. str. 


Crous et al. 

100 

10 changes 

65 

100 

Athelia epiphylla AY586633 

Paullicorticium ansatum AY586693 

65 

63 

68 

100 

99 

100 

91 

67 

94 

Coccodinium bartschii CPC 13861 

Exophiala oligosperma AF050289 

Exophiala jeanselmei AF050271 

Fonsecaea pedrosoi AF050276 

Capronia mansonii AY004338 

Exophiala dermatitidis DQ823100 

Vibrissea flavovirens AY789426 

Vibrissea truncorum AY789402 

Trimmatostroma salicis CPC 13571 

Mollisia cinerea DQ470942 

Trimmatostroma betulinum CBS 282.74 

Phaeotheca triangularis CBS 471.90 

Capnodium coffeae DQ247800 

Capnodium salicinum CBS 131.34 

Scorias spongiosa DQ678075 

63 

68 

73 

92 

65 

98 

100 

56 

58 

100 87 

100 

100 

76 

Trichosphaeria pilosa AY590297 Trichosphaeriaceae 

Cladosporium cladosporioides CBS 109.21 

Cladosporium uredinicola ATCC 46649 

Cladosporium bruhnei CBS 115683 

Cladosporium sphaerospermum CBS 188.54 

64 

Schizothyrium pomi EF134947 


80 100 

100 

99 

100 

100 

100 

63 

64 

61 

100 

Herpotrichiellaceae, 

Chaetothyriales, 

Chaetothyriomycetes 

Vibrisseaceae 

Dermateaceae 

Capnodiaceae 

Schizothyriaceae 

“Passalora” zambiae CBS 112970 

“Passalora” zambiae CBS 112971 

Dissoconium aciculare CBS 342.82 

“Mycosphaerella” communis CBS 114238 

“Mycosphaerella” lateralis CBS 567.89 

Mycosphaerella graminicola CBS 100335 

Septoria tritici CBS 110744 

Cercosporella centaureicola CBS 120253 

Mycosphaerella punctiformis AY490776 

Ramularia sp. CBS 324.87 

Ramularia miae DQ885902 

Ramularia pratensis var. pratensis CPC 11294 

Capnobotryella renispora CBS 214.90 

100 

94 

88 

80 

99 

81 

Staninwardia suttonii DQ923535 

Devriesia staurophora DQ008150 


Pseudotaeniolina globosa CBS 109889 

Batcheloromyces proteae CBS 110696 

Batcheloromyces leucadendri CBS 110892 

Teratosphaeria alistairii DQ885901 

Penidiella columbiana CBS 486.80 

Teratosphaeria sp. CBS 208.94 

Teratosphaeria ohnowa CBS 112896 

Teratosphaeria secundaria CBS 115608 

Teratosphaeria flexuosa DQ246232 

Hortaea werneckii CBS 107.67 

Catenulostroma castellanii CBS 105.75 

Teratosphaeria bellula CBS 111700 

Catenulostroma elginense CBS 111030 

Teratosphaeria parva DQ246240 

100 Teratosphaeria parva DQ246243 

Catenulostroma germanicum CBS 539.88 

100 

Catenulostroma microsporum CBS 110890 

Catenulostroma abietis CBS 290.90 

Catenulostroma chromoblastomycosum CBS 597.97 

Phaeothecoidea eucalypti CPC 13010 


Teratosphaeria suberosa DQ246235 

Teratosphaeria pseudosuberosa CBS 118911 

93 

Teratosphaeria mexicana CPC 12349 

97 Teratosphaeria mexicana DQ246237 

Teratosphaeria readeriellophora DQ246238 

100 

93 Readeriella eucalypti CPC 11186 

Cibiessia minutispora CPC 13071 

Cibiessia dimorphospora CBS 120034 

Readeriella mirabilis CBS 116293 

Readeriella novaezelandiae DQ246239 

57 Nothostrasseria dendritica CPC 12820 

Batcheloromyces eucalypti CBS 313.76 

99 Readeriella ovata complex CBS 111149 

Readeriella ovata complex CPC 18 

Readeriella destructans CBS 111369 

Readeriella pulcherrima DQ246224 

Readeriella brunneotingens CPC 13303 

Readeriella dimorpha DQ923528 

Teratosphaeria molleriana DQ246219 


Teratosphaeria molleriana CBS 111164 

Teratosphaeria fibrillosa CPC 1876 

Teratosphaeria maxii DQ885899 

Catenulostroma macowanii CBS 110756 

Teratosphaeria nubilosa CBS 114419 


Readeriella stellenboschiana CBS 116428 

Readeriella gauchensis CBS 120303 

Teratosphaeria cryptica DQ246222 

Readeriella considenianae DQ923527 

Cibiessia nontingens CBS 120725 

Readeriella zuluensis CBS 120301 

Readeriella blakelyi DQ923526 

Teratosphaeria toledana DQ246230 

100 

Teratosphaeria suttonii DQ246227 

Readeriella epicoccoides CPC 12352 

Helotiales, Leotiomycetes 

Davidiellaceae 

Mycosphaerellaceae 

Teratosphaeriaceae 

Capnodiales, Dothideomycetes 

Fig. 1. One of 1 135 equally most parsimonious trees obtained from a heuristic search with 100 random taxon additions of the LSU sequence alignment using PAUP v. 4.0b10. 

The scale bar shows 10 changes, and bootstrap support values from 1 000 replicates are shown at the nodes. Thickened lines indicate the strict consensus branches and extype 

sequences are printed in bold face. The tree was rooted to two sequences obtained from GenBank (Athelia epiphylla AY586633 and Paullicorticium ansatum AY586693).


1.00 

0.62 

0.1 expected changes per site 



0.62 

1.00 

0.75 

1.00 

1.00 

0.99 

0.99 

0.85 

0.97 


Exophiala dermatitidis DQ823100 


Coccodinium bartschii CPC 13861 



Vibrisseaceae 

0.97 

Vibrissea flavovirens AY789426 

Vibrissea truncorum AY789402 

1.00 

Trimmatostroma salicis CPC 13571 

Mollisia cinerea DQ470942 

Trimmatostroma betulinum CBS 282.74 

Phaeotheca triangularis CBS 471.90 

Capnodium coffeae DQ247800 

Capnodium salicinum CBS 131.34 

1.00 

1.00 

1.00 

0.80 

0.74 

1.00 

0.55 

1.00 

1.00 

0.91 

Dermateaceae 




Helotiales, 

Leotiomycetes 

Capnodiaceae 

Scorias spongiosa DQ678075 

1.00 

Trichosphaeria pilosa AY590297 Trichosphaeriaceae 

Cladosporium cladosporioides CBS 109.21 

Cladosporium uredinicola ATCC 46649 

1.00 Cladosporium bruhnei CBS 115683 

0.95 Cladosporium sphaerospermum CBS 188.54 


1.00 Schizothyrium pomi EF134947 


Schizothyriaceae 

1.00 

1.00 

0.83 1.00 

0.72 1.00 

1.00 0.58 

0.79 

0.89 0.62 1.00 

0.99 

1.00 

0.58 

0.94 

0.56 

0.83 

0.94 

0.77 

0.51 

1.00 

1.00 

0.77 

0.85 

Dissoconium aciculare CBS 342.82 

“Mycosphaerella” communis CBS 114238 

“Mycosphaerella” lateralis CBS 567.89 

Passalora zambiae CBS 112970 

Passalora zambiae CBS 112971 

Cercosporella centaureicola CBS 120253 


Ramularia sp. CBS 324.87 


Ramularia pratensis var. pratensis CPC 11294 

0.57 Mycosphaerella graminicola CBS 100335 

1.00 Septoria tritici CBS 110744 

Capnobotryella renispora CBS 214.90 

Pseudotaeniolina globosa CBS 109889 

Catenulostroma chromoblastomycosum CBS 597.97 

Teratosphaeria bellula CBS 111700 

Hortaea werneckii CBS 107.67 

Catenulostroma castellanii CBS 105.75 


Batcheloromyces leucadendri CBS 110892 


Penidiella columbiana CBS 486.80 

Teratosphaeria sp. CBS 208.94 

Teratosphaeria ohnowa CBS 112896 

Teratosphaeria secundaria CBS 115608 

Teratosphaeria flexuosa DQ246232 

Catenulostroma elginense CBS 111030 


1.00 

1.00 

1.00 

1.00 

1.00 

1.00 

0.66 

0.76 




Catenulostroma germanicum CBS 539.88 

Catenulostroma microsporum CBS 110890 

Catenulostroma abietis CBS 290.90 



Teratosphaeria pseudosuberosa CBS 118911 


0.97 

Teratosphaeria mexicana CPC 12349 

Teratosphaeria mexicana CBS 110502 


Nothostrasseria dendritica CPC 12820 

Readeriella mirabilis CBS 116293 

Cibiessia dimorphospora CBS 120034 

Cibiessia minutispora CPC 13071 


Readeriella eucalypti CPC 11186 

1.00 

Readeriella brunneotingens CPC 13303 

Readeriella stellenboschiana CBS 116428 

Readeriella gauchensis CBS 120303 


Teratosphaeria cryptica DQ246222 

Cibiessia nontingens CBS 120725 

Readeriella zuluensis CBS 120301 


Teratosphaeria suttonii DQ246227 

Readeriella epicoccoides CPC 12352 

1.00 

Readeriella blakelyi DQ923526 

Readeriella ovata complex CBS 111149 

Readeriella ovata complex CPC 18 

Batcheloromyces eucalypti CBS 313.76 

Readeriella destructans CBS 111369 

Readeriella pulcherrima DQ246224 



Readeriella dimorpha DQ923528 



Teratosphaeria molleriana CBS 111164 

Teratosphaeria fibrillosa CPC 1876 

Catenulostroma macowanii CBS 110756 

Teratosphaeria maxii DQ885899 


Mycosphaerellaceae Teratosphaeriaceae 


Fig. 2. Consensus phylogram (50 % majority rule) of 18 815 trees resulting from a Bayesian analysis of the LSU sequence alignment using MrBayes v. 3.1.2. Bayesian posterior 

probabilities are indicated at the nodes. Ex-type sequences are printed in bold face. The tree was rooted to two sequences obtained from GenBank (Athelia epiphylla AY586633 

and Paullicorticium ansatum AY586693). 


Crous et al. 

Treatment of phylogenetic clades 

Davidiellaceae clade 

Davidiella Crous & U. Braun, Mycol. Progr. 2: 8. 2003. 

Type species: Davidiella tassiana (De Not.) Crous & U. Braun, 

Mycol. Progr. 2: 8. 2003. 

Basionym: Sphaerella tassiana De Not., Sferiacei Italici 1: 87. 

1863. 

Description: Schubert et al. (2007 – this volume). 

Anamorph: Cladosporium Link, Ges. Naturf. Freunde Berlin Mag. 

Neuesten Entdeck. Gesammten Naturk. 7: 37. 1816. 

Type species: Cladosporium herbarum (Pers. : Fr.) Link, Ges. 

Naturf. Freunde Berlin Mag. Neuesten Entdeck. Gesammten 

Naturk. 7: 37. 1816. 

Basionym: Dematium herbarum Pers., Ann. Bot. (Usteri), 11 Stück: 

32. 1794: Fr., Syst. Mycol. 3: 370. 1832. 

Description: Schubert et al. (2007 – this volume). 

Notes: The genus Davidiella (Davidiellaceae) was recently 

introduced for teleomorphs of Cladosporium s. str. (Braun et al. 

2003). The genus Cladosporium is well-established, and contains 

around 772 names (Dugan et al. 2004), while Davidiella presently 

has 33 names (www.MycoBank.org), of which only around five 

have acknowledged Cladosporium states. 

Teratosphaeriaceae clade 

Teratosphaeria Syd. & P. Syd., Ann. Mycol. 10: 39. 1912. 

Type species: Teratosphaeria fibrillosa Syd. & P. Syd., Ann. Mycol. 

10: 40. 1912. Fig. 3. 

Description: Crous et al. (2004a; figs 182–185). 

Notes: Although similar in morphology, the genus Teratosphaeria 

was separated from Mycosphaerella based on its ascomatal 

arrangement, and periphysate ostioles (Müller & Oehrens 1982). 

It was later synonymised under Mycosphaerella by Taylor et 

al. (2003), who showed that the type species clustered within 

Mycosphaerella based on ITS DNA sequence data. The LSU 

sequence data generated in the present study, has clearly shown 

that Mycosphaerella is polyphyletic, thus contradicting earlier 

reports of monophyly by Crous et al. (2000) and Goodwin et al. 

(2001), which were based on ITS data. 

A re-examination of T. fibrillosa, the type species of 

Teratosphaeria, revealed several morphological features that 

characterise the majority of the taxa clustering in the clade, though 

several characters have been lost in some of the small-spored 

species. These characters are discussed below: 

1. Teratosphaeria fibrillosa has a superficial stroma linking 

ascomata together, almost appearing like a spider’s web on the leaf 

surface. Although this feature is not seen in other taxa in this clade, 

some species, such as M. suberosa Crous, F.A. Ferreira, Alfenas 

& M.J. Wingf. and M. pseudosuberosa Crous & M.J. Wingf. have 

a superficial stroma, into which the ascomata are inbedded (Crous 

1998, Crous et al. 2006b). 

2. Ascospores of Teratosphaeria become brown and 

verruculose while still in their asci. This feature is commonly 

observed in species such as M. jonkershoekensis P.S. van Wyk, 

Marasas & Knox-Dav., M. alistairii Crous, M. mexicana Crous, M. 

maxii Crous and M. excentricum Crous & Carnegie (Crous 1998, 

Crous & Groenewald 2006a, b, Crous et al. 2007b). 

3. A few ascomata of T. fibrillosa were found to have 

some pseudoparaphysoidal remnants (cells to distinguish 

pseudoparaphyses), though they mostly disappear with age. This 

feature is rather uncommon, though pseudoparaphyses were 

observed in ascomata of M. eucalypti (Wakef.) Hansf. 

4. Ascospores of Teratosphaeria were found to be covered in 

a mucous sheath, which is commonly observed in other taxa in this 

clade, such as M. bellula Crous & M.J. Wingf., M. pseudocryptica 

Crous, M. suberosa, M. pseudosuberosa, M. associata Crous & 

Carnegie, M. dendritica Crous & Summerell and M. fimbriata Crous 

& Summerell (Crous et al. 2004b, 2006b, 2007b). Re-examination 

of fresh collections also revealed ascospores of M. cryptica (Cooke) 

Hansf. and M. nubilosa (Cooke) Hansf. to have a weakly definable 

sheath. Germinating ascospores of species in this clade all exhibit 

a prominent mucoid sheath. 

5. Asci of T. fibrillosa were observed to have a multi-layered 

endotunica, which, although not common, can be seen in species 

such as M. excentrica, M. maxii, M. alistairii, M. pseudosuberosa, 

M. fimbriata (Crous et al. 2006b, 2007b, Crous & Groenewald 

2006a, b), and also M. nubilosa. 

6. Finally, ascomata of T. fibrillosa and T. proteae-arboreae 

P.S. van Wyk, Marasas & Knox-Dav. have well-developed 

ostiolar periphyses, which are also present in species such as M. 

suberosa, M. pseudosuberosa, M. maxii and T. microspora Joanne 

E. Taylor & Crous (Crous 1998, Crous et al. 2004a, b, 2006b). 

Morphologically thus, the Teratosphaeria clade is distinguishable 

from Mycosphaerella s. str., though these differences are less 

pronounced in some of the smaller-spored species. Based on 

these distinct morphological features, as well as its phylogenetic 

position within the Capnodiales, a new family is herewith proposed 

to accommodate species of Teratosphaeria: 

Teratosphaeriaceae Crous & U. Braun, fam. nov. MycoBank 

MB504464. 

Ascomata pseudotheciales, superficiales vel immersa, saepe in stromate ex cellulis 

brunneis pseudoparenchymatibus disposita, globulares, uniloculares, papillata, 

apice ostiolato, periphysata, saepe cum periphysoidibus; tunica multistratosa, ex 

cellulis brunneis angularibus composita, strato interiore ex cellulis applanatis hyalinis; 

saepe cum pseudoparaphysibus subcylindricis, ramosis, septatis, anastomosibus. 

Asci fasciculati, octospori, bitunicati, saepe cum endotunica multistratosa. 

Ascosporae ellipsoideae-fusiformes vel obovoideae, 1-septatae, hyalinae, deinde 

pallide brunneae et verruculosae, saepe mucosae. 

Ascomata pseudothecial, superficial to immersed, frequently 

situated in a stroma of brown pseudoparenchymatal cells, 

globose, unilocular, papillate, ostiolate, canal periphysate, with 

periphysoids frequently present; wall consisting of several layers 

of brown textura angularis; inner layer of flattened, hyaline cells. 

Pseudoparaphyses frequently present, subcylindrical, branched, 

septate, anastomosing. Asci fasciculate, 8-spored, bitunicate, 

frequently with multi-layered endotunica. Ascospores ellipsoidfusoid 

to obovoid, 1-septate, hyaline, but becoming pale brown and 

verruculose, frequently covered in mucoid sheath. 

Typus: Teratosphaeria Syd. & P. Syd., Ann. Mycol. 10: 39. 1912. 

Teratosphaeria africana (Crous & M.J. Wingf.) Crous & U. Braun, 

comb. nov. MycoBank MB504466.


Fig. 3. Teratosphaeria fibrillosa (epitype material). A. Leaf spots. B. Subepidermal ascomata linked by means of stromatic tissue. C. Paraphyses among asci. D. Periphysoids. E. 

Ascospores becoming brown in asci. F–G. Multi-layered endotunica. H–K. Ascospores, becoming brown and verruculose. L–M. Germinating ascospores. Scale bars = 10 µm. 

Basionym: Mycosphaerella africana Crous & M.J. Wingf., Mycologia 

88: 450. 1996. 

Teratosphaeria associata (Crous & Carnegie) Crous & U. Braun, 

comb. nov. MycoBank MB504467. 

Basionym: Mycosphaerella associata Crous & Carnegie, Fungal 

Diversity 26: 159. 2007. 

Teratosphaeria alistairii (Crous) Crous & U. Braun, comb. nov. 

MycoBank MB504468. 

Basionym: Mycosphaerella alistairii Crous, in Crous & Groenewald, 

Fungal Planet, No. 4. 2006. 

Anamorph: Batcheloromyces sp. 


Crous et al. 

Teratosphaeria bellula (Crous & M.J. Wingf.) Crous & U. Braun, 


Basionym: Mycosphaerella bellula Crous & M.J. Wingf., Mycotaxon 

46: 20. 1993. 

Teratosphaeria cryptica (Cooke) Crous & U. Braun, comb. nov. 


Basionym: Sphaerella cryptica Cooke, Grevillea 20: 5. 1891. 

≡ Mycosphaerella cryptica (Cooke) Hansf., Proc. Linn. Soc. New South 

Wales 81: 35. 1956. 

Anamorph: Readeriella nubilosa (Ganap. & Corbin) Crous & U. 

Braun, comb. nov. MycoBank MB504471. 

Basionym: Colletogloeum nubilosum Ganap. & Corbin, Trans. Brit. 

Mycol. Soc. 72: 237. 1979. 

≡ Colletogloeopsis nubilosum (Ganap. & Corbin) Crous & M.J. Wingf., 

Canad. J. Bot. 75: 668. 1997. 

Teratosphaeria dendritica (Crous & Summerell) Crous & U. 


Basionym: Mycosphaerella dendritica Crous & Summerell, Fungal 

Diversity 26: 161. 2007. 

Anamorph: Nothostrasseria dendritica (Hansf.) Nag Raj, Canad. 

J. Bot. 61: 25. 1983. 

Basionym: Spilomyces dendriticus Hansf., Proc. Linn. Soc. New 

South Wales 81: 32. 1956. 

Teratosphaeria excentrica (Crous & Carnegie) Crous & U. Braun, 


Basionym: Mycosphaerella excentrica Crous & Carnegie, Fungal 

Diversity 26: 164. 2007. 

Anamorph: Catenulostroma excentricum (B. Sutton & Ganap.) 

Crous & U. Braun, comb. nov. MycoBank MB504475. 

Basionym: Trimmatostroma excentricum B. Sutton & Ganap., New 

Zealand J. Bot. 16: 529. 1978. 

Teratosphaeria fibrillosa Syd. & P. Syd., Ann. Mycol. 10: 40. 

1912. 

≡ Mycosphaerella fibrillosa (Syd. & P. Syd.) Joanne E. Taylor & Crous, 

Mycol. Res. 107: 657. 2003. 

Specimens examined: South Africa, Western Cape Province, Bains Kloof near 

Wellington, on living leaves of Protea grandiflora, 26 Feb. 1911, E.M. Doidge, 

holotype PREM; Stellenbosch, Jonkershoek valley, S33° 59’ 44.7”, E18° 58’ 50.6”, 

1 Apr. 2007, on leaves of Protea sp., P.W. Crous & L. Mostert, epitype designated 

here CBS H-19913, culture ex-epitype CBS 121707 = CPC 13960. 

Teratosphaeria fimbriata (Crous & Summerell) Crous & U. Braun, 


Basionym: Mycosphaerella fimbriata Crous & Summerell, Fungal 

Diversity 26: 166. 2007. 

Teratosphaeria flexuosa (Crous & M.J. Wingf.) Crous & U. Braun, 


Basionym: Mycosphaerella flexuosa Crous & M.J. Wingf., Mycol. 

Mem. 21: 58. 1998. 

Teratosphaeria gamsii (Crous) Crous & U. Braun, comb. nov. 


Basionym: Mycosphaerella gamsii Crous, Stud. Mycol. 55: 113. 

2006. 

Teratosphaeria jonkershoekensis (P.S. van Wyk, Marasas & 

Knox-Dav.) Crous & U. Braun, comb. nov. MycoBank MB504479. 

Basionym: Mycosphaerella jonkershoekensis P.S. van Wyk, 

Marasas & Knox-Dav., J. S. African Bot. 41: 234. 1975. 

Teratosphaeria maxii (Crous) Crous & U. Braun, comb. nov. 


Basionym: Mycosphaerella maxii Crous, in Crous & Groenewald, 

Fungal Planet No. 6. 2006. 

Teratosphaeria mexicana (Crous) Crous & U. Braun, comb. nov. 


Basionym: Mycosphaerella mexicana Crous, Mycol. Mem. 21: 81. 

1998. 

Teratosphaeria microspora Joanne E. Taylor & Crous, Mycol. 

Res. 104: 631. 2000. 

≡ Mycosphaerella microspora (Joanne E. Taylor & Crous) Joanne E. 

Taylor & Crous, Mycol. Res. 107: 657. 2003. 

Anamorph: Catenulostroma microsporum (Joanne E. Taylor & 

Crous) Crous & U. Braun, comb. nov. MycoBank MB504482. 

Basionym: Trimmatostroma microsporum Joanne E. Taylor & 

Crous, Mycol. Res. 104: 631. 2000. 

Teratosphaeria molleriana (Thüm.) Crous & U. Braun, comb. 

nov. MycoBank MB504483. 

Basionym: Sphaerella molleriana Thüm., Revista Inst. Sci. Lit. 

Coimbra 28: 31. 1881. 

≡ Mycosphaerella molleriana (Thüm) Lindau, Nat. Pfanzenfam. 1: 424. 

1897. 

= Mycosphaerella vespa Carnegie & Keane, Mycol. Res. 102: 1275. 1998. 

= Mycosphaerella ambiphylla A. Maxwell, Mycol. Res. 107: 354. 2003. 

Anamorph: Readeriella molleriana (Crous & M.J. Wingf.) Crous & 

U. Braun, comb. nov. MycoBank MB504484. 

Basionym: Colletogloeopsis molleriana Crous & M.J. Wingf., 

Canad. J. Bot. 75: 670. 1997. 

Teratosphaeria nubilosa (Cooke) Crous & U. Braun, comb. nov. 


Basionym: Sphaerella nubilosa Cooke, Grevillea 19: 61. 1892. 

≡ Mycosphaerella nubilosa (Cooke) Hansf., Proc. Linn. Soc. New South 

Wales 81: 36. 1965. 

= Mycosphaerella juvenis Crous & M.J. Wingf., Mycologia 88: 453. 1996. 

Teratosphaeria ohnowa (Crous & M.J. Wingf.) Crous & U. Braun, 


Basionym: Mycosphaerella ohnowa Crous & M.J. Wingf., Stud. 

Mycol. 50: 206. 2004. 

Teratosphaeria parkiiaffinis (Crous & M.J. Wingf.) Crous & U. 


Basionym: Mycosphaerella parkiiaffinis Crous & M.J. Wingf., Fungal 

Diversity 26: 168. 2007. 

Teratosphaeria parva (R.F. Park & Keane) Crous & U. Braun, 


Basionym: Mycosphaerella parva R.F. Park & Keane, Trans. Brit. 

Mycol. Soc. 79: 99. 1982. 

= Mycosphaerella grandis Carnegie & Keane, Mycol. Res. 98: 414. 1994. 

Teratosphaeria perpendicularis (Crous & M.J. Wingf.) Crous & U. 


Basionym: Mycosphaerella perpendicularis Crous & M.J. Wingf., 

Stud. Mycol. 55: 113. 2006. 

Teratosphaeria pluritubularis (Crous & Mansilla) Crous & U. 


Basionym: Mycosphaerella pluritubularis Crous & Mansilla, Stud. 

Mycol. 55: 114. 2006. 

10


Teratosphaeria pseudafricana (Crous & T.A. Cout.) Crous & U. 


Basionym: Mycosphaerella pseudafricana Crous & T.A. Cout., 

Stud. Mycol. 55: 115. 2006. 

Teratosphaeria pseudocryptica (Crous) Crous & U. Braun, comb. 


Basionym: Mycosphaerella pseudocryptica Crous, Stud. Mycol. 55: 

6. 2006. 

Anamorph: Readeriella sp. 

Teratosphaeria pseudosuberosa (Crous & M.J. Wingf.) Crous & 


Basionym: Mycosphaerella pseudosuberosa Crous & M.J. Wingf., 

Stud. Mycol. 55: 118. 2006. 

Anamorph: Catenulostroma sp. 

Teratosphaeria quasicercospora (Crous & T.A. Cout.) Crous & U. 


Basionym: Mycosphaerella quasicercospora Crous & T.A. Cout., 

Stud. Mycol. 55: 119. 2006. 

Teratosphaeria readeriellophora (Crous & Mansilla) Crous & U. 


Basionym: Mycosphaerella readeriellophora Crous & Mansilla, 

Stud. Mycol. 50: 207. 2004. 

Anamorph: Readeriella readeriellophora Crous & Mansilla, Stud. 

Mycol. 50: 207. 2004. Fig. 18. 

Teratosphaeria secundaria (Crous & Alfenas) Crous & U. Braun, 


Basionym: Mycosphaerella secundaria Crous & Alfenas, Stud. 

Mycol. 55: 122. 2006. 

Teratosphaeria stramenticola (Crous & Alfenas) Crous & U. 


Basionym: Mycosphaerella stramenticola Crous & Alfenas, Stud. 

Mycol. 55: 123. 2006. 

Teratosphaeria suberosa (Crous, F.A. Ferreira, Alfenas & M.J. 

Wingf.) Crous & U. Braun, comb. nov. MycoBank MB504498. 

Basionym: Mycosphaerella suberosa Crous, F.A. Ferreira, Alfenas 

& M.J. Wingf., Mycologia 85: 707. 1993. 

Teratosphaeria suttonii (Crous & M.J. Wingf.) Crous & U. Braun, 


Basionym: Mycosphaerella suttonii Crous & M.J. Wingf. (suttoniae), 

Canad. J. Bot. 75: 783. 1997. 

Anamorph: Readeriella epicoccoides (Cooke & Massee) Crous & 


Basionym: Cercospora epicoccoides Cooke & Massee apud Cooke, 

Grevillea 19: 91. 1891. 

≡ Phaeophleospora epicoccoides (Cooke & Massee) Crous, F.A. Ferreira 

& B. Sutton, S. African J. Bot. 63: 113. 1997. 

≡ Kirramyces epicoccoides (Cooke & Massee) J. Walker, B. Sutton & 

Pascoe, Mycol. Res. 96: 919. 1992. 

= Hendersonia grandispora McAlp., Proc. Linn. Soc. New South Wales 28: 

99. 1903. 

= Phaeoseptoria eucalypti Hansf., Proc. Linn. Soc. New South Wales 82: 225. 

1957. 

= Phaeoseptoria luzonensis T. Kobayashi, Trans. Mycol. Soc. Japan 19: 377. 

1978. 

Synanamorph: Pseudocercospora sp. 

Teratosphaeria toledana (Crous & Bills) Crous & U. Braun, comb. 


Basionym: Mycosphaerella toledana Crous & Bills, Stud. Mycol. 50: 

208. 2004. 

Anamorph: Readeriella toledana (Crous & Bills) Crous & U. Braun, 


Basionym: Phaeophleospora toledana Crous & Bills, Stud. Mycol. 

50: 208. 2004. 

Key to treated anamorph genera of Teratosphaeria (Teratosphaeriaceae) 

1. Hyphae submerged to superficial, disarticulating into arthroconidia .......................................................................................................... 2 

1. Hyphae not disarticulating into arthroconidia ............................................................................................................................................. 3 

2. Mature, brown hyphae disarticulating into thick-walled, spherical, smooth to verruculose 0(–2) transversely septate, 

brown conidia ............................................................................................................................ Pseudotaeniolina (= Friedmanniomyces) 

2. Hyphae superficial, brown to green-brown, smooth, disarticulating to form pale brown, cylindrical, 0–3-septate conidia with subtruncate 

ends, frequently with a Readeriella synanamorph ....................................................................................................................... Cibiessia 

3. Hyphal ends forming endoconidia; hyphae pale to medium brown, verruculose, end cells dividing into several brown, verruculose, 

thick-walled, ellipsoid to obovoid endoconidia ................................................................................................................. Phaeothecoidea 

3. Endoconidia absent.................................................................................................................................................................................... 4 

4. Conidiogenous cells integrated in hyphae; well-developed conidiomata or long, solitary, macronematous, terminally penicillate 

conidiophores absent ................................................................................................................................................................................. 5 

4. Conidiomata well-developed or with long, solitary, terminally penicillate conidiophores ........................................................................... 7 

5. Conidia in chains, holoblastic, pseudocladosporium-like in morphology, but scars and hila not excessively thickened, nor refractive, 

producing chlamydospores in culture; species are mostly heat resistant .................................................................................... Devriesia 

5. Conidia solitary on indistict to well defined phialides on hyphae ............................................................................................................... 6 

6. Conidiogenous cells integrated in the distal ends of hyphae; conidia thick-walled, brown, smooth, 

-septate ......................................................................................................................................................................... Capnobotryella 

www.studiesinmycology.org 

11

Crous et al. 

6. Conidiophores short and frequently reduced to conidiogenous cells that proliferate percurrently via wide necks, giving rise to hyaline, 0(–2)- 

septate, broadly ellipsoidal conidia ................................................................................................................................................ Hortaea 

7. Conidia brown, with hyaline basal appendages; conidiomata pycnidial, conidiogenous cells phialidic, but also percurrent, 

subhyaline ....................................................................................................................................................................... Nothostrasseria 

7. Conidia brown, but basal appendages lacking, amero- to scolecospores ................................................................................................. 8 

8. Conidiomata pycnidial to acervular ............................................................................................................................................................ 9 

8. Conidiomata not enclosed by host tissue, fasciculate to sporodochial or solitary, hyphomycetous ........................................................ 10 

9. Conidia solitary, dry, without mucilaginous sheath .................................................................................................................... Readeriella 

9. Conidia catenulate, with persistant mucilaginous sheath ..................................................................................................... Staninwardia 

10. Conidiophores usually solitary, rarely densely fasciculate to synnematous (in vivo), penicillate, with a branched, apical conidiogenous 

apparatus giving rise to ramoconidia and branched chains of secondary conidia; scars not to slightly thickened and darkened-refractive 

..................................................................................................................................................................................................... Penidiella 

10. Conidiophores not penicillate, without a branched conidiogenous apparatus, in vivo fasciculate to sporodochial .................................. 11 

11. Biotrophic; fruiting composed of sporodochia and radiating layers of hyphae arising from the stromata, conidiophores arising from 

superficial sporodochia and radiating hyphae, conidiogenous cells unilocal, with conspicuous annellations, conidia solitary or in fragile 

disarticulating chains, aseptate or transversely 1–3-septate, usually with distinct frills, secession rhexolytic ............... Batcheloromyces 

. Biotrophic, leaf-inhabiting, with distinct, subepidermal to erumpent, well-developed sporodochia, or saxicolous, saprobic, sometimes 

causing opportunistic human infections; radiating layers of hyphae arising from sporodochia; conidiogenous cells without 

annellations; conidia in true simple or branched basipetal chains, transversely 1- to pluriseptate or with longitudinal and oblique septa 

(dictyosporous), occasionally distoseptate ...................................................................................................................... Catenulostroma 

To explain the arguments behind the selection and synonymies 

of some of these anamorphic genera, they are briefly discussed 

below: 

Acidomyces Baker et al., Appl. Environ. Microbiol. 70: 6270. 2004. 

(nom. inval.) 

Type species: Acidomyces richmondensis Baker et al., Appl. 

Environ. Microbiol. 70: 6270. 2004. (nom. inval.) 

Notes: The genus presently clusters among isolates in the 

Teratosphaeria clade based on sequences deposited in GenBank. 

Acidomyces lacks a Latin description and holotype specimen, and 

is thus invalidly described. The genus, which was distinguished 

from other taxa based on its DNA phylogeny (Dothideomycetes), 

forms filamentous hyphae with disarticulating cells. It is unclear 

how it differs from Friedmanniomyces Onofri and Pseudotaeniolina 

J.L. Crane & Schokn. 

Batcheloromyces Marasas, P.S. van Wyk & Knox-Dav., J. S. 

African Bot. 41: 41. 1975. 

Type species: Batcheloromyces proteae Marasas, P.S. van Wyk & 

Knox-Dav., J. S. African Bot. 41: 43. 1975. 

Description: Crous et al. (2004a; figs 4–26). 

Notes: Batcheloromyces is presently circumscribed as a genus 

that forms emergent hyphae, giving rise to superficial sporodochial 

plates, forming brown, verrucose, erect conidiophores that 

proliferate holoblastically, with ragged percurrent proliferations that 

become visible with age. Conidia are produced singly or in fragile, 

disarticulating chains, are brown, thick-walled, 0–3 transversely 

euseptate (though at times they appear as distoseptate). The genus 

Batcheloromyces has in recent years been confused with Stigmina 

(Sutton & Pascoe 1989) on the basis that some collections showed 

conidiophores to give rise to solitary conidia only, though conidial 

catenulation was clearly illustrated by Taylor et al. (1999). In culture 

colonies tend to sporulate in a slimy mass (on OA), though a 

synanamorph can be seen (in B. leucadendri, Fig. 4) to sporulate 

via holoblastic conidiogenesis on hyphal tips of the aerial mycelium, 

forming elongate-globose to ellipsoid, muriformly septate, thickwalled 

conidia, that occur in clusters. 

The finding that Stigmina s. str. [based on S. platani (Fuckel) 

Sacc., the type species] is a generic synonym of Pseudocercospora 

Speg. (Crous et al. 2006a), and that the type species of 

Trimmatostroma (T. salicis, Fig. 5) belongs to the Helotiales 

(Fig. 1), raises the question of where to place stigmina- and 

trimmatostroma-like anamorphs that reside in the Teratosphaeria 

clade. Although the stigmina-like species can be accommodated 

in Batcheloromyces (see Sutton & Pascoe 1989), a new genus is 

required for Teratosphaeria anamorphs that have a trimmatostromalike 

morphology. The recognition of Batcheloromyces and the 

introduction of a new anamorph genus for trimmatostroma-like 

anamorphs of Teratosphaeria are also morphologically justified. 

Batcheloromyces is easily distinguishable from Stigmina s. 

str. by its special structure of the fruiting body, composed of 

sporodochia and radiating layers of hyphae arising from the 

sporodochia and the conidia often formed in delicate disarticulating 

chains. Trimmatostroma-like anamorphs of Teratosphaeria are 

morphologically also sufficently distinct from Trimmatostroma s. str. 

(see notes under Catenulostroma Crous & U. Braun) as well as 

Batcheloromyces (see key above). 

Batcheloromyces eucalypti (Alcorn) Crous & U. Braun, comb. 


Basionym: Stigmina eucalypti Alcorn, Trans. Brit. Mycol. Soc. 60: 

151. 1973. 

Capnobotryella Sugiy., in Sugiyama, Pleomorphic Fungi: The 

Diversity and its Taxonomic Implications (Tokyo): 148. 1987. 

Type species: Capnobotryella renispora Sugiy., in Sugiyama, 

Pleomorphic Fungi: The Diversity and its Taxonomic Implications 

(Tokyo): 148. 1987. 

Description: Sugiyama & Amano (1987, figs 7.5–7.8). 

12


Fig. 4. Batcheloromyces leucadendri in vitro. A–B. Batcheloromyces state with synanamorph (arrows). C–D. Conidia occurring solitary or in short chains. Scale bar = 10 µm. 

Fig. 5. Trimmatostroma salicis. A. Sporodochia on twig. B–E. Chains of disarticulating conidia. Scale bars = 10 µm. 

Notes: The genus forms brown, septate, thick-walled hyphae, with 

ellipsoidal, 0–1-septate conidia forming directly on the hyphae, via 

minute phialides. Hambleton et al. (2003) also noted the occurrence 

of endoconidiation. 

Catenulostroma Crous & U. Braun, gen. nov. MycoBank 

MB504474. 

Etymology: Named after its catenulate conidia, and stromata giving 

rise to sporodochia. 

Hyphomycetes. Differt a Trimmatostromate habitu phytoparasitico, maculis 

formantibus, conidiophoris saepe fasciculatis, per stoma emergentibus vel habitu 

saxiphilo-saprophytico, interdum sejunctis ex mycosibus humanis. 


13

Crous et al. 

Habit plant pathogenic, leaf-spotting or saxicolous-saprobic, 

occasionally isolated from opportunistic human mycoses. 

Mycelium internal and external; hyphae dark brown, septate, 

branched. Conidiomata in vivo vary from acervuli to sporodochia or 

fascicles of conidiophores arising from well-developed or reduced, 

pseudoparenchymatal stromata. Setae and hyphopodia absent. 

Conidiophores arising from hyphae or stromata, solitary, fasciculate 

to sporodochial, in biotrophic, plant pathogenic species emerging 

through stomata, little differentiated, semimacronematous, branched 

or not, continuous to septate, brown, smooth to verruculose. 

Conidiogenous cells integrated, terminal or conidiophores reduced 

to conidiogenous cells, holoblastic-thalloblastic, meristematic, 

unilocal, delimitation of conidium by a single septum with 

retrogressive delimitation of next conidium giving an unconnected 

chain of conidia, brown, smooth to verruculose, conidiogenous 

scars (conidiogenous loci) inconspicuous, truncate, neither 

thickened nor darkened. Conidia solitary or usually forming simple 

to branched basipetal chains of transversely to muriformly eu- or 

distoseptate, 1– to multiseptate, brown, smooth, verruculose to 

verrucose conidia, conidial secession schizolytic. 

Type species: Catenulostroma protearum (Crous & M.E. Palm) 

Crous & U. Braun, comb. nov. 

Description: Crous & Palm (1999), Crous et al. (2004a; figs 364– 

365). 

Notes: Catenulostroma contains several plant pathogenic species 

previously placed in Trimmatostroma, a morphologically similar but, 

based on its type species, phylogenetically distinct genus belonging 

to Helotiales (Fig. 1). Trimmatostroma s. str. is well-distinguished 

from most Catenulostroma species by being saprobic, living on 

twigs and branches of woody plants, or occasionally isolated 

from leaf litter, i.e., they are not associated with leaf spots. The 

conidiomata of Trimmatostroma species are subepidermal, 

acervular-sporodochial with a well defined wall of textura angularis, 

little differentiated, micronematous conidiophores giving rise to long 

chains of conidia that disarticulate at the surface to form a greyblack 

to brown powdery mass. The generic affinity of other species 

assigned to Trimmatostroma, e.g. those having a lichenicolous 

habit, is unresolved. 

Trimmatostroma abietis Butin & Pehl (Butin et al. 1996) clusters 

together with the plant pathogenic Catenulostroma species, but 

differs from these species in having a more complex ecology. 

Trimmatostroma abietis is usually foliicolous on living or necrotic 

conifer needles on which characteristic acervuli to sporodochia 

with densely arranged, fasciculate fertile hyphae are formed, 

comparable to the fasciculate conidiomata of the plant pathogenic 

species of Catenulostroma (Butin et al. 1996: 205, fig. 1). Although 

not discussed by Butin et al. (1996), T. abietis needs to be compared 

to T. abietina Doherty, which was orginally described from Abies 

balsamea needles collected in Guelph, Canada (Doherty 1900). 

Morphologically the two species appear to be synonymous, except 

for reference to muriformly septate conidia, which is a feature not 

seen in vivo in the type of T. abietis. Furthermore, as this is clearly 

a species complex, this matter can only be resolved once fresh 

Canadian material has been collected to serve as epitype for T. 

abietina. 

Isolates from stone, agreeing with T. abietis in cultural, 

morphological and physiological characteristics, have frequently 

been found (Wollenzien et al. 1995, Butin et al. 1996, Gorbushina 

et al. 1996, Kogej et al. 2006, Krumbein et al. 1996). Furthermore, 

isolates from humans (ex skin lesions and ex chronic osteomycelitis 

of human patients) and Ilex leaves are known (Butin et al. 1996). De 

Hoog et al. (1999) included strains of T. abietis from stone, man and 

Ilex leaves in molecular sequence analyses and demonstrated their 

genetical identity based on 5.8S rDNA and ITS2 data, but strains 

from conifer needles were not included. Furthermore, we consider 

T. abietis, as presently defined, to represent a species complex, 

with Dutch isolates from Pinus again appearing distinct from 

German Abies isolates, suggesting that different conifer genera 

could harbour different Catenulostroma species. Isolates from 

stone form stromatic, durable microcolonies, which are able to grow 

under extreme xerophilic environmental conditions. Cultural growth 

resembles that of other meristematic black yeasts (Butin et al. 1996, 

Kogej et al. 2006). Another fungus isolated from stone in Germany 

is in vitro morphologically close to C. abietis, but differs in forming 

conidia with oblique septa. Furthermore, a human pathogenic 

isolate from Africa clusters together with other Catenulostroma 

species. The habit and origin of this human pathogenic fungus in 

nature and its potential morphology on “natural” substrates, which 

typically deviates strongly from the growth in vitro, are still unknown. 

However, C. abietis, usually growing as a foliicolous and saxicolous 

fungus, has already shown the potential ability of Catenulostroma 

species to cause opportunistic human infections. 

Key to Catenulostroma species 

1. Conidia formed in basipetal chains, smooth, 4-celled, consisting of two basal cells with truncate lateral sides, each giving rise to a 

secondary globose apical cell, that can extend and develop additional septa, appearing as two lateral arms ................. C. excentricum 

1. Conidia variable in shape, but without two basal cells giving rise to two lateral arms ............................................................................... 2 

2. Conidia smooth or almost so, at most very faintly rough-walled; usually foliicolous on conifer needles or saxicolous, forming stromatic, 

xerophilic durable microcolonies on stone, occasionally causing opportunistic human infections ............................................................ 3 

2. Conidia distinctly verruculose to verrucose; plant pathogenic, forming leaf spots ..................................................................................... 5 

3. Conidia (8–)20–35(–60) × 4–5(–7) µm, 1–10-septate .................................................................................... C. chromoblastomycosum 

3. Conidia much shorter, 8–20 µm long, 0–5-septate .................................................................................................................................... 4 

4. Conidia 0–5 times transversely septate, mostly two-celled; usually foliicolous on conifer needles or saxicolous ....................... C. abietis 

4. Conidia 2–4 times transversely septate and often with 1–2 oblique septa; isolated from stone ........................................ C. germanicum 

5. Conidia rather broad, usually wider than 10 µm ........................................................................................................................................ 6 

14


Fig. 6. Catenulostroma chromoblastomycosum (type material). A. Sporodochium on pine needle in vitro. B–H. Chains of disarticulating conidia. Scale bars: A = 350, B, E, G, 

H = 10 µm. 

5. Conidia narrower, width below 10 µm ....................................................................................................................................................... 7 

6. Conidia distoseptate, rather long, (12–)25–35(–45) × (7–)10–15(–25) µm; conidiomata large, up to 250 µm diam, on Protea anceps 

............................................................................................................................................................................................... C. protearum 

6. Conidia euseptate, shorter, (9–)16–20(–36) × (10–)14–18(–27) µm; sporodochia 90–100 × 40–80 µm; on Protea grandiceps 

................................................................................................................................................................................................. C. elginense 

7. Conidia 1- to multiseptate, (10–)15–17(–23) × (5–)6.5–7(–9) µm; on various Proteaceae .................................................. C. macowanii 

7. Conidia in vivo predominantly 1-septate, (8–)13–15(–21) × (3.5–)5.5–6(–8) µm; on Protea cynaroides 

......................................................................................................................................... C. microsporum (Teratosphaeria microspora) 

Catenulostroma abietis (Butin & Pehl) Crous & U. Braun, comb. 


Basionym: Trimmatostroma abietis Butin & Pehl, Antonie van 

Leeuwenhoek 69: 204. 1996. 

Notes: Catenulostroma abietis needs to be compared to 

Trimmatostroma abietina Doherty (Abies balsamea needles 

Canada), which is either an older name for this species, or a closely 

related taxon. Presently T. abietina is not known from culture, and 

needs to be recollected. 

Catenulostroma chromoblastomycosum Crous & U. Braun, sp. 

nov. MycoBank MB504505. Fig. 6. 

Etymology: Named after the disease symptoms observed due to 

opportunistic human infection. 

Differt a C. abieti et C. germanico conidiis longioribus, (8–)20–35(–60) × 4–5(–7) 

µm, 1–10-septatis. 

Description based on cultures sporulating on WA supplemented 

with sterile pine needles. Mycelium consisting of branched, 

septate, smooth to finely verruculose, medium to dark brown, 

thick-walled, 3–4 µm wide hyphae. Conidiomata brown, superficial, 


15

Crous et al. 

Fig. 7. Catenulostroma germanicum (type material). A–D. Chains of disarticulating conidia in vitro. Scale bars = 10 µm. 

sporodochial, up to 350 µm diam. Conidiophores reduced to 

inconspicuous conidiogenous loci on hyphae, 2–4 µm wide, neither 

darkened nor thickened or refractive. Conidia occurring in branched 

chains, that tend to remain attached to each other, subcylindrical 

with subtruncate ends, straight to slightly curved, (8–)20–35(–60) 

× 4–5(–7) µm, 1–10-septate, medium brown, smooth to finely 

verruculose. 

Cultural characteristics: Colonies on PDA erumpent, spreading, 

slow growing, with sparse to moderate aerial mycelium and smooth, 

irregular, submerged margins; greenish black (surface). 

Specimen examined: Zaire, Pawa, isolated from man with chromoblastomycosis, 

Mar. 1997, V. de Brouwere, holotype CBS H-19935, culture ex-type CBS 597.97. 

Notes: Catenulostroma chromoblastomycosum was originally 

identified as an isolate of Stenella araguata Syd. The latter fungus 

is morphologically distinct, however, having much shorter and 

narrower conidia, formed in acropetal chains, as well as quite 

different conidiogenous loci and conidial hila which are small, 

thickened and darkened. 

Catenulostroma elginense (Joanne E. Taylor & Crous) Crous & 


Basionym: Trimmatostroma elginense Joanne E. Taylor & Crous, 

Mycol. Res. 104: 633. 2000. 

Catenulostroma excentricum, see Teratosphaeria excentrica. 

Catenulostroma germanicum Crous & U. Braun, sp. nov. 

MycoBank MB504507. Fig. 7. 

Etymology: Named after the geographic location of its type strain 

in Germany. 

Differt a C. abieti conidiis 1–2 oblique septatis. 

Mycelium consisting of branched, septate, smooth, pale to 

medium brown, 2–4 µm wide hyphae, giving rise to conidial 

chains. Conidiophores integrated, subcylindrical, branched or not, 

septate, little differentiated, micronematous, 3–5 µm wide, 3- to 

multiseptate, medium brown, thick-walled; conidiogenous cells 

integrated, terminal, inconspicuous, unilocal, conidiogenous loci 

16


inconspicuous. Conidia in simple or branched basipetal chains, 

subcylindrical, straight to flexuous, (8–)10–15(–20) × 4–5(–6) µm, 

2–4 transversely septate or with 1–2 oblique septa, medium to dark 

brown, thick-walled, smooth. 

Cultural characteristics: Colonies on OA erumpent, spreading, with 

even, smooth margins and sparse to moderate aerial mycelium; 

olivaceous-grey, with iron-grey margins (surface). Colonies reaching 

12 mm diam after 1 mo at 25 °C in the dark; colonies fertile. 

Specimen examined: Germany (former West-Germany), isolated from stone, Oct. 

1988, J. Kuroczkin, holotype CBS H-19936, culture ex-type CBS 539.88. 

Notes: Catenulostroma germanicum was originally deposited as 

Taeniolina scripta (P. Karst.) P.M. Kirk. It is clearly distinct, however, 

as the latter fungus forms intricate, branched, brown conidia 

(Kirk 1981), unlike those of C. germanicum. Phylogenetically C. 

germanicum forms part of the C. abietis species complex. 

Catenulostroma macowanii (Sacc.) Crous & U. Braun, comb. 


Basionym: Coniothecium macowanii Sacc., Syll. Fung. 4: 512. 

1886. 

≡ Coniothecium punctiforme G. Winter, Hedwigia 24: 33. 1885, non C. 

punctiforme Corda, Icones Fungorum (Prague) 1: 2. 1837. 

≡ Trimmatostroma macowanii (Sacc.) M.B. Ellis, More Dematiacous 

Hyphomycetes: 29. 1976. 

Catenulostroma microsporum, see Teratosphaeria microspora. 

Catenulostroma protearum (Crous & M.E. Palm) Crous & U. 


Basionym: Trimmatostroma protearum Crous & M.E. Palm, Mycol. 

Res. 103: 1303. 1999. 

Cibiessia Crous, Fungal Diversity 26: 151. 2007. 

Type species: Cibiessia dimorphospora Crous & C. Mohammed, 

Fungal Diversity 26: 151. 2007. 

Description: Crous et al. (2007b; figs 3–5). 

Notes: The genus Cibiessia was introduced to accommodate 

species with chains of disarticulating conidia (arthroconidia). Some 

species have been shown to have a Readeriella synanamorph. 

Devriesia Seifert & N.L. Nick., Can. J. Bot. 82: 919. 2004. 

Type species: Devriesia staurophora (W.B. Kendr.) Seifert & N.L. 

Nick., Canad. J. Bot. 82: 919. 2004. 

Description: Seifert et al. (2004; figs 2–42). 

Notes: The genus is characterised by producing chains of pale 

brown, subcylindrical to fusiform, 0–1-septate conidia with 

somewhat thickened, darkened hila, forming chlamydospores in 

culture, and being heat resistant. Morphologically they resemble 

taxa placed in Pseudocladosporium U. Braun (= Fusicladium 

Bonord.; Venturiaceae), though phylogenetically Devriesia is not 

allied to this family. 

Hortaea Nishim. & Miyaji, Jap. J. Med. Mycol. 26: 145. 1984. 

Type species: Hortaea werneckii (Horta) Nishim. & Miyaji, Jap. J. 

Med. Mycol. 26: 145. 1984. 

Description: de Hoog et al. (2000, illust. p. 721). 


Notes: The genus forms brown, septate, thick-walled hyphae, with 

ellipsoidal, 0–1-septate (becoming muriformly septate), hyaline to 

pale brown conidia forming directly on the hyphae, via phialides 

with percurrent proliferation. Isolates of H. werneckii are restricted 

to tropical or subtropical areas, where they occur as halophilic 

saprobes, frequently being associated with tinea nigra of humans 

(de Hoog et al. 2000). The generic distinction with Capnobotryella 

is less clear, except that the latter tends to have darker, thick-walled 

conidia, and reduced, less prominent phialides. 

Penidiella Crous & U. Braun, gen. nov. MycoBank MB504463. 

Etymology: Named after its penicillate conidiophores. 

Differt a Periconiellae conidiophoris apice penicillato ex cellulis conidiogenis et 

ramoconidiis compositis, cellulis conidiogenis saepe 1–3(–4) locis conidiogenis, 

terminalibus vel subterminalibus, subdenticulatis, non vel subincrassatis, non vel 

leviter fuscatis-refractivis, ramoconidiis praesentibus, saepe numerosis, conidiis 

ramicatenatis. 

Mycelium consisting of branched, septate, smooth to verruculose, 

subhyaline to pale brown hyphae. Conidiophores macronematous, 

occasionally also with some micronematous conidiophores; 

macronematous conidiophores arising from superficial mycelium 

or stromata, solitary, fasciculate or in synnemata, erect, brown, 

thin- to thick-walled, smooth to finely verruculose; terminally 

penicillate, branched terminal part consisting of a conidiogenous 

apparatus composed of a series of conidiogenous cells and/or 

ramoconidia. Conidiogenous cells integrated, terminal, intercalary 

or pleurogenous, unbranched, pale to medium brown, smooth 

to finely verruculose, tapering to a flattened or rounded apical 

region or tips slightly inflated, polyblastic, sympodial, giving rise 

to a single or several sets of ramoconidia on different levels; with 

relatively few conidiogenous loci, 1–3(–4), terminal or subterminal, 

subdenticulate, denticle-like loci usually conical, terminally 

truncate, usually unthickened or at most very slightly thickened, not 

to slightly darkened or somewhat refractive. Conidia in branched 

acropetal chains. Ramoconidia 0–1-septate, pale to medium brown, 

smooth to verruculose, thin-walled, ellipsoidal, obovoid, fusiform, 

subcylindrical to obclavate; conidia subcylindrical, fusiform to 

ellipsoid-ovoid, 0–1-septate, pale olivaceous to brown, smooth to 

verruculose, thin-walled, catenate; hila truncate, unthickened or 

almost so, barely to somewhat darkened-refractive. 

Type species: Penidiella columbiana Crous & U. Braun, sp. nov. 

Notes: Three ramichloridium-like genera cluster within Capnodiales, 

namely Periconiella Sacc. [type: P. velutina (G. Winter) Sacc.], 

Ramichloridium Stahel ex de Hoog [type: R. apiculatum (J.H. 

Mill., Giddens & A.A. Foster) de Hoog] and Penidiella [type: P. 

columbiana Crous & U. Braun]. All three genera have brown, 

macronematous conidiophores with similar conidial scars. Within 

this complex, Ramichloridium is distinct in having a prominent 

rachis giving rise to solitary conidia. Periconiella and Penidiella 

are branched in the apical part of their conidiophores, and lack 

a rachis. In Periconiella conidia are solitary or formed in short, 

mostly simple chains, ramoconidia are lacking. The apical 

conidiogenous apparatus is composed of conidiogenous cells or 

branches with integrated, usually terminal conidiogenous cells, 

which are persistent. The conidiogenous cells are subcylindrical 

to somewhat clavate, usually not distinctly attenuated towards the 

tip, and have several, often numerous loci, aggregated or spread 

over the whole cell, terminal to usually lateral, flat, non-protuberant, 

not denticle-like, usually distinctly thickened and darkened, at least 

at the rim. In contrast, Penidiella has a quite distinct branching 

17

Crous et al. 

system, consisting of a single terminal conidiogenous cell giving 

rise to several ramoconidia that form secondary ramoconidia, etc., 

or the branched apparatus is composed of several terminal and 

sometimes lateral conidiogenous cells giving rise to sequences of 

ramoconidia (conidiogenous cells and ramoconidia are often barely 

distinguishable, with conidiogenous cells disarticulating, becoming 

ramoconidia). The branched apparatus may be loose to dense, 

metula-like. The conidiogenous cells have only few, usually 1–3 

(–4), terminal or subterminal subdenticulate loci, and ramoconidia 

are prominent and numerous, giving rise to branched chains of 

secondary conidia with flat-tipped hila. Some species of Penidiella 

with compact, metula-like branched apices are morphologically 

close to Metulocladosporiella Crous, Schroers, J.Z. Groenew., U. 

Braun & K. Schub. (Crous et al. 2006d). This genus encompasses 

two species of banana diseases belonging to Herpotrichiellaceae 

(Chaetothyriales), characterised by having conidiophore bases 

with rhizoid hyphal appendages and abundant micronematous 

conidiophores. Penidiella species with less pronounced penicillate 

apices, e.g. P. strumelloidea (Milko & Dunaev) Crous & U. Braun, are 

comparable with species of the genus Pleurotheciopsis B. Sutton 

(see Ellis 1976). The latter genus is distinct in having unbranched, 

often percurrently proliferating conidiophores, lacking ramoconidia 

and colourless conidia formed in simple chains. 

Cladosporium helicosporum R.F. Castañeda & W.B. Kendr. 

(Castañeda et al. 1997) is another penidiella-like fungus with 

terminally branched conidiophores, subdenticulate conidiogenous 

loci and conidia in long acropetal chains, but its affinity to Penidiella 

has still to be proven. 

Key to Penidiella species 

1. Conidiophores in vivo in well-developed, dense fascicles and distinct synnemata arising from a basal stroma; on fallen leaves of Ficus 

sp., Cuba ............................................................................................................................................................................... P. cubensis 

1. Conidiophores solitary, at most loosely aggregated .................................................................................................................................. 2 

2. Conidiophores with a terminal conidiogenous cell, often somewhat swollen, giving rise to several ramoconidia (on one level) that form 

chains of straight to distinctly curved conidia; isolated from leaf of Carex sp., Russia ..................................................... P. strumelloidea 

2. Penicillate apex of the conidiophores composed of a system of true branchlets, conidiogenous cells and ramoconidia or at least a sequence 

of ramoconidia on several levels; conidia usually straight ......................................................................................................................... 3 

3. Mycelium verruculose; long filiform conidiophores ending with a subdenticulate cell giving rise to sets of penicillate conidiogenous cells or 

ramoconidia which are barely distinguishable and turn into each other; ramoconidia and conidia consistently narrow, (1.5–)2(–2.5) µm 

wide, and aseptate, ramoconidia sometimes heterochromous; on living leaves of Nectandra coriacea, Cuba ................... P. nectandrae 

3. Mycelium more or less smooth; penicillate apex at least partly with true branchlets; conidia wider, 2–5 µm, at least partly septate, uniformly 

pigmented ................................................................................................................................................................................................. 4 

4. Hyphae, conidiophores and conidia frequently distinctly constricted at the septa; penicillate apex of the conidiophores sparingly developed, 

branchlets more or less divergent; isolated from leaf litter of Smilax sp., Cuba .................................................................. P. rigidophora 

4. Hyphae and conidia without distinct constrictions at the septa; penicillate apex of the conidiophores usually well-developed, with abundant 

branchings ................................................................................................................................................................................................. 5 

5. Conidiophores short, up to 120 × 3–4 µm, frequently with intercalary conidiogenous cell, swollen at the conidiogenous portion just below 

the upper septum which render the conidiophores subnodulose to distinctly nodulose, apex ± loosely penicillate; conidia (4–)5–7(–8) µm 

long; occasionally with micronematous conidiophores; isolated from man with tinea nigra, Venezuela .......................... P. venezuelensis 

5. Conidiophores much longer, up to 800 µm, 7–9 µm wide at the base, not distinctly nodulose, penicillate apex loose to often more 

compact, tight, metula-like; conidia longer, 7–25 × 2–5 µm; micronematous conidiophores lacking; isolated from dead leaf of Paepalanthus 

columbianus, Colombia ........................................................................................................................................................ P. columbiana 

Penidiella columbiana Crous & U. Braun, sp. nov. MycoBank 

MB504510. Figs 8–9. 

Etymology: Named after its country of origin, Colombia. 

Mycelium ex hyphis ramosis, septatis, levibus, pallide brunneis, 2–3 µm latis 

compositum. Conidiophora ex hyphis superficialibus oriunda, penicillata, erecta, 

brunnea, crassitunicata, minute verruculosa, ad 800 µm longa, ad basim 7–9 

µm lata, ad apicem pluriramosa, ex ramibus diversibus et cellulis conidiogenis 

composita, ramibus primariis (–2) subcylindraceis, 1–7-septatis, 50–120 × 4–6 µm; 

ramibus secundariis (–2) subcylindraceis, 1–5-septatis, 40–60 × 4–6 µm; ramibus 

tertiariis et subsequentibus 1–4-septatis, 10–30 × 3–5 µm. Cellulae conidiogenae 

terminales vel laterales, non ramosae, 5–15 × 3–5 µm, modice brunneae, minute 

verruculosae, apicem versus attenuatae, truncatae vel rotundatae, polyblasticae, 

sympodiales, cicatrices conidiales incrassatae, sed leviter fuscatae et non 

refractivae. Ramoconidia 0–1-septata, modice brunnea, levia, ellipsoidea, obclavata 

vel obovoidea, cum 1–3 hilis terminalibus, 10–20 × 3–5 µm; conidia subcylindrica 

vel ellipsoidea, 0–1-septata, pallide brunnea, catenata (–10), hila truncata, non 

incrassata, vix vel leviter fuscata. 

Mycelium consisting of branched, septate, smooth, pale brown, 2–3 

µm wide hyphae. Conidiophores arising from superficial mycelium, 

terminally penicillate, erect, brown, wall up to 1 µm wide, almost 

smooth to finely verruculose, up to 800 µm tall, 7–9 µm wide at 

the base; conidiogenous region consisting of a series of branches 

composed of true branchlets, conidiogenous cells and ramoconidia, 

branched portion usually rather compact, even metula-like, but 

also looser, with divergent branches; primary branches (–2), 

subcylindrical, 1–7-septate, 50–120 × 4–6 µm; secondary branches 

(–2), subcylindrical, 1–5-septate, 40–60 × 4–6 µm; tertiary and 

additional branches 1–4-septate, 10–30 × 3–5 µm. Conidiogenous 

cells terminal, intercalary or lateral, unbranched, 5–15 × 3–5 

µm, medium brown, finely verruculose, tapering to a flattened or 

rounded (frequently swollen) apical region, scars thickened, but 

only somewhat darkened, not refractive. Ramoconidia 0–1-septate, 

18


Fig. 8. Penidiella columbiana (type material). A. Conidiophores on pine needle in vitro. B–H. Conidiophores with chains of disarticulating conidia. Scale bars: A = 450, B–C = 

10 µm. 

medium brown, smooth, wall ≤ 1 µm wide, ellipsoidal to obclavate 

or obovoid, with 1–3 apical hila, 10–25 × 3–5 µm, ramoconidia 

with broadly truncate base, not or barely attenuated, up to 4 µm 

wide, or at least somewhat attenuated at the base, hila 1.5–3 µm 

wide. Conidia subcylindrical to ellipsoid, 0(–1)-septate, pale brown, 

in chains of up to 10, 7–15 × 2–3 µm, hila truncate, unthickened, 

barely to somewhat darkened, 1–2 µm wide. 


with moderate aerial mycelium and smooth, even, submerged 

margins; olivaceous-grey in central part, iron-grey in outer region 

(surface); colonies fertile. 

Specimen examined: Colombia, Páramo de Guasca, 3400 m alt., isolated from 

dead leaf of Paepalanthus columbianus (Eriocaulaceae), Aug. 1980, W. Gams, 

holotype CBS H-19937, culture ex-type CBS 486.80. 

Notes: This isolate was originally identified as belonging to the 

Stenella araguata species complex. The latter name has been 

somewhat confused in the literature, and has been incorrectly 

applied to isolates associated with opportunistic human infections 

(de Hoog et al. 2000). The “araguata” species complex is treated 

elsewhere in the volume (see Crous et al. 2007a – this volume). 


Penidiella cubensis (R.F. Castañeda) U. Braun, Crous & R.F. 

Castañeda, comb. nov. MycoBank MB504511. Fig. 10. 

Basionym: Cladosporium cubense R.F. Castañeda, Fungi Cubenses 

II (La Habana): 4. 1987. 

In vivo: Colonies on fallen leaves, amphigenous, effuse, pilose, 

brown. Mycelium usually external, superficial, but also internal, 

composed of branched, septate, brown, thin-walled, smooth to 

rough-walled hyphae, 2–3 µm wide. Stromata present, 40–80 µm 

diam, brown, immersed. Conidiophores densely fasciculate or in 

distinct synnemata, arising from stromata, erect, synnemata up 

to about 1000 µm long and (10–)20–40(–50) µm wide, individual 

threads filiform, pluriseptate throughout, brown, thin-walled (≤ 

0.5 µm), smooth or almost so to distinctly verruculose, apically 

penicillate. Conidiogenous cells integrated, terminal and intercalary, 

10–30 µm long, subcylindrical, terminal conidiogenous cells often 

slightly enlarged at the tip, with (1–)2–3(–4) terminal or subterminal 

subdenticulate conidiogenous loci, short conically truncate, 1–2 

µm diam, unthickened or almost so, but often slightly refractive or 

darkened-refractive, intercalary conidiogenous cells usually with a 

single lateral locus just below the upper septum, conidiogenous cells 

giving rise to a single set of primary ramoconidia, or a sequence 

of ramoconidia at different levels. Ramoconidia cylindrical to 

ellipsoid-fusoid, 8–18(–25) × 2–3 µm, aseptate, pale olivaceous, 

olivaceous-brown to brown, thin-walled, smooth or almost so to 

19

Crous et al. 

Fig. 9. Penidiella columbiana (type material). A. 

Conidiophores. B. Ramoconidia. C. Secondary conidia. 

Scale bar = 10 µm. U. Braun del. 

faintly verruculose, ramoconidia with broadly truncate base, barely 

narrowed, or ramoconidia more or less attenuated at the base, hila 

1–2 µm wide, unthickened or almost so, but often slightly refractive 

or darkened-refractive. Conidia in long acropetal chains, narrowly 

ellipsoid-ovoid, fusiform, 5–12(–15) × (1–)1.5–3 µm, aseptate, pale 

olivaceous to brownish, thin-walled, smooth to faintly rough-walled, 

ends attenuated, hila 1–1.5 µm wide, unthickened, not darkened, 

at most somewhat refractive. 

Specimen examined: Cuba, Guantánamo, Maisí, on fallen leaves of Ficus sp., 24 

Apr. 1986, M. Camino, holotype INIFAT C86/134 (HAL 2019 F, ex holotype). 

Notes: Cladosporium cubense was not available in culture and 

molecular sequence data are not available, but type material could 

be re-examined and revealed that this species is quite distinct from 

Cladosporium s. str., but agreeing with the concept of the genus 

Penidiella. Penidiella cubensis differs from all other species of this 

genus in having densely fasciculate conidiophores to synnematous 

conidiomata, arising from stromata. 

Penidiella nectandrae Crous, U. Braun & R.F. Castañeda, nom. 


Basionym: Cladosporium ferrugineum R.F. Castañeda, Fungi 

Cubenses II (La Habana): 4. 1987, homonym, non C. ferrugineum 

Allesch., 1895. 

In vivo: Colonies amphigenous, brown. Mycelium internal and 

external, superficial, composed of sparingly branched hyphae, 

septate, 1–3 µm wide, pale olivaceous-brown or brown, thinwalled 

(≤ 0.5 µm), smooth or almost so to distinctly verruculose, 

fertile cells giving rise to conidiophores somewhat swollen at the 

branching point, up to 5 µm diam, and somewhat darker. Stromata 

lacking. Conidiophores erect, straight, filiform, up to 350 µm long, 

2.5–4 µm wide, pluriseptate throughout, brown, darker below and 

paler above, thin-walled, smooth, apex penicillate, terminal cell of 

the conidiophore with 2–4 short denticle-like loci giving rise to sets 

of conidiogenous cells or ramoconidia that then form a sequence of 

new sets of ramoconidia on different levels, i.e., the loose to dense, 

metula-like branching system is composed of conidiogenous cells 

and ramoconidia which are often barely distinguishable and turn 

into each other; conidiogenous loci terminal or subterminal, usually 

1–3(–4), subdenticulate, 1–2 µm diam, conical, apically truncate, 

unthickened or almost so, not to somewhat darkened-refractive. 

Ramoconidia with truncate base, barely attenuated, or ramoconidia 

distinctly attenuated at the truncate base, up to 20 × 2 µm, aseptate, 

at the apex with 2–3(–4) subdenticulate hila, subcylindrical, 

20


very pale olivaceous, olivaceous-brown to brown, sometimes 

with different shades of brown (heterochromatous), thin-walled 

(≤ 0.5 µm), smooth to faintly verruculose. Conidia in long acropetal 

chains, narrowly ellipsoid-ovoid, fusiform to cylindrical, 5–16 × 

(1.5–)2(–2.5) µm, aseptate, very pale olivaceous, olivaceousbrown 

to brown, thin-walled, smooth to very faintly rough-walled, 

primary conidia with rounded apex and trunacte base, somewhat 

attenuated, secondary conidia truncate at both ends, hila 1–1.5 

µm diam, unthickened or almost so, at most slightly darkenedrefractive. 

Cultural characteristics: Colonies on PDA slimy, smooth, spreading; 

aerial mycelium absent, margins smooth, irregular; surface black 

with patches of cream. Colonies reaching 20 mm diam after 1 mo 

at 25 °C in the dark; colonies sterile on PDA, SNA and OA. 

Specimen examined: Cuba, Matanzas, San Miguel de los Baños, isolated from 

living leaves of Nectandra coriacea (Lauraceae), 24 Jan. 1987, R.F. Castañeda and 

G. Arnold, holotype INIFAT C87/45, culture ex-type CBS 734.87, and HAL 2018 F 

(ex-holotype). 

Notes: Although the ex-type strain of Cladosporium ferrugineum 

is sterile, its LSU DNA phylogeny reveals it to be unrelated to 

Cladosporium s. str. (see Fig. 1 in Crous et al. 2007a – this volume). 

Based on a re-examination of the type material it could clearly be 

shown that the morphology of this species fully agrees with the 

concept of the new genus Penidiella, which is supported by its 

phylogenetic position within Capnodiales. 

Fig. 10. Penidiella cubensis (type material). A. Swollen stromatic base of synnema. 

B. Conidiophores. C. Ramoconidia. D. Secondary conidia. Scale bar = 10 µm. 

U. Braun del. 

Penidiella rigidophora Crous, R.F. Castañeda & U. Braun, sp. 

nov. MycoBank MB504513. Figs 12–13. 

≡ Cladosporium rigidophorum R.F. Castañeda, nom. nud. (herbarium 

name). 

Differt a specibus Penidiellae conidiophoris dimorphosis, hyphis et conidiis ad septa 

saepe distincte constrictis. 

Mycelium consisting of strongly branched, septate, smooth or 

almost so, pale olivaceous to medium brown, guttulate, commonly 

constricted at septa, 2–6 µm wide hyphae, swollen cells up to 8 

µm wide, wall up to 1(–1.5) µm wide. Conidiophores dimorphic. 

Macronematous conidiophores separate, erect, subcylindrical, 

predominantly straight to slightly curved, terminally loosely 

penicillate, up to 120 µm long, and 4–5 µm wide at the base, which 

is neither lobed nor swollen, and lacks rhizoids, up to 10-septate, 

medium to dark brown, wall up to 1(–1.5) µm wide. Micronematous 

conidiophores erect, subcylindrical, up to 40 µm tall, 3–4 µm wide, 

1–3-septate, pale to medium brown (concolorous with hyphae). 

Conidiogenous cells predominantly terminal, rarely intercalary, 

medium brown, smooth, subcylindrical, but frequently swollen at 

apex, 10–20 × 5–6 µm, loci (predominantly single in micronematous 

conidiophores, but up to 4 in macronematous conidiophores) flattipped, 

sub-denticulate or not, 1–1.5 µm wide, barely to slightly 

darkened and thickened-refractive. Conidia in branched chains, 

medium brown, verruculose, (appearing like small spines under 

light microscope), ellipsoid to cylindrical-oblong, up to 1(–1.5) 

µm wide, frequently constricted at septa, which turn dark with 

age; ramoconidia (10–)13–17(–25) × 3–4(–5) µm, 1(–3)-septate; 

secondary conidia (7–)8–10(–12) × 3–4(–5); hila unthickened to 

very slightly thickened and darkened, not refractive, (0.5–)1(–1.5) 

µm. 

Fig. 11. Penidiella nectandrae (type material). A. Conidiophores. B. Ramoconidia. 

C. Secondary conidia. Scale bar = 10 µm. U. Braun del. 


with lobate margins and moderate aerial mycelium; iron-grey 

(surface), with a greenish black margin; reverse greenish black. 

Colonies reaching 20 mm diam after 1 mo at 25 °C in the dark; 

colonies fertile. 


21

Crous et al. 

Fig. 12. Penidiella rigidophora (type material). A–F. Micronematous conidiophores giving rise to chains of conidia. G–H. Macronematous conidiophores (note base in G, and 

apex in H). I. Conidia. Scale bars = 10 µm. 

Specimen examined: Cuba, isolated from leaf litter of Smilax sp. (Smilacaceae), 6. 

Nov. 1994, R.F. Castañeda, holotype CBS H-19938, culture ex-type CBS 314.95. 

Notes: Cladosporium rigidophorum is a herbarium name, which was 

never validly published. The ex-type strain, however, represents a 

new species of Penidiella, for which a valid name with Latin diagnosis 

is herewith provided. This species is easily distinguishable from all 

other taxa of Penidiella by forming distinct constrictions at hyphal 

and conidial septa as well as micronematous conidiophores (except 

for P. venezuelensis in which a few micronematous conidiophores 

have been observed). It is also phylogenetically distinct from the 

other taxa of Penidiella (see Fig. 1 in Crous et al. 2007a – this 

volume). 

22


Penidiella strumelloidea (Milko & Dunaev) Crous & U. Braun, 

comb. nov. MycoBank MB504514. Figs 14–15. 

Basionym: Cladosporium strumelloideum Milko & Dunaev, Novosti 

Sist. Nizsh. Rast. 23: 134. 1986. 

Mycelium consisting of branched, septate, smooth, hyaline to 

pale olivaceous, 1–4 µm wide hyphae, sometimes constricted at 

somewhat darker septa. Conidiophores solitary, erect, arising from 

superficial mycelium, micronematous, i.e., reduced to conidiogenous 

cells, or macronematous, subcylindrical, straight to slightly curved, 

subcylindrical throughout or often somewhat attenuated towards the 

apex, 12–80 × (2–)2.5–4 µm, 0–6-septate, medium brown, smooth, 

wall ≤ 0.75 µm, penicillate apex formed by a terminal conidiogenous 

cell giving rise to a single set of ramoconidia. Conidiogenous cells 

terminal, integrated, subcylindrical, straight, 8–12 × 1.5–2(–2.5) 

µm, pale brown, thin-walled, smooth, apex obtusely rounded to 

somewhat clavate; loci terminal, occasionally subterminal or lateral, 

unthickened or almost so to slightly thickened and darkened, not 

refractive, 1–1.5 µm wide. Conidia in branched chains; ramoconidia 

subcylindrical, with 1–3 terminal loci, olivaceous-brown, smooth; 

secondary conidia ellipsoidal, with one side frequently straight and 

the other convex, straight to slightly curved, (8–)10–12(–20) × 2(– 

3) µm, subhyaline to olivaceous-brown, smooth, thin-walled; hila 

unthickened or almost so to somewhat thickened and darkened, 

not refractive, 1 µm wide. 


with abundant, dense to woolly aerial mycelium, and uneven, 

feathery margins; surface pale olivaceous grey, reverse iron-grey. 



Fig. 13. Penidiella rigidophora (type material). A. Hyphae. B. Conidiophores. C. 

Ramoconidia. D. Secondary conidia. Scale bar = 10 µm. U. Braun del. 

Specimen examined: Russia, Yaroslavl Region, Rybinsk Reservoir, mouth of 

Sutka River, isolated from leaf of Carex sp. (Cyperaceae), from stagnant water, S. 

Ozerskaya, holotype BKMF-2534, culture ex-type CBS 114484. 

Fig. 14. Penidiella strumelloidea (type 

material). A–B. Micronematous conidiophores. 

C–D. Macronematous conidiophores. E–G. 

Conidia. Scale bars = 10 µm. 


23

Crous et al. 

conidiophores, about 10–15 × 2–3 µm. Conidiogenous cells 

terminal and intercalary, unbranched, subcylindrical, 5–12 × 3–4 

µm, medium brown, smooth or almost so to finely verruculose, 

apex of conidiogenous cells frequently swollen, up to 6 µm diam, 

with 1–3(–4) flat-tipped, non to slightly thickened, non to slightly 

darkened-refractive loci, 1–1.5 µm wide, frequently appearing 

subdenticulate, up to 1.5 µm long, intercalary conidiogenous cells 

also slightly swollen at the conidiogenous portion just below the 

upper septum, which render the conidiophores subnodulose to 

nodulose, swellings round about the conidiophore axis or unilateral. 

Conidia ellipsoid-ovoid, subcylindrical, pale to medium olivaceousbrown 

or brown, finely verruculose, wall ≤ 0.5 µm wide, guttulate or 

not, occurring in branched chains. Ramoconidia 0–1(–3)-septate, 

5–15(–22) × 3–4(–5) µm, with 1–3 subdenticulate apical hila; 

secondary conidia 0(–1)-septate, ellipsoid, obovoid to irregular, 

(4–)5–7(–8) × (2–)2.5–3(–4) µm; hila non to slightly thickened, non 

to slightly darkened-refractive, (0.5–)1(–1.5) µm wide. 

Cultural characteristics: Colonies on OA erumpent, spreading, 

with dense, compact aerial mycelium, and even, smooth margins; 

olivaceous-grey (surface), margins iron-grey. Colonies reaching 22 

mm diam after 1 mo at 25 °C in the dark. 

Specimen examined: Venezuela, isolated from man with tinea nigra, Jan. 1975, D. 

Borelli, holotype CBS H-19934, culture ex-type CBS 106.75. 

Fig. 15. Penidiella strumelloidea (type material). A. Hyphae. B. Conidiophores. C. 

Ramoconidia. D. Secondary conidia. Scale bars = 10 µm. U. Braun del. 

Notes: Penidiella strumelloidea resembles other species of 

Penidiella by having penicillate conidiophores with a conidiogenous 

apparatus giving rise to branched conidial chains. It differs, however, 

from all other species of this genus in having a rather simple 

penicillate apex composed of a single terminal conidiogenous 

cell giving rise to one set of ramoconidia which form frequently 

somewhat curved conidia. It is also phylogenetically distinct from 

the other taxa of Penidiella (see Fig. 1 in Crous et al. 2007a – this 

volume). 

Penidiella venezuelensis Crous & U. Braun, sp. nov. MycoBank 

MB504515. Figs 16–17. 

Etymology: Named after the geographic location of its type strain, 

Venezuela. 

Differt a P. columbiana conidiophoris bevioribus et angustioribus, ad 120 × 3–4 µm, 

subnodulosis, apice plus minusve laxe penicillatis et conidiis brevioribus, (4–)5–7(– 

8) µm longis. 

Mycelium consisting of branched, septate, smooth to faintly 

rough-walled, thin-walled, subhyaline, pale olivaceous to medium 

brown, (1.5–)2–3 µm wide hyphae. Conidiophores solitary, erect, 

macronematous, subcylindrical, straight to flexuous to once 

geniculate, up to 120 µm long, 3–4 µm wide, 1–12-septate, pale to 

medium olivaceous-brown or brown, thin-walled (up to about 1 µm), 

terminally penicillate, branched portion composed of true branchlets 

and/or a single set or several sets of ramoconidia, branchlets up 

to 50 µm long; occasionally with a few additional micronematous 

Notes: The type culture of Penidiella venezuelensis was originally 

determined as Stenella araguata from which it is, however, 

quite distinct by having smooth mycelium, long penicillate 

conidiophores with subdenticulate conidiogenous loci, smaller 

conidia, and agreeing with the concept of the genus Penidiella. It is 

phylogenetically distinct from the other taxa of Penidiella (see Fig. 

1 in Crous et al. 2007a – this volume). 

Pseudotaeniolina J.L. Crane & Schokn., Mycologia 78: 88. 1986. 

? = Friedmanniomyces Onofri, Nova Hedwigia 68: 176. 1999. 

Type species: Pseudotaeniolina convolvuli (Esfand.) J.L. Crane & 

Schokn., Mycologia 78: 88. 1986. 

Description: Crane & Schoknecht (1986, figs 3–19). 

Notes: No cultures or sequence data are available of the type 

species, and Pseudotaeniolina globosa De Leo, Urzì & de Hoog was 

placed in Pseudotaeniolina based on its morphology and ecology. 

The genus Friedmanniomyces is presently known from two species 

(Selbmann et al. 2005). Morphologically Friedmanniomyces is 

similar to Pseudotaeniolina, but fresh material of Pseudotaeniolina 

convolvuli needs to be recollected before this can be clarified. 

Readeriella Syd. & P. Syd., Ann. Mycol. 6: 484. 1908. 

= Kirramyces J. Walker, B. Sutton & Pascoe, Mycol. Res. 96: 919. 1992. 

= Colletogloeopsis Crous & M.J. Wingf., Canad. J. Bot. 75: 668. 1997. 

Synanamorphs: Cibiessia Crous, Fungal Diversity 26: 151. 2007; 

also pseudocercospora-like, see Crous (1998). 

Type species: Readeriella mirabilis Syd. & P. Syd., Ann. Mycol. 6: 

484. 1908. 


Notes: Several coelomycete genera are presently available 

to accommodate anamorphs of Capnodiales that reside in 

Teratosphaeriaceae, for which Readeriella is the oldest name. Other 

genera such as Phaeophleospora Rangel, Sonderhenia H.J. Swart 

& J. Walker and Lecanosticta Syd. belong to Mycosphaerellaceae. 

24


Fig. 16. Penidiella venezuelensis (type material). A. Microconidiophore. B. Apical part of macroconidiophore. C–F. Chains of conidia. Scale bars = 10 µm. 

Readeriella is polyphyletic within Teratosphaeriaceae. The 

recognition and circumscription (synonymy) of this genus follows the 

principles for anamorph genera within Capnodiales as outlined in the 

introduction to this volume. The only unifying character is conidial 

pigmentation, and the mode of conidiogenesis. Conidiogenous 

cells range from mono- to polyphialides with periclinal thickening, 

to phialides with percurrent proliferation, as observed in the type 

species, R. mirabilis (Fig. 18). Within the form genus conidia vary 

from aseptate to multiseptate, smooth to rough, and have a range 

of synanamorphs. Readeriella mirabilis has a synanamorph with 

cylindrical, aseptate conidia, while other species of Readeriella 

again have Cibiessia synanamorphs (scytalidium-like, with chains 

of dry, disarticulating conidia), suggesting the conidial morphology 

to be quite plastic. A re-examination of R. readeriellophora Crous & 

Mansilla revealed pycnidia to form a central cushion on which the 

conidiogenous cells are arranged (Fig. 18). This unique feature is 

commonly known in genera such as Coniella Höhn. and Pilidiella 

Petr. & Syd. (Diaporthales) (Van Niekerk et al. 2004), and has never 

been observed among anamorphs of the Capnodiales. Another 

species of Readeriella, namely “Phaeophleospora” toledana Crous 

& Bills, again forms paraphyses interspersed among conidiogenous 

cells, a rare feature in this group of fungi, while several species 

Fig. 17. Penidiella venezuelensis (type material). A. Hypha. B. Micronematous 

conidiophores. C. Macronematous conidiophores. D–E. Ramoconidia. F. Secondary 

conidia. Scale bar = 10 µm. U. Braun del. 


25

Crous et al. 

have conidiomata ranging from acervuli to pycnidia (Cortinas et 

al. 2006). Phylogenetically this coelomycete morphology, with 

its characteristic conidiogenesis, has evolved several times in 

Teratosphaeriaceae. 

Readeriella blakelyi (Crous & Summerell) Crous & U. Braun, 


Basionym: Colletogloeopsis blakelyi Crous & Summerell, Fungal 

Diversity 23: 342. 2006. 

Readeriella brunneotingens Crous & Summerell, sp. nov. 


Etymology: Named after the diffuse brown pigment visible in agar 

when cultivated on MEA. 

Readeriellae gauchensi similis, sed coloniis viridi-atris et pigmento brunneo in agaro 

diffundente distinguenda. 

Leaf spots amphigenous, irregular specks up to 3 mm diam, 

medium brown with a thin, raised, concolorous border. Conidiomata 

amphigenous, substomatal, exuding conidia in black masses; 

conidiomata pycnidial in vivo and in vitro, globose, brown to black, 

up to 120 µm diam; wall consisting of 3–4 cell layers of brown 

cells of textura angularis. Conidiogenous cells brown, verruculose, 

aseptate, doliiform to ampulliform, or reduced to inconspicuous loci 

on hyphae (in vitro), proliferating percurrently near the apex, 5–7 × 

3–5 µm; sympodial proliferation also observed in culture. Conidia 

brown, smooth to finely verruculose, ellipsoidal to subcylindrical, 

apex obtuse to subobtuse, tapering to a subtruncate or truncate 

base (1–1.5 µm wide) with inconspicuous, minute marginal frill, 

(5–)6–7(–8) × 2–3(–3.5) µm in vitro, becoming 1-septate; in older 

cultures becoming swollen, and up to 2-septate, 15 µm long and 

5 µm wide. 

Cultural characteristics: Colonies on MEA reaching 20 mm diam 

after 2 mo at 25 °C; colonies erumpent, aerial mycelium sparse to 

absent, margins smooth but irregularly lobate; surface irregularly 

folded, greenish black, with profuse sporulation, visible as oozing 

black conidial masses; a diffuse dark-brown pigment is also 

produced, resulting in inoculated MEA plates appearing darkbrown. 

Specimen examined: Australia, Queensland, Cairns, Eureka Creek, 48 km from 

Mareeba, S 17° 11’ 13.2”, E 145° 02’ 27.4”, 468 m, on leaves of Eucalyptus 

tereticornis, 26 Aug. 2006, P.W. Crous, CBS-H 19838 holotype, culture ex-type 

CPC 13303 = CBS 120747. 

Notes: Conidial dimensions of R. brunneotingens closely 

match those of Readeriella gauchensis (M.-N. Cortinas, Crous 

& M.J. Wingf.) Crous (Cortinas et al. 2006). The two species 

can be distinguished in culture, however, in that colonies of R. 

brunneotingens are greenish black in colour, sporulate profusely, 

and exude a diffuse, brown pigment into the agar, whereas colonies 

of R. gauchensis are more greenish olivaceous, and exude a yellow 

pigment into the agar (Cortinas et al. 2006). 

Readeriella considenianae (Crous & Summerell) Crous & U. 


Basionym: Colletogloeopsis considenianae Crous & Summerell, 

Fungal Diversity 23: 343. 2006. 

≡ Phaeophleospora destructans (M.J. Wingf. & Crous) Crous, F.A. Ferreira 

& B. Sutton, S. African J. Bot. 63: 113. 1997. 

Readeriella dimorpha (Crous & Carnegie) Crous & U. Braun, 


Basionym: Colletogloeopsis dimorpha Crous & Carnegie, Fungal 

Diversity 23: 345. 2006. 

Readeriella gauchensis (M.-N. Cortinas, Crous & M.J. Wingf.) 

Crous & U. Braun, comb. nov. MycoBank MB504521. 

Basionym: Colletogloeopsis gauchensis M.-N. Cortinas, Crous & 

M.J. Wingf., Stud. Mycol. 55: 143. 2006. 

Readeriella pulcherrima (Gadgil & M. Dick) Crous & U. Braun, 


Basionym: Septoria pulcherrima Gadgil & M. Dick, New Zealand J. 

Bot. 21: 49. 1983. 

≡ Stagonospora pulcherrima (Gadgil & M. Dick) H.J. Swart, Trans. Brit. 

Mycol. Soc. 90: 285. 1988. 

= Cercospora eucalypti Cooke & Massee, Grevillea 18: 7. 1889. 

≡ Kirramyces eucalypti (Cooke & Massee) J. Walker, B. Sutton & Pascoe, 

Mycol. Res. 96: 920. 1992. 

≡ Phaeophleospora eucalypti (Cooke & Massee) Crous, F.A. Ferreira & B. 

Sutton, S. African J. Bot. 63: 113. 1997. 

Notes: The epithet “eucalypti” is preoccupied by Readeriella 

eucalypti (Gonz. Frag.) Crous (Summerell et al., 2006), and thus 

the synonym “pulcherrima” becomes the next available name for 

this species. 

Readeriella readeriellophora, see Teratosphaeria readeriellophora. 

Fig. 18. 

Readeriella stellenboschiana (Crous) Crous & U. Braun, comb. 


Basionym: Colletogloeopsis stellenboschiana Crous, Stud. Mycol. 

55: 110. 2006. 

Readeriella zuluensis (M.J. Wingf., Crous & T.A. Cout.) Crous & 


Basionym: Coniothyrium zuluense M.J. Wingf., Crous & T.A. Cout., 

Mycopathologia 136: 142. 1997. 

≡ Colletogloeopsis zuluensis (M.J. Wingf., Crous & T.A. Cout.) M.-N. 

Cortinas, M.J. Wingf. & Crous (zuluense), Mycol. Res. 110: 235. 2006. 

Staninwardia B. Sutton, Trans. Br. Mycol. Soc. 57: 540. 1971. 

Type species: Staninwardia breviuscula B. Sutton, Trans. Br. Mycol. 

Soc. 57: 540. 1971. 

Description: Sutton (1971; fig. 1). 

Notes: The genus Staninwardia presently contains two species, 

namely S. breviuscula and Staninwardia suttonii Crous & Summerell 

(Summerell et al. 2006), though its placement in Capnodiales was 

less well resolved. The genus forms acervuli on brown leaf spots, 

with brown, catenulate conidia covered in a mucilaginous sheath. 

Readeriella destructans (M.J. Wingf. & Crous) Crous & U. Braun, 


Basionym: Kirramyces destructans M.J. Wingf. & Crous, S. African 

J. Bot. 62: 325. 1996. 

26


Fig. 18. A–E. Readeriella mirabilis. A. Conidium with conidial cirrus. B. Conidiogenous cells with percurrent proliferation. C. Macroconidia. D. Slightly pigmented, verruculose 

conidiogenous cell. E. Macro- and microconidia. F–I. Readeriella readeriellophora (type material). F. Colony on OA. G. Central stromatal tissue giving rise to conidiophores. H. 

Conidiogenous cells. I. Conidia. Scale bars = 10 µm. 

Fig. 19. Readeriella brunneotingens (type material). A. Leaf spot. B. Colony on MEA. C–D. Conidia. Scale bar = 10 µm. 


27

Crous et al. 

Schizothyriaceae clade 

Schizothyrium Desm., Ann. Sci. Nat., Bot., sér. 3: 11. 1849. 

Type species: Schizothyrium acerinum Desm., Ann. Sci. Nat., Bot., 

sér. 3: 11. 1849. 

Description: Batzer et al. (2007; figs 3–7). 

Notes: Species of Schizothyrium (Schizothyriaceae) have 

Zygophiala E.W. Mason anamorphs, and were recently shown to 

be allied to Mycosphaerellaceae (Batzer et al. 2007). Although 

species of Schizothyrium have thyrothecia, they cluster among 

genera with pseudothecial ascomata, questioning the value of this 

character at the family level. Based on its bitunicate asci and 1- 

septate ascospores, the teleomorph is comparable to others in the 

Capnodiales. 

Mycosphaerellaceae clade 

Mycosphaerella subclade 

Mycosphaerella Johanson, Öfvers. Förh. Kongl. Svenska 

Vetensk.-Akad. 41(9): 163. 1884. 

Type species: Mycosphaerella punctiformis (Pers. : Fr.) Starbäck, 

Bih. Kongl. Svenska Vetensk.-Akad. Handl. 15(3, 2): 9. 1889. 

Anamorph: Ramularia endophylla Verkley & U. Braun, Mycol. Res. 

108: 1276. 2004. 

Description: Verkley et al. (2004; figs 3–16). 

Notes: The genus Mycosphaerella has in the past been linked to 23 

anamorph genera (Crous et al. 2000), while additional genera have 

been linked via DNA-based studies, bringing the total to at least 30 

genera (Crous & Braun 2003, Crous et al. 2007b). However, based 

on ITS and SSU DNA phylogenetic studies and a reassessment of 

morphological characters and conidiogenesis, several anamorph 

genera have recently been reduced to synonymy (Crous & Braun 

2003, Crous et al. 2006a). Furthermore, the DNA sequence data 

generated to date clearly illustrate that the anamorph genera in 

Mycosphaerella are polyphyletic, residing in several clades within 

Mycosphaerella. If future collections not known from culture or DNA 

sequences are to be described in form genera, we recommend that 

the concepts as explained in Crous & Braun (2003) be used until 

such stage as they can be placed in Mycosphaerella, pending 

a modification of Art. 59 of the International Code of Botanical 

Nomenclature. The genus Mycosphaerella and its anamorphs 

represent a future topical issue of the Studies in Mycology, and will 

thus be treated separately. 

Dissoconium subclade 

Dissoconium de Hoog, Oorschot & Hijwegen, Proc. K. Ned. Akad. 

Wet., Ser. C, Biol. Med. Sci. 86(2): 198. 1983. 

Type species: Dissoconium aciculare de Hoog, Oorschot & 

Hijwegen, Proc. K. Ned. Akad. Wet., Ser. C, Biol. Med. Sci. 86(2): 

198. 1983. 

? = Uwebraunia Crous & M.J. Wingf., Mycologia 88: 446. 1996. 

Teleomorph: Mycosphaerella-like. 

Description: de Hoog et al. (1983), Crous (1998), Crous et al. 

(2004b; figs 3–10). 

Notes: The genus Dissoconium presently encompasses six 

species (Crous et al. 2007b), of which two, M. lateralis Crous & 

M.J. Wingf. (D. dekkeri de Hoog & Hijwegen), and M. communis 

Crous & Mansilla (D. commune Crous & Mansilla) are also known 

from their Mycosphaerella-like teleomorphs. No teleomorph genus 

will be introduced for this clade, however, until more sexual species 

have been collected to help clarify the morphological features of 

this genus. A further complication lies in the fact that yet other 

species, morphologically distinct from Dissoconium, also cluster in 

this clade (Crous, unpubl. data). 

“Passalora” zambiae subclade 

“Passalora” zambiae Crous & T.A. Cout., Stud. Mycol. 50: 209. 

2004. 


Notes: This fungus was placed in the form genus “Passalora” 

based on its smooth mycelium, giving rise to conidiophores forming 

branched chains of brown conidia with thickened, darkened, 

refractive hila. Although derived from single ascospores, the 

teleomorph material was lost, and thus it needs to be recollected 

before the relavance of its phylogenetic position can be fully 

understood. 

Additional teleomorph genera considered 

Coccodinium A. Massal., Atti Inst. Veneto Sci. Lett. Arti, Série 2, 

5: 336. 1860. (Fig. 20). 

Type species: Coccodinium bartschii A. Massal., Atti Inst. Veneto 

Sci. Lett. Arti, Série 2, 5: 337. 1860. 

Description: Eriksson (1981, figs 34–35). 

Notes: The genus Coccodinium (Coccodiniaceae) is characterised 

by having ascomata that are sessile on a subiculum, or somewhat 

immersed, semiglobose, collapsed when dry, brownish, uniloculate, 

with a centrum that stains blue in IKI (iodine potassium iodide). 

Asci are bitunicate, stalked, 8-spored, saccate, and have a thick, 

undifferentiated endotunica. Periphyses and periphysoids are 

well-developed and numerous. Ascospores are elongate, fusiform, 

ellipsoidal or clavate, transversely septate or muriform, hyaline or 

brownish (Eriksson 1981), and lack a mucous sheath. Based on a 

SSU sequence (GenBank accession U77668) derived from a strain 

identified as C. bartschii (Winka et al. 1998), Coccodinium appears 

to be allied to the taxa treated here in Teratosphaeria. Freshly 

collected cultures are relatively slow growing, and on MEA they 

form erumpent round, black colonies with sparse hyphal growth. On 

the surface of these colonies hyphal strands, consisting of brown, 

globose cells, give rise to conidia. Older cells (up to 15 µm diam) 

become fertile, giving rise to 1–3 conidia via inconspicuous phialidic 

loci. Conidia are fusoid-ellipsoidal to clavate, 3–5-septate, becoming 

constricted at the transverse septa, apex obtuse, base subtruncate, 

guttulate, smooth, widest in the upper third of the conidium, 15– 

40 × 4–7 µm. Phylogenetically Coccodinium is thus allied to the 

Chaetothyriales (Fig. 1), and not the Teratosphaeriaceae. 

28


Fig. 20. Coccodinium bartschii. A. Ascomata on host. B. Ostiolar area. C. Periphysoids. D–E. Ascospores shot onto agar. F–I. Asci with thick ectotunica. J–K. Young ascospores. 

L–M. Mature ascospores. N. Colony on MEA. O–Q. Conidiogenous cells giving rise to conidia. R–S. Conidia. Scale bars: A, N = 250, B, D, F–G, I, L–M, O = 10 µm. 


29

Crous et al. 

Fig. 21. Stigmidium schaereri. A. Lichenicolous habit on Dacampia hookeri. B. Vertical section through an ascoma. C–D. Asci. E–G. Ascospores. H. Older, brown ascospores. 

Scale bars = 10 µm. 

Stigmidium Trevis., Consp. Verruc.: 17. 1860. (Fig. 21). 

Type species: Stigmidium schaereri (A. Massal.) Trevis., Consp. 

Verruc.: 17. 1860. 

Description: Roux & Triebel (1994, figs 47–50). 

Notes: The type species of the genus is lichenicolous, characterised 

by semi-immersed, black, globose ascomata with ostiolar 

periphyses and periphysoids. Asci are 8-spored, fasciculate, 

bitunicate, (endotunica not giving a special reaction in Congo red 

or toluidine blue). Ascospores are fusoid-ellipsoidal, medianly 1- 

septate, guttulate, thin-walled, lacking a sheath. Presently no 

culture is available, and thus the placement of Stigmidium remains 

unresolved. 

Discussion 

From the LSU sequence data presented here, it is clear that 

Mycosphaerella is not monophyletic as previously suggested (Crous 

et al. 2001, Goodwin et al. 2001). The first step to circumscribe 

natural genera within this complex was taken by Braun et al. (2003), 

who separated Cladosporium anamorphs from this complex, 

and erected Davidiella (Davidiellaceae; Schoch et al. 2006) to 

accommodate their teleomorphs. The present study reinstates 

the genus Teratosphaeria for a clade of largely extremotolerant 

fungi (Selbmann et al. 2005) and foliar pathogens of Myrtaceae 

and Proteaceae (Crous et al. 2004a, b, 2006b, 2007b), and further 

separates generic subclades within the Mycosphaerellaceae, 

while Batzer et al. (2007) again revealed Schizothyrium Desm. 

(Schizothyriaceae) to cluster within the Mycosphaerellaceae. Our 

results, however, provide support for recognition of Schizothyrium as 

a distinct genus, although Schizothyriaceae was less well supported 

as being separate from Mycosphaerellaceae (Capnodiales). 

Although pleomorphism represents a rather unstudied 

phenomenon in this group of fungi, it has been observed in several 

species. Within the Teratosphaeria clade, Crous et al. (2007b) 

recently demonstrated teleomorphs to have Readeriella and 

Cibiessia synanamorphs, while the black yeast genera that belong 

to this clade, commonly have more than one anamorph state in 

culture. The present study also revealed Readeriella mirabilis to 

have two conidial types in culture, and to be highly plastic regarding 

its mode of conidiogenesis, and Readeriella to be the oldest generic 

name available for a large group of leaf-spotting coelomycetes in 

the Teratosphaeriaceae (Capnodiales). 

Although not commonly documented, there are ample 

examples of synanamorphs in Capnodiales. Within Mycosphaerella, 

Beilharz et al. (2004) described Passalora perplexa Beilharz, 

Pascoe, M.J. Wingf. & Crous as a species with a coelomycete 

and yeast synanamorph, while Crous & Corlett (1998) described 

Mycosphaerella stigmina-platani F.A. Wolf to have a Cercostigmina 

U. Braun and Xenostigmina Crous synanamorph, and recent 

collections also revealed the presence of a similar species that has 

typical “Stigmina” (distoseptate conidia) and Pseudocercospora 

(euseptate conidia) synanamorphs (Crous, unpubl. data), and 

Crous (1998) reported Readeriella epicoccoides (coelomycete) to 

30


have a Cercostigmina (hyphomycete) synanamorph in culture. 

Although the Mycosphaerella complex encompasses 

thousands of names, it may appear strange that it is only now that 

more clarity is obtained regarding the phylogenetic relationships 

among taxa in this group. This is partly due to the fact that these 

organisms are cultivated with difficulty, and also that the first paper 

to address the taxonomy of this complex based on DNA sequence 

data was only relatively recently published (Stewart et al. 1999). 

In the latter study, the genus Paracercospora Deighton (scars 

minutely thickened along the rim), was shown to be synonymous 

with the older genus Pseudocercospora. Similarily, Crous et al. 

(2001) showed that Cercostigmina (rough, irregular percurrent 

proliferations) was also synonymous with Pseudocercospora. 

This led Crous & Braun (2003) to conclude that conidiomatal type, 

conidial catenulation, septation and proliferation of conidiogenous 

cells were of less importance in separating species at the generic 

level. Mycovellosiella Rangel and Phaeoramularia Munt.-Cvetk. 

were subsequently reduced to synonymy with the older name, 

Passalora Fr., and characters identified as significant at the generic 

level were pigmentation (Cercospora vs. Passalora), scar structure 

(Passalora vs. Pseudocercospora), and verruculose superficial 

hyphae (Stenella vs. Passalora). Due to the unavailability of 

cultures, no decision was made regarding Stenella (verrucose 

conidia and mycelium), Stigmina (distoseptate conidia), and 

several other, less well-known genera such as Asperisporium 

Maubl., Denticularia Deighton, Distocercospora N. Pons & B. 

Sutton, Prathigada Subram., Ramulispora, Pseudocercosporidium 

Deighton, Stenellopsis B. Huguenin and Verrucisporota D.E. Shaw 

& Alcorn. In a recent study, however, Crous et al. (2006a) were 

able to show that Phaeoisariopsis (synnemata, conidia with slightly 

thickened hila) and Stigmina (distoseptate conidia) were also 

synonyms of Pseudocercospora. 

The present study shows that most anamorph genera are 

polyphyletic within Teratosphaeria, and paraphyletic within 

Capnodiales. In some cases, generic concepts of anamorphs 

based on morphology and conidium ontogeny conform well with 

phylogenetic relationships, though this is not true in all cases due 

to convergence. Nevertheless, anamorphs still convey valuable 

morphological information that is contained in the anamorph name, 

and naming anamorphs continue to provide a practical system to 

identify the various asexual taxa encountered. 

Acknowledgements 

We gratefully acknowledge several colleagues in different countries who have 

collected the material studied here, without which this work would not have been 

possible. We thank M. Vermaas for preparing the photographic plates. M. Grube is 

gratefully acknowledged for sending us fresh material of Stigmidium to include in 

this study, and D. Triebel and M. Grube are thanked for advice regarding the strain of 

hamathecial tissue in Stigmidium. K.A. Seifert is thanked for numerous discussions 

about anamorph concepts and advice pertaining to this paper, and for recollecting 

Coccodinium bartschii to enable us to clarify its phylogeny. J.K. Stone is thanked for 

commenting on an earlier draft version of this paper. 

References 



Barnes I, Crous PW, Wingfield BD, Wingfield MJ (2004). Multigene phylogenies 

reveal that red band needle blight of Pinus is caused by two distinct species of 

Dothistroma, D. septosporum and D. pini. Studies in Mycology 50: 551–565. 

Barr ME (1972). Preliminary studies on the Dothideales in temperate North America. 

Contributions from the University of Michigan Herbarium 9: 523–638. 

Batzer JC, Mercedes Diaz Arias M, Harrington TC, Gleason ML, Groenewald JZ, 

Crous PW (2007). Four species of Zygophiala (Schizothyriaceae, Capnodiales) 

are associated with the sooty blotch and flyspeck complex on apple. Mycologia: 

in press. 

Beilharz VC, Pascoe IG, Wingfield MJ, Tjahjono B, Crous PW (2004). Passalora 

perplexa, an important pleoanamorphic leaf blight pathogen of Acacia 

crassicarpa in Australia and Indonesia. Studies in Mycology 50: 471–479. 

Braun U, Crous PW, Dugan F, Groenewald JZ, Hoog GS de (2003). Phylogeny and 

taxonomy of cladosporium-like hyphomycetes, including Davidiella gen. nov., 

the teleomorph of Cladosporium s.str. Mycological Progress 2: 3–18. 

Butin H, Pehl L, Hoog GS de, Wollenzien U (1996). Trimmatostroma abietis sp. nov. 

(hyphomycetes) and related species. Antonie van Leeuwenhoek 69: 203–209. 

Castañeda RF, Kendrick B, Gené J (1997). Notes on conidial fungi. XIII. A new 

species of Cladosporium from Cuba. Mycotaxon 63: 183–187. 

Chupp C (1954). A monograph of the fungus genus Cercospora. Ithaca, New York. 

Published by the author. 

Cortinas M-N, Crous PW, Wingfield BD, Wingfield MJ (2006). Multi-gene phylogenies 

and phenotypic characters distinguish two species within the Colletogloeopsis 

zuluensis complex associated with Eucalyptus stem cankers. Studies in 

Mycology 55: 133–146. 

Crane JL, Schoknecht JD (1986). Revision of Torula and Hormiscium species. 

New names for Hormiscium undulatum, Torula equina, and Torula convolvuli. 

Mycologia 78: 86–91. 

Crous PW (1998). Mycosphaerella spp. and their anamorphs associated with leaf 

spot diseases of Eucalyptus. Mycologia Memoir 21: 1–170. 

Crous PW, Aptroot A, Kang JC, Braun U, Wingfield MJ (2000). The genus 

Mycosphaerella and its anamorphs. Studies in Mycology 45: 107–121. 

Crous PW, Braun U (2003). Mycosphaerella and its anamorphs. 1. Names published 

in Cercospora and Passalora. CBS Biodiversity Series 1: 1–571. 

Crous PW, Braun U, Schubert K, Groenewald JZ (2007a). Delimiting Cladosporium 

from morphologically similar genera. Studies in Mycology 58: 33–56. 

Crous PW, Corlett M (1998). Reassessment of Mycosphaerella spp. and their 

anamorphs occurring on Platanus. Canadian Journal of Botany 76: 1523– 

1532. 

Crous PW, Denman S, Taylor JE, Swart L, Palm ME (2004a). Cultivation and 

diseases of Proteaceae: Leucadendron, Leucospermum and Protea. CBS 

Biodiversity Series 2: 1–228. 

Crous PW, Groenewald JZ (2006a). Mycosphaerella alistairii. Fungal Planet No. 4. 

(www.fungalplanet.org). 

Crous PW, Groenewald JZ (2006b). Mycosphaerella maxii. Fungal Planet No. 6. 

(www.fungalplanet.org). 

Crous PW, Groenewald JZ, Gams W (2003). Eyespot of cereals revisited: ITS 

phylogeny reveals new species relationships. European Journal of Plant 

Pathology 109: 841–850. 

Crous PW, Groenewald JZ, Mansilla JP, Hunter GC, Wingfield MJ (2004b). 

Phylogenetic reassessment of Mycosphaerella spp. and their anamorphs 

occurring on Eucalyptus. Studies in Mycology 50: 195–214. 


genera in Mycosphaerella based on ITS rDNA sequence and morphology. 

Mycologia 93: 1081–1101. 

Crous PW, Liebenberg MM, Braun U, Groenewald JZ (2006a). Re-evaluating the 

taxonomic status of Phaeoisariopsis griseola, the causal agent of angular leaf 

spot of bean. Studies in Mycology 55: 163–173. 

Crous PW, Palm ME (1999). Systematics of selected foliicolous fungi associated 

with leaf spots of Proteaceae. Mycological Research 103: 1299–1304. 

Crous PW, Schroers HJ, Groenewald JZ, Braun U, Schubert K (2006d). 



Crous PW, Summerell BA, Carnegie AJ, Mohammed C, Himaman W, Groenewald 

JZ (2007b). Foliicolous Mycosphaerella spp. and their anamorphs on Corymbia 

and Eucalyptus. Fungal Diversity 26: 143–185. 

Crous PW, Wingfield MJ, Mansilla JP, Alfenas AC, Groenewald JZ (2006b). 


occurring on Eucalyptus. II. Studies in Mycology 55: 99–131. 

Crous PW, Slippers B, Wingfield MJ, Rheeder J, Marasas WFO, Phillips AJL, Alves 

A, Burgess T, Barber P, Groenewald JZ (2006c). Phylogenetic lineages in the 

Botryosphaeriaceae. Studies in Mycology 55: 235–253. 

Doherty MW (1900). New Species of Trimmatostroma. Botanical Gazette 30: 400– 

403. 



Ellis MB (1976). More dematiaceous hyphomycetes. Commonwealth Mycological 


Eriksson O (1981). The families of bitunicate ascomycetes. Opera Botanica 60: 

1–220. 


31

Crous et al. 

Gams W, Verkley GJM, Crous PW (2007). CBS course of mycology, 5 th ed. 

Centraalbureau voor Schimmelcultures, Utrecht. 

Goodwin SB, Dunkley LD, Zismann VL (2001). Phylogenetic analysis of Cercospora 

and Mycosphaerella based on the internal transcribed spacer region of 

ribosomal DNA. Phytopathology 91: 648–658. 

Gorbushina AA, Panina LK, Vlasov DY, Krumbein WE (1996). Fungi deteriorating 

Chersonessus marbles. Mikologia i Fitopatologia 30: 23–28. 

Hambleton S, Tsuneda A, Currah RS (2003). Comparative morphology and 

phylogenetic placement of two microsclerotial black fungi from Sphagnum. 

Mycologia 95: 959–975. 

Hoog GS de, Gerrits van den Ende AHG (1998). Molecular diagnostics of clinical 

strains of filamentous Basidiomycetes. Mycoses 41: 183–189. 

Hoog GS de, Guarro J, Gené J, Figueras MJ (2000). Atlas of Clinical Fungi. 2 nd 

Edn. Centraalbureau voor Schimmelcultures, Utrecht, and Universitat Rovira 

I Virgili, Reus. 

Hoog GS de, Oorschot CAN van, Hijwegen T (1983). Taxonomy of the Dactylaria 

complex. II. Proceedings van de Koninklijke Nederlandse Akademie van 

Wetenschappen, Series C, 86(2): 197–206. 

Hoog GS de, Zalar P, Urzì C, Leo de F, Yurlova NA, Sterflinger K (1999). Relationships 

of dothideaceous black yeasts and meristematic fungi based on 5.8S and ITS2 

rDNA sequence comparision. Studies in Mycology 43: 31–37. 

Hunter GC, Wingfield BD, Crous PW, Wingfield MJ (2006). A multi-gene phylogeny 

for species of Mycosphaerella occurring on Eucalyptus leaves. Studies in 

Mycology 55: 147–161. 

Kirk PM (1981). New or interesting microfungi. 1. Dematiaceous hyphomycetes from 

Devon. Transactions of the British Mycological Society 76: 71–87. 

Kogej T, Gorbushina AA, Gunde-Cimerman N (2006). Hypersaline conditions 

induce changes in cell-wall melanization and colony structure in a halophilic 

and a xerophilic black yeast species of the genus Trimmatostroma. Mycological 

Research 110: 713–724. 

Krumbein WE, Gorbushina AA, Sterflinger K, Haroska U, Kunert U, Drewello R, 

Weißmann R (1996). Biodetorioration of historical window panels of the 

former Cistercian Monastery church of Haina (Hessen, Germany). DECHEMA 

Monographs 133: 417–424. 

Moncalvo J-M, Rehner SA, Vilgalys R (1993). Systematics of Lyophyllum section 

Difformia based on evidence from culture studies and ribosomal DNA 

sequences. Mycologia 85: 788–794. 

Müller E, Oehrens E (1982). On the genus Teratosphaeria (Ascomycetes). Sydowia 

35: 138–142. 

Niekerk JM van, Groenewald JZ, Verkley GJM, Fourie PH, Wingfield MJ, Crous PW 

(2004). Systematic reappraisal of Coniella and Pilidiella, with specific reference 

to species occurring on Eucalyptus and Vitis in South Africa. Mycological 

Research 108: 283–303. 

Rayner RW (1970). A mycological colour chart. CMI and British Mycological Society. 

Kew. 

Rehner SA, Samuels GJ (1994). Taxonomy and phylogeny of Gliocladium analysed 

from nuclear large subunit ribosomal DNA sequences. Mycological Research 

98: 625–634. 

Roux C, Triebel D (1994). Révision des espèces de Stigmidium et de 

Sphaerellothecium (champignons lichénicoles non lichénisés, Ascomycetes) 

correspondant à Pharcidia epicymatia sensu Keissler ou à Stigmidium schaereri 

auct. Bulletin de la Societe Linneenne de Provence 45: 451–542. 

Schoch C, Shoemaker RA, Seifert, KA, Hambleton S, Spatafora JW, Crous PW 


Mycologia 98: 1043–1054. 





Seifert KA, Nickerson NL, Corlett M, Jackson ED, Louis-Seize G, Davies RJ 

(2004). Devriesia, a new hyphomycete genus to accommodate heat-resistant, 


Selbmann L, Hoog GS de, Mazzaglia A, Friedmann EI, Onofri S (2005). Fungi at 

the edge of life: cryptoendolithic black fungi from Antarctic desert. Studies in 

Mycology 51: 1–32. 

Stewart EL, Liu Z, Crous PW, Szabo L (1999). Phylogenetic relationships among 

some cercosporoid anamorphs of Mycosphaerella based on rDNA sequence 

analysis. Mycological Research 103: 1491–1499. 

Sugiyama J, Amano N (1987). Two metacapnodiaceous sooty moulds from Japan: 

their identity and behavior in pure culture. In: Pleomorphic fungi: the diversity 

and its taxonomic implications. (Sugiyama J, ed.). Kodansha, Tokyo/Elsevier, 

Amsterdam: 141–156. 

Summerell BA, Groenewald JZ, Carnegie AJ, Summerbell RC, Crous PW (2006). 

Eucalyptus microfungi known from culture. 2. Alysidiella, Fusculina and 

Phlogicylindrium genera nova, with notes on some other poorly known taxa. 


Sutton BC (1971). Staninwardia gen. nov. (Melanconiales) on Eucalyptus. 

Transactions of the British Mycological Society 57: 539–542. 

Sutton BC, Pascoe IG (1989). Reassessment of Peltosoma, Stigmina and 

Batcheloromyces and description of Hyphothyrium gen. nov. Mycological 

Research 92: 210–222. 

Taylor JE, Crous PW, Wingfield MJ (1999). The genus Batcheloromyces on 

Proteaceae. Mycological Research 103: 1478–1484. 

Taylor JE, Groenewald JZ, Crous PW (2003). A phylogenetic analysis of 

Mycosphaerellaceae leaf spot pathogens of Proteaceae. Mycological Research 

107: 653–658. 

Verkley GJM, Crous PW, Groenewald JZ, Braun U, Aptroot A (2004). Mycosphaerella 

punctiformis revisited: morphology, phylogeny, and epitypification of the type 

species of the genus Mycosphaerella (Dothideales, Ascomycota). Mycological 

Research 108: 1271–1282. 

Vilgalys R, Hester M (1990). Rapid genetic identification and mapping of 

enzymatically amplified ribosomal DNA from several Cryptococcus species. 

Journal of Bacteriology 172: 4238–4246. 

White TJ, Bruns T, Lee S, Taylor J (1990). Amplification and direct sequencing of 

fungal ribosomal RNA genes for phylogenetics. In: PCR Protocols: a guide to 

methods and applications (Innis MA, Gelfand DH, Sninsky JJ, White TJ, eds). 

Academic Press, San Diego, California: 315–322. 

Winka K, Eriksson OE, Bång Å (1998). Molecular evidence for recognizing the 

Chaetothyriales. Mycologia 90: 822–830. 

Wollenzien U, Hoog GS de, Krumbein WE, Urzì C (1995). On the isolation of 

microcolonial fungi occurring on and in marble and other calcareous rocks. 

Science of the Total Environment 167: 287–294. 

32


doi:10.3114/sim.2007.58.02 


Delimiting Cladosporium from morphologically similar genera 

P.W. Crous 1* , U. Braun 2 , K. Schubert 3 and J.Z. Groenewald 1 

1 

CBS Fungal Biodiversity Centre, P.O. Box 85167, 3508 AD Utrecht, The Netherlands; 2 Martin-Luther-Universität, Institut für Biologie, Geobotanik und Botanischer Garten, 

Herbarium, Neuwerk 21, D-06099 Halle (Saale), Germany; 3 Botanische Staatssammlung München, Menzinger Straße 67, D-80638 München, Germany 

*Correspondence: Pedro Crous, p.crous@cbs.knaw.nl 

Abstract: The genus Cladosporium is restricted to dematiaceous hyphomycetes with a coronate scar type, and Davidiella teleomorphs. In the present study numerous 

cladosporium-like taxa are treated, and allocated to different genera based on their morphology and DNA phylogeny derived from the LSU nrRNA gene. Several species are 

introduced in new genera such as Hyalodendriella, Ochrocladosporium, Rachicladosporium, Rhizocladosporium, Toxicocladosporium and Verrucocladosporium. A further new 

taxon is described in Devriesia (Teratosphaeriaceae). Furthermore, Cladosporium castellanii, the etiological agent of tinea nigra in humans, is confirmed as synonym of Stenella 

araguata, while the type species of Stenella is shown to be linked to the Teratosphaeriaceae (Capnodiales), and not the Mycosphaerellaceae as formerly presumed. 

Taxonomic novelties: Devriesia americana Crous & Dugan, sp. nov., Hyalodendriella Crous, gen. nov., Hyalodendriella betulae Crous sp. nov., Ochrocladosporium Crous & 

U. Braun, gen. nov., Ochrocladosporium elatum (Harz) Crous & U. Braun, comb. nov., Ochrocladosporium frigidarii Crous & U. Braun, sp. nov., Rachicladosporium Crous, U. 

Braun & Hill, gen. nov., Rachicladosporium luculiae Crous, U. Braun & Hill, sp. nov., Rhizocladosporium Crous & U. Braun, gen. nov., Rhizocladosporium argillaceum (Minoura) 

Crous & U. Braun, comb. nov., Toxicocladosporium Crous & U. Braun, gen. nov., Toxicocladosporium irritans Crous & U. Braun, sp. nov., Verrucocladosporium K. Schub., Aptroot 

& Crous, gen. nov., Verrucocladosporium dirinae K. Schub., Aptroot & Crous, sp. nov. 

Key words: Cladosporium, Davidiella, food spoilage, hyphomycetes, indoor air, LSU phylogeny, taxonomy. 

Introduction 

Cladosporioid hyphomycetes are common, widespread fungi. 

The genus Cladosporium Link is based on the type species, 

Cladosporium herbarum (Pers. : Fr.) Link, which in turn has been 

linked to Davidiella Crous & U. Braun teleomorphs (Braun et al. 

2003, Schubert et al. 2007b – this volume). Cladosporium is one 

of the largest, most heterogeneous genera of hyphomycetes, 

comprising more than 772 names (Dugan et al. 2004), and including 

endophytic, fungicolous, human pathogenic, phytopathogenic and 

saprobic species. Species of this genus affect daily human life in 

various ways. The common saprobic members of Cladosporium 

occur on all kinds of senescing and dead leaves and stems of 

herbaceous and woody plants, as secondary invaders on necrotic 

leaf lesions caused by other fungi, are frequently isolated from air, 

soil, food stuffs, paint, textiles and other organic matters, are also 

known to be common endophytes (Riesen & Sieber 1985, Brown 

et al. 1998, El-Morsy 2000) as well as phylloplane fungi (Islam & 

Hasin 2000, De Jager et al. 2001, Inacio et al. 2002, Stohr & Dighton 

2004, Levetin & Dorsey 2006). Furthermore, some Cladosporium 

species are known to be potential agents of medical relevance. 

Cladosporium herbarum is, for instance, a common contaminant in 

clinical laboratories and causes allergic lung mycoses (de Hoog et 

al. 2000, Schubert et al. 2007b – this volume). 

In spite of the enormous relevance of this genus, there is 

no comprehensive modern revision of Cladosporium, but some 

attempts to revise and monograph parts of it have been initiated 

during the last decade (David 1997, Partridge & Morgan-Jones 

2002, Wirsel et al. 2002, Braun et al. 2003, Dugan et al. 2004, Park 

et al. 2004, Seifert et al. 2004, Schubert & Braun 2004, 2005a, 

b, 2007, Heuchert et al. 2005, Schubert 2005a, b, Schubert et al. 

2006). 

Previous molecular studies employing rDNA ITS sequence 

data (Crous et al. 2001) have shown Cladosporium spp. to cluster 

adjacent to the main monophyletic Mycosphaerella Johanson 

cluster, suggesting a position apart from the latter genus. Braun 

et al. (2003) carried out more comprehensive sequence analyses, 

based on ITS (ITS-1, 5.8S and ITS-2) and 18S rDNA data, providing 

further evidence that Cladosporium s. str. represents a sister clade 

of Mycosphaerella. 

Various authors discussed the taxonomy and circumscription 

of Cladosporium (von Arx 1981, 1983, McKemy & Morgan-Jones 

1990, Braun 1995), reaching different conclusions. However, a 

first decisive revision of Cladosporium, leading to a more natural 

concept of this genus, was published by David (1997), who carried 

out comprehensive scanning electron microscopic examinations of 

the scar and hilum structure in Cladosporium and Heterosporium 

Klotzsch ex Cook. The first Scanning Electron Micrograph (SEM) 

studies of these structures, published by Roquebert (1981), indicated 

that the conidiogenous loci and conidial hila in Cladosporium are 

characterised by having a unique structure. David (1997) confirmed 

these observations, based on a wide range of Cladosporium and 

Heterosporium species, and demonstrated that the structures of the 

conidiogenous loci and hila in the latter genus fully agree with those 

of Cladosporium, proving that Heterosporium was indeed a synonym 

of Cladosporium s. str. He introduced the term “coronate” for the 

Cladosporium scar type, which is characterised by having a central 

convex part (dome), surrounded by a raised periclinal rim (David 

1997), and showed that this type is confined to anamorphs, as far 

as experimentally proven, connected with teleomorphs belonging 

in “Mycosphaerella” s. lat. These results were confirmed in a later 

phylogenetic study by Braun et al. (2003). Cladosporium s. str. was 

shown to be a sister clade to Mycosphaerella s. str., for which the 

new teleomorph genus Davidiella was proposed. Although no clear 

morphological differences were reported between Davidiella and 

Mycosphaerella, a further study by Aptroot (2006) found ascospores 

of Davidiella to have characteristic irregular cellular inclusions 

(lumina), which are absent in species of Mycosphaerella, along with 

periphysoids and pseudoparaphyses (Schubert et al. 2007b – this 

volume). Furthermore, a higher order phylogeny study by Schoch 

et al. (2006), which employed DNA sequence data of four loci (SSU 

nrDNA, LSU nrDNA, EF-1α, RPB2), revealed species of Davidiella 

to cluster in a separate family (Davidiellaceae) from species of 

Mycosphaerella (Mycosphaerellaceae), with both families residing 

in the Capnodiales (Dothideomycetes), and not Dothideales as 

always presumed. 

33

Crous et al. 

Table 1. Isolates for which new sequences were generated. 

Anamorph Teleomorph Accession number 1 Host Country Collector GenBank numbers 2 

(ITS, LSU) 

Cladoriella eucalypti CBS 115899*; CPC 10954 Eucalyptus sp. South Africa P.W. Crous EU040224, EU040224 

Coniothyrium palmarum CBS 758.73; CMW 5283 Phoenix dactylifera Israel Y. Pinkas DQ240000, EU040225 

Devriesia acadiensis CBS 115874; DAOM 232211 Soil Canada N. Nickerson AY692095, EU040226 

Devriesia americana CBS 117726; ATCC 96545; CPC 5121 Air U.S.A. F.M. Dugan AY251068, EU040227 

Devriesia shelburniensis CBS 115876; DAOM 232217 Soil Canada N. Nickerson AY692093, EU040228 

Devriesia thermodurans CBS 115878*; DAOM 225330 Soil Canada N. Nickerson AY692087, EU040229 

Hormoconis resinae CBS 365.86 – – – EU040230, EU040230 

CBS 184.54; ATCC 11841; CPC 3692; IMI 089837; IFO 31706 Creosote-treated wooden pole U.S.A. – AY251067, EU040231 

Hyalodendriella betulae CBS 261.82* Alnus glutinosa Netherlands W. Gams EU040232, EU040232 

Ochrocladosporium elatum CBS 146.33*; ATCC 11280; ATHUM 2862; IFO 6372; IMI 049629; Wood pulp Sweden E. Melin EU040233, EU040233 

MUCL 10094 

Ochrocladosporium frigidarii CBS 103.81* Cooled room Germany B. Ahlert EU040234, EU040234 

Parapleurotheciopsis inaequiseptata MUCL 41089; INIFAT C98/30-1 Rotten leaf Brazil R.F. Castañeda EU040235, EU040235 

Passalora daleae CBS 113031* Dalea spinosa Mexico L.B. Sparrius EU040236, EU040236 

Rachicladosporium luculiae CPC 11407* Luculia sp. New Zealand F. Hill EU040237, EU040237 

Ramularia aplospora Mycosphaerella alchemillicola CBS 545.82* Powdery mildew on Alchemilla vulgaris Germany T. Hijwegen EU040238, EU040238 

Retroconis fusiformis CBS 330.81; IMI 170799 Gossypium sp. Pakistan – EU040239, EU040239 

Rhizocladosporium argillaceum CBS 241.67*; ATCC 38103; IFO 7055; OUT 4262 Decayed myxomycete Japan K. Tubaki EU040240, EU040240 

Subramaniomyces fusisaprophyticus CBS 418.95; INIFAT C94/134 Leaf litter Cuba R.F. Castañeda EU040241, EU040241 

Thedgonia ligustrina W1877 Ligustrum sp. – H. Evans EU040242, EU040242 

Toxicocladosporium irritans CBS 185.58* Mouldy paint Suriname M.B. Schol-Schwarz EU040243, EU040243 

Verrucocladosporium dirinae CBS 112794* Dirina massiliensis U.K. A. Aptroot EU040244, EU040244 

1 ATCC: American Type Culture Collection, Virginia, U.S.A.; ATHUM: Culture Collection of Fungi, University of Athens, Department of Biology, Section of Ecology and Systematics, Athens, Greece; CBS: Centraalbureau voor Schimmelcultures, Utrecht, 

The Netherlands; CMW: Culture collection of Mike Wingfield, housed at FABI, Pretoria, South Africa; CPC: Culture collection of Pedro Crous, housed at CBS; DAOM: Plant Research Institute, Department of Agriculture (Mycology), Ottawa, Canada; IFO: 

Institute For Fermentation, Osaka, Japan; IMI: International Mycological Institute, CABI-Bioscience, Egham, Bakeham Lane, U.K.; INIFAT: Alexander Humboldt Institute for Basic Research in Tropical Agriculture, Ciudad de La Habana, Cuba; MUCL: 

Mycotheque de l’ Université Catholique de Louvain, Louvain-la-Neuve, Belgium; OUT: Department of Fermentation Technology, Faculty of Engineering, Osaka University, Yamadaue, Suita-shi, Osaka, Japan. 

2 ITS: internal transcribed spacer regions and 5.8S rRNA gene; LSU: partial 28S rRNA gene. 


34

Cladosporium and morphologically similar genera 

The current circumscription of Cladosporium emend. can be 

summarised as follows: Dematiaceous hyphomycetes; Davidiella 

anamorphs; mycelium internal and external; hyphae branched, 

septate, pigmented; stromata lacking or occasionally present; 

conidiophores mononematous, solitary to fasciculate, cylindrical, 

geniculate-sinuous to nodulose, simple to branched, subhyaline 

to usually distinctly pigmented, continuous to septate, smooth 

to verruculose; conidiogenous cells integrated, terminal and 

intercalary, usually sympodial, with a single to several scars; 

conidiogenesis holoblastic; conidiogenous loci coronate, i.e., 

more or less protuberant, composed of a central convex dome, 

surrounded by a raised periclinal rim, barely to distinctly darkened; 

conidia solitary or in short to long, simple to branched acropetal 

chains, amero- to phragmosporous, subhyaline to usually distinctly 

pigmented, smooth, verruculose, verrucose, echinulate, cristate, 

hila coronate, more or less protuberant. 

The new concept of Cladosporium s. str., supported by 

molecular data and typical coronate conidiogenous loci and conidial 

hila, rendered it possible to initiate a comprehensive revision 

of Cladosporium s. lat. The preparation of a general, annotated 

check-list of Cladosporium s. lat. was the first step in this direction 

(Dugan et al. 2004). The aim of the present study, therefore, was to 

delineate Cladosporium s. str. from other taxa that have in recent 

years been described in Cladosporium s. lat. To attain this goal 

isolates were studied under standardised conditions on a set of 

predescribed media (Schubert et al. 2007b – this volume), and 

subjected to DNA sequence analysis of the LSU nrRNA gene. 

were only sequenced for isolates of which these data were not 

available. The ITS data were not included in the analyses but 

deposited in GenBank where applicable. Gaps longer than 10 

bases were coded as single events for the phylogenetic analyses; 

the remaining gaps were treated as missing data. Sequence data 

were deposited in GenBank (Table 1) and alignments in TreeBASE 

(www.treebase.org). 

Morphology 

Wherever possible, 30 measurements (× 1 000 magnification) 

were made of structures mounted in lactic acid or Shear’s solution 

(Gams et al. 2007), with the extremes of spore measurements 

given in parentheses. Microscopic observations were made from 

colonies cultivated for 7 d under continuous near-ultraviolet light 

at 25 °C on SNA as explained in Schubert et al. (2007b – this 

volume). Three classes of conidia are distinguished. Ramoconidia 

are defined as short apical branches (often conidiogenous cells) 

of a conidiophore which secede and function as conidia. They are 

characterised by having a truncate, undifferentiated base, i.e., 

they differ from true conidia by lacking characteristic basal hila 

caused by conidiogenesis. Ramoconidia give rise to branched or 

unbranched conidia. Secondary ramoconidia are branched conidia 

with a narrowed base, bearing a true hilum, that can occur in chains, 

giving rise to conidia, which differ from secondary ramoconidia 

with regards to shape, size and septation. In previous literature on 

Cladosporium and allied genera, the true ramoconidia have often 

been classified as “ramoconidia s. str.” whereas the secondary 

ramoconidia have been named “ramoconidia s. lat.” 


Isolates 

Isolates used were obtained from the Centraalbureau voor 

Schimmelcultures (CBS), or freshly isolated from various 

substrates (Table 1). Strains were cultured on 2 % malt extract 

plates (MEA; Gams et al. 2007), by obtaining single conidial 

colonies as explained in Crous (1998). Colonies were subcultured 

onto fresh MEA, oatmeal agar (OA), potato-dextrose agar (PDA) 

and synthetic nutrient-poor agar (SNA) (Gams et al. 2007), and 

incubated under near-ultraviolet light to study their morphology. 

Cultural characteristics were assessed after 2–4 wk on OA and 

PDA at 25 °C in the dark, using the colour charts of Rayner (1970). 

Nomenclatural novelties and descriptions were deposited in 

MycoBank (www.MycoBank.org). 

DNA isolation, sequencing and phylogeny 


DNA was isolated following the CTAB-based protocol described in 

Gams et al. (2007). The primers V9G (de Hoog & Gerrits van den 

Ende 1998) and LR5 (Vilgalys & Hester 1990) were used to amplify 

part of the nuclear rDNA operon spanning the 3’ end of the 18S 

rRNA gene (SSU), the first internal transcribed spacer (ITS1), the 

5.8S rRNA gene, the second ITS region and the 5’ end of the 28S 

rRNA gene (LSU). Four internal primers, namely ITS4 (White et al. 

1990), LR0R (Rehner & Samuels 1994), LR3R (www.biology.duke. 

edu/fungi/mycolab/primers.htm), and LR16 (Moncalvo et al. 1993), 

were used for sequencing to ensure that good quality overlapping 

sequences are obtained. The PCR conditions, sequence alignment 

and subsequent phylogenetic analysis followed the methods of 

Crous et al. (2006d). The ITS1, ITS2 and 5.8S rRNA gene (ITS) 


Results 

DNA extraction, amplification and phylogeny 

Amplicons of approximately 1 700 bases were obtained for the 

isolates listed in Table 1. The newly generated sequences were 

used to obtain additional sequences from GenBank, which were 

added to the alignment. The manually adjusted LSU alignment 

contained 73 sequences (including the two outgroup sequences) 

and 996 characters including alignment gaps. Of the 849 characters 

used in the phylogenetic analysis, 336 were parsimony-informative, 

77 were variable and parsimony-uninformative, and 436 were 

constant. Neighbour-joining analyses using three substitution 

models on the sequence data yielded trees with identical topologies 

to one another. The neighbour-joining trees support the same 

clades as obtained from the parsimony analysis, but with a different 

arrangement at the deeper nodes, which were poorly supported 

in the bootstrap analyses or not at all (for example, the Helotiales 

and Pleosporales are swapped around). Performing a parsimony 

analysis with gaps treated as new characters increases the number 

of equally parsimonious trees to 94; the same topology is observed 

but with less resolution for the taxa in the Helotiales (data not 

shown). Forty-four equally most parsimonious trees (TL = 1 572 

steps; CI = 0.436; RI = 0.789; RC = 0.344), one of which is shown 

in Fig. 1, were obtained from the parsimony analysis of the LSU 

sequence data. The cladosporium-like taxa were found to belong to 

the Helotiales, Pleosporales, Sordariales and as sister taxa to the 

Davidiellaceae in the Capnodiales. 

The LSU alignment used for parsimony and distance analysis 

was supplemented with sequences for Parapleurotheciopsis 

inaequiseptata (Matsush.) P.M. Kirk and Subramaniomyces 

fusisaprophyticus (Matsush.) P.M. Kirk, as well as related sequences 

35

Crous et al. 

100 

61 

100 

52 

100 

10 changes 



Rhizocladosporium argillaceum CBS 241.67 

Myxotrichum deflexum AY541491 

Hormoconis resinae CBS 365.86 


56 

99 

99 

67 

100 84 

90 

93 

81 

100 

65 

75 

100 

Bisporella citrina AY789385 

Sarcoleotia turficola AY789277 

Torrendiella eucalypti DQ195799 


Hyalodendriella betulae CBS 261.82 

Thedgonia ligustrina W1877 

Blumeria graminis f. sp. bromi AB022362 Erysiphaceae 

Neofabraea alba AY064705 Dermateaceae 

Neofabraea malicorticis AY544662 

Phoma herbarum DQ678066 

Ascochyta pisi DQ678070 

Didymella bryoniae AB266850 

Didymella cucurbitacearum AY293792 

Leptospora rubella DQ195792 

Phaeosphaeria avenaria AY544684 

Coniothyrium palmarum CBS 758.73 

Ochrocladosporium frigidarii CBS 103.81 

Ochrocladosporium elatum CBS 146.33 

Cladoriella eucalypti DQ195790 

100 Cladoriella eucalypti CBS 115899 

100 

83 

87 

85 

69 

Helotiaceae 

Chaetomium homopilatum AF286404 

Retroconis fusiformis CBS 330.81 

Chaetomium globosum AF286403 

Aporothielavia leptoderma AF096186 

Rachicladosporium luculiae CPC 11407 

Dichocladosporium chlorocephalum EU009456 




Verrucocladosporium dirinae CBS 112794 

Toxicocladosporium irritans CBS 185.58 

97 

93 

89 

100 

Cladosporium cladosporioides DQ008145 

Cladosporium uredinicola DQ008147 

Cladosporium bruhnei DQ008149 

Cladosporium iridis DQ008148 

Cladosporium cladosporioides DQ008146 

Mycosphaerella marksii AF309578 

Penidiella nectandrae EU019275 

“Stenella” cerophilum AF050286 

84 

55 

100 

75 

94 

100 

99 

100 

94 

58 

52 

58 

Incertae sedis 

Myxotrichaceae 

Amorphotecaceae 



“Mycosphaerella” lateralis AF309583 

Pseudocercospora paraguayensis AF309574 

Passalora eucalypti AF309575 

Mycosphaerella irregulariramosa AF309576 


Ramularia aplospora CBS 545.82 

Passalora daleae CBS 113031 

Mycosphaerella africana AF309581 

Passalora fulva DQ008163 


Devriesia americana CBS 117726 

Penidiella strumelloidea EU019277 

Devriesia thermodurans CBS 115878 

64 

66 

Devriesia shelburniensis CBS 115876 

Devriesia acadiensis CBS 115874 



Catenulostroma germanicum EU019253 

Catenulostroma chromoblastomycosum EU019251 

Penidiella rigidophora EU019276 


Penidiella columbiana EU019274 

Penidiella venezuelensis EU019278 

Catenulostroma castellanii EU019250 

Teratosphaeria suttonii AF309587 

Teratosphaeria molleriana AF309584 

Teratosphaeria cryptica AF309585 

Teratosphaeria juvenis AF309586 

Phaeosphaeriaceae 

Leptosphaeriaceae 



Chaetomiaceae, Sordariales 



Pleosporales 

Fig. 1. One of 44 equally most parsimonious trees obtained from a heuristic search with 100 random taxon additions of the LSU sequence alignment using PAUP v. 4.0b10. The 

scale bar shows 10 changes, and bootstrap support values from 1 000 replicates are shown at the nodes. Thickened lines indicate the strict consensus branches and ex-type 

sequences are printed in bold face. The tree was rooted to two sequences obtained from GenBank (Athelia epiphylla AY586633 and Paullicorticium ansatum AY586693). 



Helotiales 

Capnodiales 

36




Blumeria graminis f. sp. bromi AB022362 Erysiphaceae 

Bisporella citrina AY789385 Helotiaceae 

0.80 

Hyalodendriella betulae CBS 261.82 

Thedgonia ligustrina W1877 


1.00 

0.69 Neofabraea alba AY064705 

Dermateaceae 

1.00 Neofabraea malicorticis AY544662 

Helotiales 

0.60 Sarcoleotia turficola AY789277 Helotiaceae 

1.00 Torrendiella eucalypti DQ195799 

0.90 



1.00 Hormoconis resinae CBS 365.86 

Amorphotecaceae 


0.94 

Rhizocladosporium argillaceum CBS 241.67 Incertae sedis 

Myxotrichum deflexum AY541491 Myxotrichaceae 

1.00 Chaetomium globosum AF286403 

0.88 

Aporothielavia leptoderma AF096186 

0.95 

Chaetomiaceae, Sordariales 

1.00 0.79 

Chaetomium homopilatum AF286404 

Retroconis fusiformis CBS 330.81 

1.00 Fasciatispora petrakii AY083828 Xylariaceae 

0.81 

Phlogicylindrium eucalypti DQ923534 Incertae sedis 

Plectosphaera eucalypti DQ923538 Phyllachoraceae 

0.72 

Xylariales 

0.74 Subramaniomyces fusisaprophyticus CBS 418.95 Incertae sedis 

0.68 Parapleurotheciopsis inaequiseptata MUCL 41089 Incertae sedis 

Pseudomassaria carolinensis DQ810233 Hyponectriaceae 

Phoma herbarum DQ678066 

0.63 1.00 Ascochyta pisi DQ678070 

1.00 0.97 


Didymella bryoniae AB266850 

1.00 Didymella cucurbitacearum AY293792 

Leptospora rubella DQ195792 

Pleosporales 

1.00 

Phaeosphaeriaceae 

Phaeosphaeria avenaria AY544684 

0.78 Coniothyrium palmarum CBS 758.73 Leptosphaeriaceae 

0.93 Ochrocladosporium frigidarii CBS 103.81 

1.00 Ochrocladosporium elatum CBS 146.33 


1.00 Cladoriella eucalypti CBS 115899 


1.00 Cladoriella eucalypti DQ195790 

Rachicladosporium luculiae CPC 11407 

0.85 Dichocladosporium chlorocephalum EU009456 


0.68 Dichocladosporium chlorocephalum EU009457 Incertae sedis 


0.89 Verrucocladosporium dirinae CBS 112794 

Toxicocladosporium irritans CBS 185.58 

0.90 Cladosporium cladosporioides DQ008145 

1.00 Cladosporium uredinicola DQ008147 

0.97 1.00 Cladosporium bruhnei DQ008149 


0.61 Cladosporium iridis DQ008148 

1.00 Cladosporium cladosporioides DQ008146 

Mycosphaerella marksii AF309578 

1.00 Penidiella nectandrae EU019275 

“Stenella” cerophilum AF050286 

0.71 “Mycosphaerella” lateralis AF309583 

1.00 Pseudocercospora paraguayensis AF309574 

0.99 Passalora eucalypti AF309575 

0.51 Mycosphaerella irregulariramosa AF309576 

1.00 Mycosphaerella punctiformis AY490776 

0.86 Ramularia aplospora CBS 545.82 

Passalora daleae CBS 113031 

1.00 Mycosphaerella africana AF309581 

1.00 Passalora fulva DQ008163 

0.94 1.00 Staninwardia suttonii DQ923535 

Devriesia americana CBS 117726 

Penidiella strumelloidea EU019277 

1.00 Devriesia thermodurans CBS 115878 

1.00 

Devriesia shelburniensis CBS 115876 

0.99 Devriesia acadiensis CBS 115874 

0.86 Devriesia staurophora DQ008150 


0.67 

Catenulostroma germanicum EU019253 

0.65 Penidiella venezuelensis EU019278 

Catenulostroma castellanii EU019250 

0.1 expected changes per site 0.73 Teratosphaeria suttonii AF309587 

1.00 Teratosphaeria molleriana AF309584 

1.00 Teratosphaeria cryptica AF309585 

Teratosphaeria juvenis AF309586 

1.00 Catenulostroma chromoblastomycosum EU019251 

0.64 

1.00 

Penidiella rigidophora EU019276 


Penidiella columbiana EU019274 



Capnodiales 

Fig. 2. Consensus phylogram (50 % majority rule) of 800 trees resulting from a Bayesian analysis of the LSU sequence alignment using MrBayes v. 3.1.2. Bayesian posterior 

probabilities are indicated at the nodes. Ex-type sequences are printed in bold face. The tree was rooted to two sequences obtained from GenBank (Athelia epiphylla AY586633 

and Paullicorticium ansatum AY586693). 


37

Crous et al. 

Fig. 3. Rachicladosporium luculiae (type material). A–F. Conidiophores with conidial chains, and conidiogenous loci aggregated in the upper region. G. Conidia. Scale bar = 

10 µm. 

from GenBank. This alignment was subjected to a Bayesian 

analysis using a general time-reversible (GTR) substitution model 

with inverse gamma rates and dirichlet base frequencies and the 

temp value set to 0.5. The Markov Chain Monte Carlo (MCMC) 

analysis of 4 chains started from a random tree topology and lasted 

1 000 000 generations. Trees were saved each 1 000 generations, 

resulting in 1 000 trees. Burn-in was set at 200 000 generations 

after which the likelihood values were stationary, leaving 800 trees 

from which the consensus tree (Fig. 2) and posterior probabilities 

(PP’s) were calculated. The average standard deviation of split 

frequencies was 0.018459 at the end of the run. The same overall 

topology as that observed using parsimony was obtained, with 

the main exception that the Helotiales and Pleosporales swapped 

around, as observed with the distance analysis. 

Taxonomy 

The present study has delineated several cladosporium-like genera 

which are phylogenetically unrelated to, and morphologically distinct 

from Cladosporium s. str. (Davidiellaceae, Capnodiales). These are 

treated below: 

Capnodiales, incertae sedis 

Rachicladosporium Crous, U. Braun & C.F. Hill, gen. nov. 


Etymology: Named after the apical rachis on conidiophores, and its 

cladosporium-like appearance. 

Differt a Cladosporio conidiophoris cum rachibus terminalibus, locis conidiogenis 

inconspicuis vel subconspicuis, margine leviter incrassatis, non fuscatis et non 

refractivis, hilis inconspicuis. 

Mycelium consisting of branched, septate, smooth, hyaline to 

pale brown, thin-walled hyphae. Conidiophores erect, solitary, 

macronematous, arising from superficial hyphae, subcylindrical, 

straight to somewhat geniculate-sinuous, medium brown, 

finely verruculose; basal foot cell without swelling or rhizoids. 

Conidiogenous cells integrated, terminal, subcylindrical or tips 

slightly swollen, forming an apical rachis, multilocal, loci terminal 

and lateral, without evident sympodial proliferation (non-geniculate); 

conidiogenous loci inconspicuous or subconspicuous by being very 

slightly thickened along the rim, but neither darkened nor refractive, 

giving rise to simple or branched chains or solitary conidia. 

Ramoconidia medium brown, finely verruculose, 0–1-septate, 

subcylindrical to narrowly ellipsoid; conidia ellipsoid, pale brown, 

0(–1)-septate, smooth to finely verruculose; hila inconspicuous; 

secession schizolytic. 

Type species: Rachicladosporium luculiae Crous, U. Braun & C.F. 

Hill, sp. nov. 

38


Rachicladosporium luculiae Crous, U. Braun & C.F. Hill, sp. nov. 


Etymology: Named after its host genus, Luculia. 

Mycelium ex hyphis ramosis, septatis, levibus, hyalinis vel pallide brunneis, 2–3 

µm latis compositum. Conidiophora erecta, solitaria, macronemata, ex hyphis 

superficialibis oriunda, subcylindrica, recta to geniculata-sinuosa, ad 60 µm longa et 

6 µm lata, 3–6-septata, modice brunnea, subtiliter verruculosa, non crassitunicata, ad 

basim non inflatae et non rhizoideae. Cellulae conidiogenae integratae, terminales, 

8–15 × 4–5 µm, subcylindricae, apicem versus attenuatae, apice obtuso, rachidi 

terminali, locis conidialibus numerosis, 1–2 µm latis, margine leviter incrassatis, 

non fuscatis et non refractivis. Conidia catenata vel solitaria. Ramoconidia modice 

brunnea, subtile verruculosa, 0–1-septata, subcylindrica vel anguste ellipsoidea, 

10–17 × 4–5 µm; conidia secundaria ellipsoidea, pallide brunnea, 0(–1)-septata, 

levia vel subtile verruculosa, interdum guttulata, (7–)9–12(–15) × 3(–4) µm; hila 

inconspicua. 

Mycelium consisting of branched, septate, smooth, thin-walled, 

hyaline to pale brown, 2–3 µm wide hyphae. Conidiophores 

erect, solitary, macronematous, arising from superficial hyphae, 

subcylindrical, straight to somewhat geniculate-sinuous, up to 60 µm 

long, and 6 µm wide, 3–6-septate, medium brown, finely verruculose, 

thin-walled (≤ 1 µm), rarely with a single percurrent proliferation; 

basal foot cell without swelling or rhizoids. Conidiogenous cells 

integrated, terminal, 8–15 × 4–5 µm, subcylindrical, tapering to an 

obtuse apex, occasionally slightly swollen at the tip, without distinct 

sympodial proliferation (non-geniculate), forming a rachis, with 

several conidiogenous loci, terminal and lateral, 1–2 µm wide, nonprotuberant, 

quite inconspicuous to subconspicuous, very slightly 

thickened along the rim, but not darkened and refractive; giving rise 

to simple or branched chains or solitary conidia, thin-walled (≤ 0.75 

µm). Ramoconidia medium brown, finely verruculose, 0–1-septate, 

subcylindrical to narrowly ellipsoid, 10–17 × 4–5 µm; conidia 

ellipsoid, pale brown, 0(–1)-septate, smooth to finely verruculose, 

at times guttulate, (7–)9–12(–15) × 3(–4) µm; hila inconspicuous, 

neither thickened nor darkened-refractive. 


with moderate aerial mycelium and smooth, even margins; irongrey 

in the centre, olivaceous-grey in the outer region (surface); 

iron-grey underneath. Colonies reaching 4 cm diam after 1 mo at 

25 °C in the dark. 

Specimen examined: New Zealand, Auckland, isolated from leaf spots on Luculia 

sp. (Rubiaceae), 25 Jul. 2004, F. Hill 1059, holotype CBS H-19891, culture ex-type 

CBS 121620 = CPC 11407. 

Notes: Rachicladosporium is morphologically quite distinct from 

Cladosporium s. str. and allied cladosporioid genera by having an 

apical conidiophore rachis with inconspicuous to subconspicuous 

scars and unthickened, not darkened-refractive conidial hila. Due 

to the structure of the conidiogenous cells, R. luculiae superficially 

resembles species of the tretic genus Diplococcium Grove (Ellis 

1971, 1976; Goh & Hyde 1998). However, there is no evidence 

for a tretic conidiogenesis in R. luculiae. The conidia are formed 

holoblastically and separated by a thin septum. Furthermore, 

in Diplococcium the conidiogenous cells are terminal as well as 

intercalary, the conidiophores are often branched, and branched 

conidial chains are lacking or at least less common. Molecular 

sequence data about Diplococcium species are not yet available, 

though taxa that have been analysed show affinities to the 

Pleosporaceae and Helotiales (Wang et al., unpubl. data), whereas 

Rachicladosporium is allied with the Capnodiales. The ecology 

of R. luculiae is still unclear, although it has been isolated from 

lesions on Luculia sp. Fruiting of this species in vivo has not yet 

been observed, and its pathogenicity remains unproven. 


Toxicocladosporium Crous & U. Braun, gen. nov. MycoBank 

MB504426. 

Etymology: Named after ample volatile metabolites produced in 

culture, and cladosporium-like morphology. 

Differt a Cladosporio locis conidiogenis denticulatis, incrassatis et fuscatis-refractivis, 

sed non coronatis, conidiophoris et conidiis cum septis incrassatis et atrofuscis, et 

culturis cum metabolitis volaticis toxicis. 

Mycelium consisting of branched, septate, dark brown, finely 

verruculose hyphae. Conidiophores solitary, dimorphic, solitary, 

macronematous or micronematous, reduced to conidiogenous 

cells. Macronematous conidiophores subcylindrical, straight 

to geniculate-sinuous, or irregularly curved, unbranched or 

branched above, septate, dark brown, finely verruculose, walls 

thick, septa dark brown; micronematous conidiophores reduced 

to conidiogenous cells, erect, doliiform to subcylindrical, with slight 

taper towards the apex. Conidiogenous cells integrated, terminal or 

lateral, subcylindrical with slight taper towards apex; proliferating 

sympodially with apical loci protruding and denticle-like, thickened, 

darkened and refractive, but not coronate. Conidia catenulate in 

branched or unbranched chains, medium to dark brown, thickwalled, 

with dark, thick septa, smooth to finely verruculose; 

ramoconidia septate, prominently constricted at septa, broadly 

ellipsoid to subcylindrical; conidia ellipsoid to ovoid, pale to medium 

brown, 0(–1)-septate; hila not coronate, but protruding, thickened, 

darkened and refractive in ramoconidia, but less obvious in young 

conidia. 

Type species: Toxicocladosporium irritans Crous & U. Braun, sp. 

nov. 

Toxicocladosporium irritans Crous & U. Braun, sp. nov. 


Etymology: Named after the skin irritation resulting from exposure 

to the fungus. 

Mycelium (in PDA) ex hyphis ramosis, septatis, atro-brunneis, minute verruculosis, 

(2–)3–4 µm latis, ultimo crassitunicatis et crassiseptatis. Conidiophora solitaria, 

dimorphosa, macronemata et solitaria vel micronemata. Conidiophora macronemata 

ex hyphis modice brunneis lateraliter oriunda, erecta, subcylindrica, recta, 

geniculata-sinuosa vel irregulariter curvata, non ramosa vel ad apicem ramosa, 

2–7-septata, atro-brunnea, leviter verruculosa, crassitunicata, septa atro-brunnea, 

30–60 × 4–6 µm; conidiophora micronemata saepe non septata, raro 1–2-septata, 

erecta, doliiformes vel subcylindrica, apicem versus leviter attenuata, 10–30 × 2.5– 

4 µm. Cellulae conidiogenae integratae, terminales vel laterales, subcylindricae, 

apicem versus leviter attenuatae, 7–12 × 3–4 µm, sympodiales, cum 1–3 locis 

conidiogenibus, denticulatis, 1–1.5 µm latis, incrassatis, fuscatis-refractivis. Conidia 

catenulata vel rami-catenulata, modice vel atro-brunnea, crassitunicata, septis 

incrassatis, fuscatis, levia vel subtile verruculosa; ramoconidia (0–)1(–3)-septata, 

constricta, late ellipsoidea vel subcylindrica, 7–15 × 3–5 µm; conidia secundaria 

ellipsoidea vel ovoidea, pallide vel modice brunnea, 0(–1)-septata, (5–)6–8(–10) × 

(3–)4(–5) µm; hila protuberantes, 1–1.5 µm lata, hila ramoconidiorum incrassata 

et fuscata-refractiva, vel hila conidiorum secundariorum 0.5–1 µm lata et 

subconspicua. 

Mycelium on PDA consisting of branched, septate, dark brown, finely 

verruculose, (2–)3–4 µm wide hyphae; walls and septa becoming 

thickened and darkened with age. Conidiophores solitary, dimorphic, 

macronematous and solitary, or micronematous, reduced to 

conidiogenous cells. Macronematous conidiophores subcylindrical, 

straight to geniculate-sinuous, or irregularly curved, unbranched 

or branched above, 2–7-septate, dark brown, finely verruculose, 

walls thick, septa dark brown, 30–60 × 4–6 µm; medium brown 

hyphae giving rise to lateral, erect branches that become swollen, 

dark brown, and develop into macronematous conidiophores 

with thick-walled and dark septa; micronematous conidiophores 

39

Crous et al. 

Fig. 4. Toxicocladosporium irritans (type material). A–B, F. Microconidiophores. C–E. Macroconidiophores. G–H. Ramoconidia and conidia. Scale bars = 10 µm. 

aseptate, reduced to conidiogenous cells (rarely 1–2-septate, i.e., 

with 1–2 supporting cells), erect, doliiform to subcylindrical, with 

slight taper towards the apex, 10–30 × 2.5–4 µm. Conidiogenous 

cells integrated, terminal or lateral, subcylindrical with slight taper 

towards apex, 7–12 × 3–4 µm; proliferating sympodially with 1–3 

apical loci that can be slightly protruding and denticle-like, 1–1.5 

µm wide, thickened, darkened and refractive. Conidia catenulate 

in branched or unbranched chains, medium to dark brown, thickwalled, 

with dark, thick septa, smooth to finely verruculose; 

ramoconidia (0–)1(–3)-septate, prominently constricted at septa, 

broadly ellipsoid to subcylindrical, 7–15 × 3–5 µm; conidia ellipsoid 

to ovoid, younger apical conidia pale to medium brown, 0(–1)- 

septate, (5–)6–8(–10) × (3–)4(–5) µm; hila protruding, 1–1.5 µm 

wide, thickened, darkened and refractive in ramoconidia, but less 

obvious in young conidia, where hila are 0.5–1 µm wide. 


with dense aerial mycelium and smooth, even margins; surface 

olivaceous-black (centre), olivaceous-grey in outer region; reverse 

olivaceous-black. Colonies reaching 35 mm diam after 1 mo at 25 

°C in the dark; colonies fertile. 

40


Specimen examined: Suriname, Paramaribo, isolated from mouldy paint, Feb. 

1958, M.B. Schol-Schwarz, holotype CBS-H 19892, culture ex-type CBS 185.58. 

Notes: Toxicocladosporium irritans produces ample amounts 

of volatile metabolites, which cause a skin rash within minutes 

of opening an inoculated dish for microscopic examination. 

Morphologically and phylogenetically it is very similar to 

Cladosporium s. str., and produces dimorphic conidiophores, which 

is also commonly observed in Cladosporium. It is distinct by having 

dark, thick-walled conidial and conidiophore septa, and lacking the 

typical coronate Cladosporium scar type (David 1997). 

Verrucocladosporium K. Schub., Aptroot & Crous, gen. nov. 


Etymology: Named after its frequently coarsely verrucose to 

warted hyphae, conidiophores and conidia, and cladosporium-like 

morphology. 

Differt a Cladosporio hyphis saepe verrucosis, hyalinis, conidiophoris cylindraceisfiliformibus, 

rectis, non vel vix geniculatis, non nodulosis, locis conidiogenis leviter 

incrassatis, distincte fuscatis-refractivis, sed non coronatis, conidiis saepe valde 

variantibus, saepe irregulariter formatis, grosse verrucosis-rugosis. 

Mycelium sparingly branched, hyphae septate, not constricted at 

septa, hyaline, almost smooth to irregularly rough-walled, coarsely 

verrucose to warted. Conidiophores arising laterally from creeping 

hyphae, erect, straight, or somewhat flexuous, narrowly cylindrical 

to filiform, neither geniculate nor nodulose, unbranched, septate, 

pale brown, thin-walled, smooth to often irregularly rough-walled or 

verrucose. Conidiogenous cells integrated, terminal or intercalary, 

cylindrical, polyblastic, with sympodial proliferation, with loci often 

crowded at the apex, truncate, barely to slightly thickened, but 

distinctly darkened-refractive. Ramoconidia cylindrical, aseptate, 

concolorous with conidiophores, thin-walled, irregularly roughwalled, 

coarsely verruculose to verrucose-rugose; hila unthickened 

but somewhat refractive. Conidia in long unbranched or loosely 

branched chains, obovoid, ellipsoid, fusiform to subcylindrical, with 

swollen and constricted parts, often appearing irregular in shape 

and outline, 0–1-septate, pale brown, thin-walled and irregularly 

rough-walled, verruculose-rugose; hila truncate, barely to slightly 

thickened, but distinctly darkened-refractive. 

Type species: Verrucocladosporium dirinae K. Schub., Aptroot & 

Crous, sp. nov. 

Verrucocladosporium dirinae K. Schub., Aptroot & Crous, sp. 


Etymology: Named after its host, Dirina massiliensis. 

Mycelium sparse ramosum. Hyphae 1–3 µm latae, septatae, non constrictae, 

hyalinae, leviae, vel irregulariter verruculosae, interdum verrucosae, tuberculatae, 

tenuitunicatae. Conidiophora ex hyphis repentibus lateraliter oriunda, erecta, 

recta, interdum leviter flexuosa, anguste cylindrica vel filiformes, non geniculta, 

non nodulosa, non ramosa, ad 85 µm longa, 2–3 µm lata, septata, tenuitunicata 

(≤ 0.75 µm), pallide brunnea, levia vel saepe irregulariter verrucosa, leviter 

crassitunicata. Cellulae conidiogenae integratae, saepe terminales, interdum 

intercalares, cylindricae, angustae, 9–20 µm longae, holoblasticae, sympodiales, 

locis conidiogenibus 1–3, saepe ad apicem aggregatis, interdum protuberantibus, 

truncatis, 1–1.8(–2) µm latis, incrassatis et fuscatis-refractivis. Ramoconidia 

cylindrica, 16–21 × (2–)2.5–3 µm, non septata, pallide brunnea, tenuitunicata, 

irregulariter verruculosa vel crosse verrucosa-rugosa, ad 4 hilis terminalibus, 

ad basim late truncata, non attenuata, 2–2.5 µm lata, non incrassata, sed leviter 

refractiva. Conidia catenata, in catenis longis, non ramosis vel laxe ramosis, plus 

minusve recta, obovoidea, ellipsoidea, fusiformes vel subcylindricae, sed saepe 

irregulares, 4–18(–23) × (2–)2.5–3.5 µm, 0–1-septata, ad septa interdum constricta, 

pallide brunnea, tenuitunicata (≤ 0.5 µm), irregulariter verruculosa-rugosa, utrinque 

leviter attenuata, hila truncata, (0.5–)0.8–1.5(–2) µm lata, vix vel leniter incrassata, 

sed distincte fuscata-refractiva. 

Mycelium sparingly branched; hyphae 1–3 µm wide, septate, not 

constricted at septa, hyaline, smooth to irregularly rough-walled, 

sometimes coarsely verrucose, with small to large drop-like, 

tuberculate warts, walls unthickened. Conidiophores arising laterally 

from creeping hyphae, erect, straight, sometimes slightly flexuous, 

narrowly cylindrical to filiform, not geniculate, non nodulose, 

unbranched, up to 85 µm long, 2–3 µm wide, septate, thin-walled 

(≤ 0.75 µm), pale brown, smooth to often irregularly rough-walled, 

verrucose, walls slightly thickened. Conidiogenous cells integrated, 

mostly terminal, sometimes also intercalary, cylindrical, narrow, 

9–20 µm long, conidiogenesis holoblastic, proliferation sympodial, 

with a single or up to three conidiogenous loci, often crowded at 

the apex, sometimes situated on small lateral prolongations, loci 

truncate, 1–1.8(–2) µm wide, thickened and darkened-refractive. 

Ramoconidia cylindrical, 16–21 × (2–)2.5–3 µm, aseptate, 

concolorous with conidiophores, thin-walled, irregularly roughwalled, 

verruculose to coarsely verrucose-rugose, apically with up 

to 4 hila, with a broadly truncate, non-attenuated base, 2–2.5 µm 

wide, unthickened but somewhat refractive. Conidia catenate, in 

long unbranched or loosely branched chains, more or less straight, 

obovoid, ellipsoid, fusiform to subcylindrical, but often appearing 

to form band-like structures, with swollen and constricted parts, 

accordion or fir tree-like and also due to ornamentation often 

appearing irregular in shape and outline, 4–18(–23) × (2–)2.5– 

3.5 µm, 0–1-septate, sometimes constricted at the more or less 

median septum, pale brown, thin-walled (≤ 0.5 µm), irregularly 

rough-walled, verruculose-rugose, somewhat attenuated towards 

apex and base, hila truncate, (0.5–)0.8–1.5(–2) µm wide, barely 

or slightly thickened, but distinctly darkened-refractive; microcyclic 

conidiogenesis not observed. 

Cultural characteristics: Colonies erumpent, spreading, with 

catenate, feathery margins and moderate aerial mycelium on PDA. 

Surface grey-olivaceous, reverse iron-grey. Colonies reaching 25 

mm after 1 mo at 25 °C. 

Specimen examined: U.K., Somerset, Kingsbury Episcopi, isolated from the lichen 

Dirina massiliensis (Roccelaceae, Arthoniales), Mar. 2003, A. Aptroot, holotype 

CBS-H 19883, culture ex-type CBS 112794. 

Notes: Verrucocladosporium dirinae was deposited as 

Cladosporium arthoniae M. Christ. & D. Hawksw., but the name 

was misapplied. The latter species, described from apothecia of 

Arthonia impolita on Quercus from Sweden, does not possess 

clearly visible, distinct conidiogenous loci and hila, and therefore 

has to be excluded from the genus Cladosporium s. str. and is also 

easily distinguishable from the newly introduced species above. 

Furthermore the conidiophores are apically frequently branched and 

the catenate, ellipsoid conidia are smaller and wider, 6–10 × 4–5 µm 

(Hawksworth 1979). Due to the conidiogenesis and the structure of 

the conidiogenous loci and conidia, C. arthoniae is rather close to 

lichenicolous Taeniolella S. Hughes species. The unique feature 

of the new genus Verrucocladosporium is its unusual conidial and 

hyphal ornamentation. Furthermore, it differs from Cladosporium 

s. str. in having cylindrical-filiform conidiophores, which are neither 

geniculate nor nodulose, quite distinct, thickened and darkened, 

but non-coronate conidiogenous loci and often irregularly shaped 

conidia. Phylogenetially, it is also distinct as a sister taxon to 

Cladosporium s. str. Concerning differences to other cladosporioid 

genera, see “key to the genera”. Verrucocladosporium dirinae has 

been isolated from the lichen species Dirina massiliensis, i.e., this 

species is probably lichenicolous, although its ecology is not quite 

clear. Fruiting of this species in vivo has not yet been observed. A 

second unnamed, taeniolella-like, lichenicolous hyphomycete was 

also present on the thallus of this lichen. 


41

Crous et al. 

Fig. 5. Verrucocladosporium dirinae (type material). A. Colonies on MEA. B–C. Conidial chains. D–H. Ramoconidia and conidia. Scale bars = 10 µm. 

Capnodiales, Teratosphaeriaceae 

Devriesia americana Crous & Dugan, sp. nov. MycoBank 

MB504434. Fig. 6. 

Etymology: Named after the geographic location of its type strain, 

New York, U.S.A. 

Differt a D. shelburniensi conidiophoris brevioribus (ad 30 µm longis), leviter 

latioribus (2–3 µm), ramoconidiis saepe nullis et conidiis 0–1-septatis. 

Mycelium consisting of branched, septate, 1.5–3 µm wide hyphae, 

irregular in width, predominantly guttulate, smooth, forming hyphal 

strands and hyphal coils; hyphae frequently forming dark brown, 

thick-walled, intercalary, muriformly septate chlamydospores on 

PDA in culture. Conidiophores subcylindrical, medium brown, 

straight to irregularly curved, up to 7-septate and 30 µm tall, 2–3 

µm wide, or reduced to conidiogenous cells. Conidiogenous cells 

terminal or lateral on hyphae, 5–12 × 2–3 µm, medium brown, 

smooth, guttulate, subcylindrical, mono- to polyblastic; scars 

somewhat darkened and thickened, but not refractive. Conidia 

medium brown, guttulate, smooth, in mostly unbranched chains, 

subcylindrical to narrowly ellipsoidal, tapering towards subtruncate 

ends, 0–1-septate, (7–)8–12(–16) × 2(–2.5) µm; hila darkened, 

somewhat thickened, not refractive, 1–1.5 µm wide. 

42


Fig. 6. Devriesia americana (type material). A–B. Chlamydospore-like structures formed in culture. C–F. Conidiophores giving rise to conidial chains. G–H. Conidia. Scale bars 

= 10 µm 

Cultural characteristics: Colonies erumpent, with sparse aerial 

mycelium on PDA, and smooth, uneven, wide margins, submerged 

under the agar surface; greenish-black (surface); reverse 

olivaceous-black; on OA iron-grey (surface). Colonies reaching 

8–15 mm diam on PDA after 14 d at 25 °C in the dark; colonies 

fertile, but sporulation sparse. 

Specimen examined: U.S.A., New York, Long Island, isolated from air, F.M. Dugan, 

holotype CBS-H 19894, culture ex-type CBS 117726 = ATCC 96545 = CPC 5121. 

Notes: Until recently, this species was treated as part of the 

“Phaeoramularia” hachijoensis species complex (Braun et al. 

2003). The present strain has conidia that are smaller than those 

of “Phaeoramularia” hachijoensis, which has ramoconidia that are 

1–3-septate, up to 30 µm long, and conidia that are predominantly 

1-septate, 10–21 × 2–4 µm (Matsushima 1975). From the illustration 

provided by Matsushima, it appears that “Phaeoramularia” 

hachijoensis is indeed a species of Pseudocladosporium U. Braun, 

a finding which is in agreement with the name Pseudocladosporium 

hachijoense (Matsush.) U. Braun proposed by Braun (1998). 

Devriesia americana is both morphologically and phylogenetically 

more allied to Teratosphaeria Syd. & P. Syd. than Venturia 

Sacc. Based on its pigmented conidiophores and catenulate 

conidia, and scars that are somewhat darkened and thickened, 

and the formation of chlamydospores in culture, it is allocated to 

Devriesia Seifert & N.L. Nick. Species of the genus Devriesia are 

ecologically different, however (Seifert et al. 2004), being soil-borne 


and thermotolerant. It is possible, therefore, that further collections 

of this fungus may eventually indicate that it needs to be placed in a 

distinct genus within the Teratosphaeriaceae. Devriesia americana 

is the second species of Devriesia with muriform chlamydospores, 

beside D. shelburniensis N.L. Nick. & Seifert, but the latter species 

is easily distinguishable by its long and narrow conidiophores (ca 

100–200 × 1.5–2.5 µm) and abundant ramoconidia, up to 25.5 

µm long, with 0–3 septa. Furthermore, D. shelburniensis is a 

thermotolerant soil-borne hyphomycete. 

Stenella araguata Syd., Ann. Mycol. 28(1/2): 205. 1930. Figs 7–8. 

≡ Cladosporium araguatum (Syd.) Arx, Genera of Fungi Sporulating in 

pure Culture, Edn 2 (Vaduz): 224. 1974. 

= Cladosporium castellanii Borelli & Marcano, Castellania 1: 154. 1973. 

Leaf spots hypophyllous, irregular to subcircular, up to 8 mm 

diam, indistinct, yellow to pale brown with indistinct margins on 

IMI 15728(a); on IMI 34905 (Fig. 7) lesions are amphigenous, and 

fascicles and sporodochia are rare, with superficial mycelium being 

predominant. Mycelium consisting of internal and external, medium 

brown, septate, branched, verruculose, 3–4 µm wide hyphae. 

Caespituli fasciculate to sporodochial, hypophyllous, medium brown, 

up to 120 µm wide and 60 µm high. Conidiophores arising singly 

from superficial mycelium, or aggregated in loose to dense fascicles 

arising from the upper cells of a brown stroma up to 70 µm wide and 

30 µm high; conidiophores medium brown, finely verruculose, 1–5- 

septate, subcylindrical, straight to geniculate-sinuous, unbranched 

43

Crous et al. 

Fig. 7. Stenella araguata (syntype material, IMI 34905). A. Leaf spot. B. Conidiophore, conidia and verruculose hypha on leaf surface. C–D. Conidiophore with terminal 

conidiogenous cells. E–G. Ramoconidia and conidia. Scale bars = 10 µm. 

or branched, 20–40 × 3–4 µm. Conidiogenous cells terminal or 

lateral, unbranched, medium brown, finely verruculose, tapering 

to slightly or flat-tipped loci, proliferating sympodially, 5–20 × 3–4 

µm; scars thickened, darkened and refractive. Conidia solitary 

or catenulate, in simple chains, medium brown, verruculose, 

subcylindrical to narrowly obclavate, apex obtuse, base bluntly 

rounded with truncate hilum, straight, 0–3-septate, (7–)13–20(–25) 

× 3(–3.5) µm; hila thickened, darkened, refractive, 1–1.5 µm wide. 

Description based on CBS 105.75 (Fig. 8): Mycelium consisting 

of branched, septate, verruculose, medium brown, 2–4 µm wide 

hyphae. Conidiomata brown, superficial, sporodochial, up to 200 µm 

diam Conidiophores solitary, erect, micro- to macronematous, 1–12- 

septate, subcylindrical, straight to geniculate-sinuous or irregularly 

curved, 10–70 × 3–4 µm; frequently swollen and constricted at 

septa, thick-walled, medium brown, verruculose. Conidiogenous 

cells terminal and intercalary, subcylindrical, straight, but frequently 

branched laterally, 6–20 × 3–4 µm, with 1–3 flat-tipped loci that 

can be subdenticulate, 1.5–2 µm wide, somewhat darkened and 

thickened, not prominently refractive. Conidia medium brown, thickwalled, 

verruculose, septa becoming darkly pigmented, occurring in 

branched chains. Ramoconidia subcylindrical to narrowly ellipsoid, 

12–25 × 3.5–4(–5) µm, 1(–4)-septate. Conidia occurring in short 

chains (–8), subcylindrical to narrowly ellipsoid, 0–1(–3)-septate, 

(7–)10–15(–20) × (2–)3–3.5(–4) µm; hila somewhat thickened, 

darkened but not refractive, 1.5–2(–2.5) µm wide. 

Cultural characteristics: Colonies on OA erumpent, spreading, with 

moderate aerial mycelium and smooth, even margins; olivaceousgrey 

(surface); on PDA olivaceous-black (surface), margins 

feathery, uneven, with moderate aerial mycelium; reverse iron-grey. 

Colonies reaching 20 mm diam after 1 mo at 25°C in the dark on 

OA. 

Specimens examined: Venezuela, Aragua, La Victoria, on leaf spots of 

Pithecellobium lanceolatum (Mimosaceae), Jan. 1928, H. Sydow, lectotype of S. 

araguata (selected here!) IMI 15728(a); 3 Feb. 1928, syntype of S. araguata, IMI 

34905. Venezuela, isolated from man with tinea nigra, 1973, D. Borelli, holotype of 

C. castellanii, IMI 183818, culture ex-type CBS 105.75. 

Notes: Stenella araguata is a leaf spot pathogen of Pithecellobium 

in Venezuela, and represents the type species of the genus Stenella 

[Two collections were cited, viz. no. 407, ‘La Victoria’, and no. 370, 

‘inter La Victoria et Suata’, both without any date, and without 

any specific type indication. Thus, the two collections have to be 

considered syntypes. The two IMI collections with different dates 

are parts of the syntypes, of which IMI 15728(a) is proposed here 

to serve as lectotype]. Stenella araguata was incorrectly seen as 

a species of Cladosporium by von Arx (1974), which has recently 

been morphologically circumscribed (Braun et al. 2003, Schubert et 

al. 2007b – this volume), and is linked to Davidiella teleomorphs. 

In a study by McGinnis & Padhye (1978), Cladosporium 

castellanii (tinea nigra of human in Venezuela) was shown to be 

synonymous to Stenella araguata (leaf spots of Pithecellobium 

lanceolatum in Venezuela). In the present study we re-examined 

the ex-type strain of C. castellanii (CBS 105.75), and found conidia 

to be 0–1(–3)-septate, (7–)10–15(–20) × (2–)3–3.5(–4) µm, while 

those of the type specimen of S. araguata were similar, namely 

0–3-septate, (7–)13–20(–25) × 3(–3.5) µm. Furthermore, both 

collections have verruculose hyphae, which is the primary feature 

distinguishing Stenella from Passalora Fr. (Crous & Braun 2003). 

44


Fig. 8. Stenella araguata (CBS 105.75). A–B. Conidiophore fascicles on a pine needle and tap-water agar, respectively. C–D, G. Conidiophores giving rise to conidial chains. 

E–F, H–J. Conidial chains with ramoconidia and conidia. Scale bars = 10 µm. 

Stenella has always been used for anamorphs of Mycosphaerella 

(Crous et al. 2004, 2006c), and the fact that it belongs to 

Teratosphaeria (Teratosphaeriaceae), and not Mycosphaerella 

(Mycosphaerellaceae), raised the question of how to treat stenellalike 

anamorphs in Mycosphaerella. Due to insufficient availability of 

cultures (Crous et al. 2000, 2001), the status of Stenella was left 

unresolved (Crous & Braun 2003). Presently (Crous & Groenewald, 

unpubl. data), it is clear that the stenella-like morphology type is 

polyphyletic within the Mycosphaerellaceae, and paraphyletic 

within the Capnodiales. Several species are known that represent 

morphological transitions between Stenella and Passalora. It 

seems logical, therefore, that future studies should favour using 

Passalora to also accommodate Mycosphaerella anamorphs with 

superficial, verruculose hyphae, which have traditionally been 

placed in Stenella. This is in spite of the fact that there are other 

generic names available within the Mycosphaerellaceae for taxa 

with a stenella-like morphology (pigmented structures, darkened, 

thickened, refractive scars, and superficial, verruculose mycelium), 

namely Zasmidium Fr. (1849) (see Arzanlou et al. 2007 – this 

volume), and Verrucisporota D.E. Shaw & Alcorn (1993). Based 

on the phylogenetic position of the type species, Stenella s. str. 

is an anamorph of Teratosphaeria (Teratosphaeriaceae). Using 

the generic concept as employed in this volume of the Studies in 

Mycology, however, the anamorph genus is accepted as being 

poly- and paraphyletic within the order Capnodiales. 


45

Crous et al. 

Helotiales, incertae sedis 

Hyalodendriella Crous, gen. nov. MycoBank MB504435. 

Etymology: Morphologically similar to Hyalodendron Diddens. 

Differt a Hyalodendro et Retroconi conidiophoris dimorphis, cicatricibus incrassatis 

et conidiis ultimo brunneis. 

Morphologically similar to Hyalodendron and Retroconis, but 

distinct in that it has dimorphic conidiophores, conidia that turn 

brown with age, and have thickened scars. Microconidiophores 

forming as lateral branches on hyphae, subcylindrical, subhyaline 

to pale brown, smooth, septate, with terminal conidiogenous cells. 

Macroconidiophores septate, subcylindrical, straight to curved, 

subhyaline to pale brown, smooth, with an apical rachis that is pale 

brown, smooth, subcylindrical, with numerous, aggregated loci. 

Conidia limoniform to ellipsoid, aseptate, smooth, pale brown, in 

short chains, tapering towards ends that are prominently apiculate, 

prominently thickened and darkened, but not refractive. 

Type species: Hyalodendriella betulae Crous, sp. nov. 

Hyalodendriella betulae Crous sp. nov. MycoBank MB504436. 

Fig. 9. 

Mycelium ex hyphis ramosis, septatis, 1.5–2 µm latis, levibus, hyalinis vel pallide 

brunnei compositum. Conidiophora dimorphosa: (A) Conidiophora ex hyphis 

lateraliter oriunda, subcylindrical, subhyalina vel pallide brunnea, levia, 1–6- 

septata, ad 40 µm longa et 2–3 µm lata. Cellulae conidiogenae terminales, 5–15 

× 2–3 µm, loco conidiogeno singulare et terminale, cellula ellipsoidea (conidio ?), 

persistente, interdum cellulis catenulatis (ad 6), pallide brunneo, apice subacute 

rotundato, basi truncata, 5–7 × 3–4 µm. (B) Conidiophoris 10–20 × 2–3 µm, 

1–2-septatis, subcylindraceis, rectis vel curvatis, subhyalinis vel pallide brunneis, 

levibus. Cellulae conidiogenae pallide brunneae, leviae, subcylindraceae, locis 

numerosis, aggregatis, inconspicuis vel subdenticulatis, leviter protuberantes, 0.5 

µm diam, incrassatis et fuscatis. Conidia catenulata (2–3), (4–)5–6(–7) × 2.5–3 µm, 

limoniformes vel ellipsoidea, non septata, levia, pallide brunnea, utrinque attenuata, 

apiculata, 0.5–1 × 0.5 µm, incrassata et fuscata, non refractiva. 

Mycelium consisting of branched, septate, 1.5–2 µm wide 

hyphae, smooth, hyaline to pale brown. Conidiophores dimorphic. 

Type A: Conidiophores forming as lateral branches on hyphae, 

subcylindrical, subhyaline to pale brown, smooth, 1–6-septate, up 

to 40 µm long, and 2–3 µm wide. Conidiogenous cells terminal, 5– 

15 × 2–3 µm, with a single, apical locus, giving rise to an ellipsoidal 

cell (conidium?) which mostly remains attached, pale brown, with 

a subacutely rounded apex and truncate base, 5–7 × 3–4 µm, at 

times forming chains of up to 6 such cells. Type B: Conidiophores 

10–20 × 2–3 µm, 1–2-septate, subcylindrical, straight to curved, 

subhyaline to pale brown, smooth. Conidiogenous cells pale 

brown, smooth, subcylindrical with numerous, aggregated loci, 

inconspicuous to subdenticulate and somewhat protruding, 0.5 µm 

wide, somewhat thickened and darkened. Conidia in chains of 2–3, 

limoniform to ellipsoid, widest in the middle, aseptate, smooth, pale 

brown, tapering towards ends that are prominently apiculate, 0.5–1 

µm long, 0.5 µm wide, prominently thickened and darkened, but 

not refractive. 

Cultural characteristics: Colonies on PDA slimy, spreading, 

somewhat erumpent in the centre, with even, catenulate margins, 

lacking aerial mycelium; surface fuscous-black to olivaceous-black, 

with patches of cream; reverse fuscous-black with patches of 

cream. Colonies reaching 25 mm diam on PDA after 1 mo at 25 °C 

in the dark; colonies fertile with profuse sporulation on SNA. 

Specimen examined: Netherlands, Oostelijk Flevoland, Jagersveld, isolated from 

Alnus glutinosa (Betulaceae), May 1982, W. Gams, holotype CBS-H 19895, culture 

ex-type CBS 261.82. 

Notes: Morphologically Hyalodendriella resembles the genera 

Hyalodendron and Retroconis de Hoog & Bat. Vegte (de Hoog 

& Batenburg van der Vegte 1989). It is distinct, however, in its 

pigmentation, dimorphic conidiophores and conidia. Furthermore, 

a strain of Retroconis fusiformis (S.M. Reddy & Bilgrami) de Hoog 

& Bat. Vegte (CBS 330.81) clusters apart from Hyalodendriella, 

namely in the Chaetomiaceae, Sordariales. 

Pleosporales, incertae sedis 

Ochrocladosporium Crous & U. Braun, gen. nov. MycoBank 

MB504437. 

Etymology: Named after its pale brown, cladosporium-like conidia. 

Differt a Cladosporio et generis cladosporioidibus diversis conidiophoris cum cellulis 

basalibus T-formibus et/vel cicatricibus non incrassatis, non vel leviter fuscatisrefractivis. 

Mycelium consisting of branched, septate hyphae, subhyaline to 

pale brown, smooth, giving rise to two types of conidiophores. 

Macronematous conidiophores solitary, erect, arising from 

superficial hyphae, composed of a subcylindrical stipe, without a 

swollen or lobed base or rhizoids, with or without a T-shaped foot cell, 

pale to dark brown; apical conidiogenous apparatus with or without 

additional branches, branched part, if present, with short branchlets 

composed of conidiogenous cells and ramoconidia, continuous to 

septate, wall thin or slightly thicked, pale brown. Conidiogenous cells 

integrated, terminal or intercalary, subcylindrical to doliiform, pale 

brown, thin-walled, smooth; unilocal or multilocal, determinate to 

sympodial, loci conically truncate, subdenticulate, neither thickened, 

nor darkened-refractive or only slightly darkened-refractive. 

Micronematous conidiophores integrated in hyphae, reduced to a 

lateral peg-like locus or erect, frequently reduced to conidiogenous 

cells, pale brown, smooth, subcylindrical. Conidia occurring in 

branched chains, fusiform, ellipsoid-ovoid to subcylindrical, 0(–1)- 

septate, ramoconidia present, pale brown, thin-walled, smooth to 

finely verruculose, ends attenuated, hila obconically truncate to 

almost pointed, neither thickened nor darkened-refractive. 

Type species: Ochrocladosporium elatum (Harz) Crous & U. Braun, 

comb. nov. 

Ochrocladosporium elatum (Harz) Crous & U. Braun, comb. 


Basionym: Hormodendrum elatum Harz, Bull. Soc. Imp. Naturalistes 

Moscou 44: 140. 1871. 

≡ Cladosporium elatum (Harz) Nannf., in Melin & Nannfeldt, Svenska 

Skogsvardsfoereren Tidskr. 32: 397. 1934. 

≡ Cadophora elatum (Harz) Nannf., in Melin & Nannfeldt, Svenska 

Skogsvardsfoereren Tidskr. 32: 422. 1934. 

Mycelium consisting of branched, septate, smooth, hyaline, 2–4 

µm wide, thin-walled, hyphae, becoming darker brown in places, 

giving rise to erect conidiophores. Conidiophores either reduced 

to conidiogenous cells, or well-differentiated, terminal and lateral 

on hyphae, erect, highly variable, arising from superficial and 

submerged hyphae, reduced to subdenticulate loci, 1–1.5 µm 

wide, or well-differentiated, up to 60 µm long, 1–3-septate, 3–4 

µm wide, hyaline to medium brown, smooth, thin-walled (≤ 1 µm). 

Conidiogenous cells integrated as lateral peg-like loci on hyphal 

cells, or erect, subcylindrical, up to 25 µm long, 2.5–4 µm wide, 

with 1–3 terminal loci, occasionally also lateral, 1–1.5 µm wide, 

not thickened and darkened, but frequently somewhat refractive 

(mounted in Shear’s solution, not lactic acid). Ramoconidia 

46


Fig. 9. Hyalodendriella betulae (type material). A. Conidiophores on PDA. B–C. Microconidiophores. D–H. Macroconidiophores with fascicles of conidiogenous cells. I. Conidia 

with darkened, thickened hila. Scale bars = 10 µm. 

subcylindrical to ellipsoid, hyaline to pale brown, smooth to finely 

verruculose, 10–40 × 3–5 µm, 0(–1)-septate, giving rise to branched 

chains of conidia (up to 20 per chain) that are subcylindrical to 

ellipsoid, aseptate, (7–)8–10(–14) × (3–)4(–4.5) µm, smooth to 

finely verruculose, olivaceous-brown, thin-walled (up to 0.5 µm), 

hila 0.5–1 µm wide, neither thickened nor, or barely, darkened 

refractive. 

Cultural characteristics: Colonies erumpent, spreading, fast 

growing, covering the plate within 1 mo at 25 °C; aerial mycelium 

abundant, margins smooth on PDA; surface isabelline in centre, 

umber in outer region; olivaceous-black in reverse. 

Specimen examined: Sweden, Iggesund, isolated from wood pulp, Jan. 1976, E. 

Melin, specimen CBS-H 19896, culture CBS 146.33. 

Notes: “Hormodendrum” elatum was originally described from 

a wooden stump in Germany. The culture examined here was 

deposited by Melin in 1933 as culture 389:14, isolated from wood 

chips in Sweden, and described by Nannfeldt, and has since 

been accepted as authentic for the species. Earlier publications 

(de Vries 1952, Ho et al. 1999, de Hoog et al. 2000), clearly state 

that this species does not belong in Cladosporium s. str., and this 

statement is supported by the phylogenetic analysis placing it in 

the Pleosporales. 


Ochrocladosporium frigidarii Crous & U. Braun, sp. nov. 


Etymology: Named after it collection site, within a cooled incubation 

room. 

Differt a O. elato conidiophoris distincte dimorphis, macroconidiophoris majoribus, 

ad 600 × 5–7 µm, septis incrassates, cellulis basalibus T-formibus et conidiis leniter 

brevioribus et latioribus, (6–)7–8(–10) × (4–)4.5–5(–6) µm. 

Mycelium consisting of branched, septate, 2–7 µm wide hyphae, 

occasionally constricted at septa with hyphal swellings, subhyaline 

to pale brown, smooth, thin-walled, giving rise to two types of 

conidiophores. Macronematous conidiophores solitary, erect, 

arising from superficial hyphae, up to 600 µm long, composed of 

a subcylindrical stipe, 5–7 µm wide, 10–15(–20)-septate, without 

a swollen or lobed base or rhizoids, but with a T-shaped foot cell, 

wall ≤ 1 µm wide, guttulate, with thick septa, dark brown, finely 

verruculose, apical 1–2 cells at times medium brown, giving rise 

to 1–2 primary branches, 0–1-septate, subcylindrical, thin-walled, 

pale brown, smooth to finely verruculose, 10–20 × 4–6 µm, giving 

rise to (1–)2–4 secondary branches, 0–1-septate, subcylindrical, 

8–13(–20) × 4–5 µm, or giving rise directly to conidiogenous cells. 

Conidiogenous cells subcylindrical to doliiform, pale brown, smooth, 

8–15 × 3–4 µm, loci somewhat protruding 1–2 µm wide, neither 

47

Crous et al. 

Fig. 10. Ochrocladosporium elatum (CBS 146.33). A–C, E. Microconidiophores. D. Macro- and microconidiophore. F–H. Macroconidiophores. I–J. Ramoconidia and conidia. 


thickened, darkened, nor refractive. Micronematous conidiophores 

erect, pale brown, smooth, subcylindrical, reduced to conidiogenous 

cells, or up to 4-septate, 15–90 × 2–3.5 µm, mostly unbranched, 

rarely branched below; conidiogenous cells subcylindrical, 

pale brown, smooth to finely verruculose, tapering at apex and 

sometimes at base, proliferating sympodially via 1(–3) loci, 1–1.5 

µm wide, denticle-like, which can appear somewhat darkened; 

micronematous conidiophores frequently occurring at the base of 

macronematous conidiophores. Ramoconidia, if present, up to 30 

µm long, 0–1-septate. Conidia and ramoconidia ellipsoid to ovoid, 

aseptate, pale brown, thin-walled (≤ 0.75 µm), finely verruculose, 

occurring in branched chains; conidia (6–)7–8(–10) × (4–)4.5–5(–6) 

µm; hila 0.5–1 µm wide, not darkened, thickened or refractive. 

Cultural characteristics: Colonies on PDA erumpent, spreading, with 

profuse sporulation and moderate aerial mycelium, even margins, 

olivaceous-grey (surface); reverse olivaceous-black. Colonies 

covering the dish after 1 mo at 25 °C in the dark. 

Specimen examined: Germany, Hannover, isolated from a cooled room, Jan. 1981, 

B. Ahlert, holotype CBS-H 19897, culture ex-type CBS 103.81. 

Notes: Ochrocladosporium frigidarii is characterised by its 

dimorphic fruiting, and inconspicuous scars and conidial hila, which 

48


Fig. 11. Ochrocladosporium frigidarii (type material). A. Macro- and microconidiophores. B. Foot cell of macroconidiophore. C–D. Microconidiophore. E–J. Macroconidiophores. 

K. Conidia. Scale bars = 10 µm. 

are distinct from Cladosporium s. str. The phylogenetic analysis of 

its LSU sequence places it in the Pleosporales, together with O. 

elatum. 

The dimorphic conidiophores seen in O. frigidarii (CBS 103.81) 

are less obvious in O. elatum (CBS 146.33), but the scars and hila 

are similar. The macronematous conidiophores of O. frigidarii are 

much longer and wider and the conidia are shorter and slightly 

wider, (6–)7–8(–10) × (4–)4.5–5(–6) µm, than those of O. elatum 

which are (7–)8–10(–14) × (3–)4(–4.5) µm. 


49

Crous et al. 

Fig. 12. Rhizocladosporium argillaceum (type material). A–E. Conidiophores with pigmented ramoconidia and hyaline conidia. F. Rhizoids forming at the foot cells of 

macroconidiophores. Scale bar = 10 µm. 


Rhizocladosporium Crous & U. Braun, gen. nov. MycoBank 

MB504440. 

Etymology: Named after the presence of rhizoids on its 

conidiophores, and cladosporium-like conidia. 

Differt a Cladosporio et generis cladosporioidibus diversis hyphis hyalinis, 

conidiophoris cum cellulis basalibus lobatis vel rhizoidibus, cellulis conidiogenis 

monoblasticis, determinatis, locis margine leviter incrassatis et fuscatis, non 

refractivis, non coronatis, ramoconidiis brunneis sed conidiis hyalinis, hilis non 

incrassatis, non fuscatis-refractivis. 

Mycelium consisting of branched, septate, smooth, hyaline hyphae. 

Conidiophores solitary, macronematous, subcylindrical, erect, 

arising from superficial mycelium, septate, pigmented, smooth; 

base somewhat inflated, lobed or with rhizoids. Conidiogenous 

cells integrated, terminal, monoblastic, determinate, subcylindrical, 

tapering towards a single flat-tipped locus, straight to once 

geniculate, occasionally with two loci, pigmented, smooth; locus 

flattened, undifferentiated to somewhat darkened and thickened 

along the rim, not refractive, giving rise to a single conidial chain 

or a single ramoconidium with several simple acropetal chains of 

secondary ramoconidia or conidia. Conidia occurring in branched 

chains; ramoconidia subcylindrical to narrowly ellipsoidal, straight 

to geniculate-sinuous, with apical and lateral conidial hila; 

ramoconidia with broadly truncate base medium brown; secondary 

ramoconidia with narrowed base subhyaline or hyaline, smooth; 

conidia aseptate, in chains, hyaline, guttulate, ellipsoidal with 

obtuse ends; hila inconspicuous, neither darkened nor refractive 

or thickened. 

Type species: Rhizocladosporium argillaceum (Minoura) Crous & 

U. Braun, comb. nov. 

Rhizocladosporium argillaceum (Minoura) Crous & U. Braun, 

comb. nov. MycoBank MB504441. Fig. 12. 

Basionym: Cladosporium argillaceum Minoura, J. Ferment. Technol. 

44: 140. 1966. 

Mycelium consisting of branched, septate, smooth, hyaline, 

thin-walled, 1.5–2 µm wide hyphae. Conidiophores solitary, 

macronematous, erect, arising from superficial mycelium; base 

somewhat inflated, lobed or with rhizoids, up to 10 µm wide; 

conidiophore stipe subcylindrical, straight to curved, rarely 

geniculate-sinuous, wall up to 1 µm wide, medium brown, 

sometimes paler towards the tip, smooth, 1–6-septate, 35–160 

50


µm tall, 4–6 µm wide. Conidiogenous cells terminal, straight, 

subcylindrical, tapering towards a flat-tipped locus, occasionally 

once geniculate, with two loci, medium brown, smooth, 15–35 × 

4–6 µm; locus flattened, undifferentiated or very slightly darkened 

and thickened along the rim, not refractive, 1.5–2 µm wide. Conidia 

occurring in branched chains. Ramoconidia subcylindrical to 

narrowly ellipsoidal, straight to geniculate-sinuous, 17–35 × 4–5 

µm, medium brown, smooth, thin-walled, frequently branching 

laterally, with apical and lateral subdenticulate conidial hila, 1.5–2.5 

µm wide; secondary ramoconidia hyaline or subhyaline. Conidia 

aseptate, (10–)12–17(–20) × (3.5–)4(–4.5) µm, in branched 

chains (–6), hyaline or subhyaline, guttulate, ellipsoidal-fusiform, 

with obtuse ends, or tapering to obconically subtruncate ends with 

hila that are inconspicuous (neither darkened nor refractive or 

thickened), 0.5–1 µm wide. 

Cultural characteristics: Colonies on PDA spreading, erumpent, with 

smooth, even margins and sparse to moderate aerial mycelium; 

hazel to fawn (surface); reverse hazel to fawn. Colonies reaching 

35 mm diam after 1 mo at 25 °C in the dark; colonies fertile. 

Specimen examined: Japan, Yoku Island, isolated from decayed myxomycete, 21 

Oct. 1961, K. Tubaki No. 4262 holotype, culture ex-type CBS 241.67 = IFO 7055. 

Notes: The lobed-rhizoid conidiophore base, and brown, 

disarticulating ramoconidia, with hyaline chains of conidia, are 

characteristic of Rhizocladosporium. Although Minoura (1966) 

illustrated some conidiophores that were micronematous (reduced 

to conidiogenous cells on superficial mycelium), these were 

not observed in the present study. Metulocladosporiella Crous, 

Schroers, J.Z. Groenew., U. Braun & K. Schub. (Crous et al. 

2006a) (Herpotrichiellaceae), comprising two banana leaf-spotting 

pathogens, is another cladosporioid hyphomycete genus having 

distinct rhizoid hyphae at the swollen base of conidiophores. It 

differs, however, in having conidiophores terminally branched in 

a metula-like manner and distinct conidiogenous loci and conidial 

hila. Pleurotheciopsis B. Sutton (Ellis 1976) is also characterised 

by having pigmented conidiophores and hyaline or pale, septate 

conidia formed in acropetal chains, but the conidiophores 

proliferate percurrently, the conidiogenous cells are polyblastic and 

ramoconidia are lacking, i.e., the conidia are formed in unbranched 

chains. Parapleurotheciopsis P.M. Kirk (Kirk 1982) is very similar to 

Rhizocladosporium. The conidiophores possess a single terminal 

unilocal conidiogenous cell giving rise to a single ramoconidium 

which forms several chains of acropetal, aseptate, hyaline to pale 

olivaceous conidia. The base of the conidiophores is somewhat 

swollen and lobed [except for Parapleurotheciopsis coccolobae 

R.F. Castañeda & B. Kendr., Castañeda & Kendrick (1990), with at 

most slightly swollen, but unlobed base]. However, R. argillaceum 

occasionally has once-geniculate conidiogenous cells with two 

loci. Furthermore, it clusters in the Helotiales (Fig. 1), whereas a 

sequenced strain of Parapleurotheciopsis inaequiseptata (MUCL 

41089), belongs to the Xylariales (Fig. 2). The occasionally 

occurring conidiogenous cells with two loci and the aseptate conidia 

connect Rhizocladosporium with Subramaniomyces Varghese & 

V.G. Rao (Varghese & Rao 1979, Kirk 1982) in which, however, 

terminal ramoconidia are lacking. Furthermore, the type species, 

S. fusisaprophyticus (Matsush.) P.M. Kirk, frequently has branched 

conidiophores. Subramaniomyces simplex U. Braun & C.F. Hill 

(Braun & Hill 2002), a species with unbranched conidiophores is, 

however, morphologically similar to R. argillaceum, but the genus 

Subramaniomyces is phylogenetically distinct and also belongs to 

the Xylariales (CBS 418.95, Fig. 2). 

Key to Cladosporium and morphologically similar genera 

(bearing simple or branched acropetal chains of amero- to phragmosporous blastoconidia) 

1. Conidiophores and conidia hyaline............................................................................................................................................................. 2 

1. At least conidiophores pigmented .............................................................................................................................................................. 4 

2. Conidia in simple chains ........................................................................................................................................................... Hormiactis 

2. Conidia in branched chains ....................................................................................................................................................................... 3 

3. Conidiogenous cells sympodial, with distinct conidiogenous loci (scars), thickened and darkened; conidia amero- to phragmosporous; 

plant pathogenic, leaf-spotting fungi (Mycosphaerella anamorphs; Mycosphaerellaceae) ......................................................... Ramularia 

3. Terminal conidiogenous cells with denticle-like loci, giving rise to ramoconidia which form simple or branched conidial chains; 

lignicolous ........................................................................................................................................................................... Hyalodendron 

[Conidiophores dimorphic; mycelium, conidiophores and conidia at first hyaline, later turning pale brown; conidia in short chains, see 

Hyalodendriella] 

4(1). Conidia distoseptate, in simple chains .............................................................................................................................................. Lylea 

4. Conidia aseptate or euseptate ................................................................................................................................................................... 5 

5. Conidiophores little differentiated, micronematous to semimacronematous; conidiogenous loci undifferentiated, truncate, neither 

distinctly thickened nor darkened or only very slightly so .......................................................................................................................... 6 

5. Conidiophores well-differentiated, semimacronematous (but multilocal and/or conidiogenous loci well-differentiated) to 

macronematous ....................................................................................................................................................................................... 12 

6. Conidiophores and conidia delicate, thin-walled, in long, easily disarticulating chains .............................................................................. 7 

6. Conidiophores and conidia robust, wall thickened, dark, conidial chains often seceding with difficulty .................................................... 9 

7. Conidiophores semimacronematous, simple to often irregularly branched; conidia delicate, narrow, 1–3 µm wide, hyaline to pale 

olivaceous ............................................................................................................................................................................ Polyscytalum 


51

Crous et al. 

7. Conidiophores unbranched, micronematous or semimicronematous, integrated in ordinary hyphae, forming minute, lateral, 

monoblastic, determinate, peg-like protuberances to semimacronematous, forming short lateral branches (conidiophores) with several 

inconspicuous to denticle-like loci .............................................................................................................................................................. 8 

8. Phialidic synanamorphs often present, but sometimes also lacking; saprobic, rarely plant pathogenic, often human pathogenic 

(Herpotrichiellaceae, Chaetothyriales) .......................................................................................................................... Cladophialophora 

8. Without phialidic synanamorphs; saprobic or plant pathogenic (Venturia, Venturiaceae).............................................................................. 

.......................................................................................................................................... Fusicladium s. lat. (incl. Pseudocladosporium) 

9(6). Conidia aseptate, rarely 1-septate; lignicolous, on dead wood ............................................................................................... Xylohypha 

9. Conidia septate ........................................................................................................................................................................................ 10 

10. Conidia 1-septate, with a dark brown to blackish band at the septum; on dead wood .................................................................. Bispora 

10. Conidia at least partly 2- to pluriseptate and/or without dark brown to blackish band at the septum ...................................................... 11 

11. Conidia branched ....................................................................................................................................................................... Taeniolina 

11. Conidia unbranched ................................................................................................................................................................... Taeniolella 

12(5). Conidiogenous loci and conidial hila distinctly coronate, i.e., composed of a central convex dome surrounded by a periclinal raised rim, 

mostly at least somewhat protuberant (anamorphs of Davidiella, Davidiellaceae, Capnodiales) ............................. Cladosporium s. str. 

12. Conidiogenous loci non-coronate (either inconspicuous, thickened and darkened or denticle-like) ........................................................ 13 

13. Mycelium, conidiophores and conidia at first hyaline or subhyaline, later turning pale brown; conidiophores dimorphic, either 

conidiogenous cells with a single conidiogenous locus, giving rise to an ellipsoid cell (conidium?) which mostly remains attached, 

base truncate, apex subacutely rounded, at times forming chains of such cells; or conidiophores with numerous aggregated loci, 

inconspicuous to subdenticulate; conidia in short chains, of mostly 2–3 (isolated from Alnus in Europe) ........................ Hyalodendriella 

13. Fruiting different; at least conidiophores consistently pigmented or conidiophores uniform or loci distinct; conidia mostly in long, often 

branched chains ...................................................................................................................................................................................... 14 

14. Conidiophores with verruculose conidiogenous apices, otherwise smooth; conidia distinctly verruculose-verrucose; conidiogenous loci 

and conidial hila inconspicuous ............................................................................................................................................................... 15 

14. Conidiophores either smooth throughout or verruculose below and smooth above or verruculose throughout; and/or conidiogenous loci 

conspicuous, i.e., thickened and darkened or denticle-like ...................................................................................................................... 16 

15. Conidiophores macronematous, unbranched, base swollen, with percurrent regenerative proliferations, unrelated to conidiation; 

conidiogenous cells terminal, occasionally also subterminal; conidia terminally and laterally formed, aseptate (saprobic on leaves) 

................................................................................................................................................................................................ Castanedaea 

15. Conidiophores little differentiated, semimacronematous, unbranched or with short lateral branchlets, base undifferentiated, 

without percurrent proliferations; conidiogenous cells terminal and occasionally intercalary-pleurogenous; conidia terminally and 

subterminally formed, 0–2-septate (lignicolous, on decorticated wood) .......................................................................... Websteromyces 

16(14). Conidiophores unbranched, with a simple terminal conidiogenous cell, non-geniculate-sinuous, subcylindrical to somewhat inflated at 

the tip; conidiogenous loci terminal and lateral, inconspicuous or subconspicuous, neither thickened nor darkened, non-protuberant; 

conidia attached with a very narrow, pointed hilum ................................................................................................................................. 17 

16. Conidiophores with a branched terminal conidiogenous apparatus, composed of conidiogenous cells and/or ramoconidia or conidiophores 

unbranched, with a single terminal conidiogenous cell or additional intercalary ones, but conidiogenous loci different, conspicuous, 

thickened and darkened or denticle-like .................................................................................................................................................. 18 

17. Conidiophores with distinct rhizoid-digitate base; tips of the conidiogenous cells somewhat swollen, usually unilaterally swollen or 

somewhat curved; conidia solitary or only in very short unbranched chains; hyperparasitic on rusts .................................. Digitopodium 

17. Conidiophores without rhizoid-digitate base; tips of the conidiogenous cells subcylindrical to somewhat swollen, but swellings not unilateral 

and not curved; conidia solitary and in simple or branched chains; associated with leaf spots ................................. Rachicladosporium 

18(16). Conidiophores in synnematous conidiomata ..................................................................................................................................... 19 

18. Synnemata lacking .................................................................................................................................................................................. 20 

19. Conidiogenous cells with a single or several truncate to subdenticulate, relatively broad conidiogenous loci; conidia with truncate, flat hila; 

on wood, resin ............................................................................................................................................................................ Sorocybe 

19. Conidiogenous loci with few, mostly 1–2 conidiogenous loci formed as minute spicules; conidia with narrow hila (shallowly apiculate); 

plant pathogenic, causing bud blast and twig blight ....................................................................................................................... Seifertia 

20(18). Conidiophores unbranched or occasionally branched; conidiogenous cells distinctly inflated, ampulliform, doliiform or clavate, nondenticulate; 

conidia at least partly globose, dark brown when mature; colonies effuse, dark; wood-inhabiting .......... Phaeoblastophora 

52


20. Conidiogenous cells not inflated, if somewhat inflated, loci denticle-like or conidia non-globose ............................................................ 21 

21. Conidiophores penicillate, i.e., with an unbranched stipe and distinct terminal branched “head” composed of branchlets, conidiogenous 

cells and/or ramoconidia .......................................................................................................................................................................... 22 

21. Conidiophores non-penicillate, i.e., irregularly and loosely branched, branchings not confined to the apical portion, sometimes only with 

short lateral branchlets, or unbranched ................................................................................................................................................... 27 

22. Penicillate apex simple, only composed of a single terminal conidiogenous cells giving rise to several ramoconidia which form secondary 

ramoconidia and conidia ......................................................................................................................... Penidiella p.p. [P. strumelloidea] 

22. Penicillate apex more complex, composed of true branchlets and/or conidiogenous cells and ramoconidia .......................................... 23 

23. Conidiophores with a compact, dense, subglobose to broadly ovoid head; conidiogenous loci and conidial hila unthickened or almost so, 

but distinct by being darkened-refractive [fruiting dimorphic, periconioid branched conidiophores formed on overwintered 

stem of Paeonia spp., unbranched cladosporioid conidiophores on leaf spots, biotrophic] (belonging to the Capnodiales) 

.................................................................................................................................................................................... Dichocladosporium 

23. Penicillate apex looser, neither compact nor subglobose ........................................................................................................................ 24 

24. Branched head composed of short branchlets and conidiogenous cells; ramoconidia lacking; conidiogenous cells subcylindrical to 

subclavate, non-geniculate; conidiogenous loci usually numerous and aggregated, terminal and lateral, non-protuberant, flat, conspicuous, 

thickened and darkened, at least around the rim; conidia solitary or in short chains ............................................................... Periconiella 

24. Ramoconidia often present; conidiogenous cells distinct, sympodial, somewhat geniculate or subdenticulate; conidiogenous loci 

inconspicuous or somewhat protruding, denticle-like, unthickened or almost so, not or somewhat darkened-refractive; conidia in long, 

often branched chains ............................................................................................................................................................................. 25 

25. Branched apex composed of short branchlets consisting of conidiogenous cells or ramoconidia, in pairs or whorls of 3–4, mostly distinctly 

constricted at the base; hyperparasitic on Asterina spp. ................................................................................................. Parapericoniella 

25. Branched apex distinct, composed of branchlets, conidiogenous cells and/or ramoconidia, if true branchlets lacking conidiogenous cells 

and ramoconidia not in whorls and not distinctly constricted at the base; saprobic or biotrophic ............................................................ 26 

26. Penicillate apex of the conidiophores loosely to densely branched, occasionally metula-like, base of the conidiophores simple, 

undifferentiated; saprobic or biotrophic (Teratosphaeriaceae, Capnodiales) .............................................................................. Penidiella 

26. Penicillate apex always dense, metula-like, base of the conidiophores swollen or lobed, often with rhizoid hyphae; plant pathogenic [on 

banana] (Chaetothyriales) ........................................................................................................................................ Metulocladosporiella 

27(21). Conidiophores simple or branched; septa of the conidiophores and conidia becoming thick-walled and dark; conidiogenous loci 

subdenticulate, somewhat thickened and conspicuously darkened-refractive; cultures producing ample amounts of volatile metabolites 

causing skin irritation after exposure to the fungus; saprobic (isolated from mouldy paint) ...................................... Toxicocladosporium 

27. Without conspicuously thickened-darkened septa; cultures without irritant, volatile metabolites ............................................................ 28 

28. Conidiogenous loci conspicuous, distinctly thickened and darkened (visible as small dark circles when viewed upon the scar), sometimes 

on small shoulders formed by sympodial proliferation, but not distinctly denticulate (Capnodiales) ....................................................... 29 

28. Conidiogenous loci inconspicuous or conspicuous by being denticle-like, not or barely thickened, not darkened or at most upper truncate 

end very slightly thickened and somewhat darkened-refractive .............................................................................................................. 31 

29. Mycelium smooth; conidiophores and conidia smooth or almost so, at most faintly rough-walled; conidiophores solitary, fasciculate, 

sporodochial to synnematous; biotrophic, usually leaf-spotting (Mycosphaerella anamorphs, Mycosphaerellaceae) 

................................................................................................................. Passalora emend. (incl. Mycovellosiella, Phaeoramularia, etc.) 

29. At least mycelium distinctly verruculose .................................................................................................................................................. 30 

30. Mycelium, conidiophores and conidia coarsely verruculose-verrucose; conidial shape variable, often irregular; isolated from a lichen 

(Dirina) .................................................................................................................................................................... Verrucocladosporium 

30. Mycelium verruculose; conidiophores mostly smooth, sometimes somewhat rough-walled, conidia smooth to distinctly verruculose; 

biotrophic, often leaf-spotting ......................................................................................................................................................... Stenella 

31(28). Conidiophores with swollen, often lobed base ................................................................................................................................... 32 

31. Conidiophores without swollen base, at most slightly swollen, but not lobed .......................................................................................... 35 

32. Conidia septate ........................................................................................................................................................................................ 33 

32. Conidia aseptate ...................................................................................................................................................................................... 34 

33. Conidiophores with a single, terminal, monoblastic, determinate conidiogenous cell giving rise to a single ramoconidium that forms 

simple or branched chains of conidia ...................................................................................................................... Parapleurotheciopsis 


53

Crous et al. 

33. Terminal conidiogenous cells polyblastic, with several denticle-like conidiogenous loci ................. Anungitea p.p. (e.g. A. longicatenata) 

34(32). Conidiogenous cells terminal, monoblastic, with a single ramoconidium giving rise to conidial chains or occasionally with 2(–3) 

denticle-like loci; base of the conidiophores often with rhizoid hyphae ...................................................................... Rhizocladosporium 

34. Conidiogenous cells polyblastic, with two or several denticle-like loci; base of the conidiophores without rhizoid hyphae 

...................................................................................................................................................................................... Subramaniomyces 

35(31). Conidiophores unbranched, with a terminal monoblastic conidiogenous cell, determinate or percurrent ......................................... 36 

35. Conidiophores branched or unbranched, but conidiogenous cells at least partly polyblastic .................................................................. 38 

36. Conidiogenous cell giving rise to a single ramoconidium which forms simple or branched chains of 0(–1)-septate conidia 

................................................................................................................................................ Parapleurotheciopsis p.p. (P. coccolobae) 

36. Conidiogenous cells giving rise to simple conidial chains without ramoconidia; conidia septate ............................................................ 37 

37. Conidiophores sometimes with percurrent proliferations; conidiophores and conidia with somewhat thickened, dark walls; conidia 1–10- 

septate, width usually exceeding 4 µm ................................................................................................................................ Heteroconium 

37. Percurrent proliferations lacking; conidiophores and conidia delicate, thin-walled and paler; conidia usually 0–1(–3)-septate and narrow, 

usually below 4 µm wide (Chaetothyriales) ......................................................................... Cladophialophora p.p. (e.g. C. chaetospira) 

[similar anamorphs of the Venturiaceae, see Fusicladium (incl. Pseudocladosporium)] 

38(35). Conidiophores often branched; conidiogenous loci distinctly denticle-like or subdenticulate; conidia aseptate; lignicolous, on dead 

wood, resin or isolated from hydrocarbone-rich substrates (jet-fuel, cosmetics, etc.) ............................................................................. 39 

38. Either with unbranched conidiophores or conidiogenous loci not distinctly denticle-like, or conidia septate, or on other substrates 

.................................................................................................................................................................................................................. 42 

39. Conidiogenous cells distinctly denticulate; conidia rather broad, approx. 7–13 µm ............................................................. Haplotrichum 

39. Conidiogenous cells non-denticulate or at most subdenticulate; conidia narrower, approx. 3–6 µm ...................................................... 40 

40. Colonies effuse, dense, but felted, black, brittle and appearing carbonaceous when dry; conidiophores solitary, brown; conidiogenous cells 

terminal and pleurogenous; conidia pale to dark brown, lateral walls conspicuously thicker than the hila; on conifer resin 

......................................................................................................................... Sorocybe (mononematous form, Hormodendrum resinae) 

40. Colonies effuse, dense, resupinate, hypochnoid, powdery, chocolate-brown and/or conidiophores lightly pigmented; conidia subhyaline 

to lightly pigmented and/or lateral walls not thickener than poles; on dead wood or isolated from hydrocarbone-rich substrates (jet-fuel, 

cosmetics, etc.) ........................................................................................................................................................................................ 41 

41. Colonies effuse, dense, resupinate, hypochnoid, powdery, chocolate-brown; conidiophores smooth; conidia subhyaline to very pale 

yellowish, hila very thin; on dead wood ......................................................................................................................... Parahaplotrichum 

41. Colonies neither resupinate nor hypochnoid; conidiophores warty; lateral walls of the conidia not thicker than the hila; isolated from 

hydrocarbone-rich substrates (jet-fuel, cosmetics, etc.) ......................................................................................................... Hormoconis 

42(38). Conidiophores simple or branched; conidiogenous cells monoblastic or occasionally polyblastic; conidiogenous loci subdenticulate, 

neither thickened nor darkened, forming simple or branched chains of regular conidia, uniform in shape, size and septation 

................................................................................................................................................................................................... Septonema 

42. Conidia not uniform in shape, size and septation; conidiogenous loci flat-tipped, subdenticulate, unthickened or slightly so, not to 

somewhat darkened-refractive ................................................................................................................................................................. 43 

43. Conidiophores simple or branched; in culture forming abundant chlamydospores; mostly soil-borne and heat-resistant (Teratosphaeriaceae, 

Capnodiales) ................................................................................................................................................................................ Devriesia 

43. Without chlamydospores in culture; phylogenetically distinct .................................................................................................................. 44 

44. Conidiophores dimorphic; conidia mostly aseptate, hila inconspicuous, neither thickened nor darkened (Pleosporales) 

.................................................................................................................................................................................... Ochrocladosporium 

44. Conidiophores either uniform or conidia at least partly septate or hila more conspicuous by being slightly thickened or at least somewhat 

darkened or refractive; phylogenetically distinct ...................................................................................................................................... 45 

45. Phialidic synanamorphs often present, but sometimes also lacking; saprobic, rarely plant pathogenic, often human pathogenic 

(Herpotrichiellaceae, Chaetothyriales) .......................................................................................................................... Cladophialophora 

45. Without phialidic synanamorphs; saprobic or plant pathogenic; phylogenetically distinct ....................................................................... 46 

46. Conidiophores usually unbranched (Venturia, Venturiaceae) .......................................... Fusicladium s. lat. (incl. Pseudocladosporium) 

[similar, barely distinguishable taxa, also clustering in the Venturiaceae, but apart from the Venturia clade are tentatively referred to as 

Anungitea until this genus will be resolved by sequences of its type species] 

54


46. Conidiophores simple to often irregularly branched; conidia delicate, narrow, 1–3 µm wide, hyaline to pale olivaceous (not belonging to 

the Venturiaceae) .................................................................................................................................................................. Polyscytalum 

Discussion 

Phylogenetic studies conducted on species of Cladosporium s. lat. 

proved the genus to be highly heterogeneous (Braun et al. 2003). It 

could be demonstrated that various anamorphs, previously referred 

to as Cladosporium, e.g. Cladosporium fulvum Cooke [≡ Passalora 

fulva (Cooke) U. Braun & Crous], have to be excluded since they 

clustered in the Mycosphaerella clade (Mycosphaerellaceae). 

Previous re-examinations and reassessments of human pathogenic 

Cladosporium species, including morphology, biology/ecology, 

physiology and molecular data (Masclaux et al. 1995, Untereiner 

1997, Gerrits van den Ende & de Hoog 1999, Untereiner & Naveau 

1999, Untereiner et al. 1999; de Hoog et al. 2000), could also be 

confirmed. In all phylogenetic analyses, it could be shown that the 

human pathogenic fungi concerned formed a clade belonging to 

the Herpotrichiellaceae (Capronia Sacc./Cladophialophora Borelli). 

Venturia anamorphs with catenate conidia, previously often 

assigned to Cladosporium s. lat., clustered together with other 

anamorphs of the Venturiaceae, and formed a monophyletic clade 

(Braun et al. 2003, Schubert et al. 2003, Beck et al. 2005). Venturia 

has now also been shown to accommodate less well-known 

anamorph genera such as Pseudocladosporium, which represent 

an additional synonym of Fusicladium Bonord. (Crous et al. 2007 

– this volume). 

Seifert et al. (2004) examined morphological, ecological and 

molecular characters of Cladosporium staurophorum (W.B. Kendr.) 

M.B. Ellis and three allied heat-resistant species and placed them 

in the new genus Devriesia, which formed a monophyletic group 

apart from the Cladosporium clade. Crous et al. (2006b) erected 

the genus Cladoriella Crous for a saprobic species (incertae sedis) 

characterised by having narrowly ellipsoidal to cylindrical or fusoid, 

0–1-septate, medium brown, thick-walled, finely verruculose 

conidia arranged in simple or branched chains, with thickened, 

darkened, refractive hila, with a minute central pore. Cladosporium 

musae E.W. Mason, the causal agent of banana speckle disease, 

has recently been shown to be allied to the Chaetothyriales (Crous 

et al. 2006a), and was placed in a new genus, Metulocladosporiella 

with C. musae as type species. Digitopodium U. Braun, Heuchert 

& K. Schub. (type species: Cladosporium hemileiae Steyaert) 

and Parapericoniella U. Braun, Heuchert & K. Schub. (type 

species: Cladosporium asterinae Deighton) represent two new 

genera of hyperparasitic hyphomycetes, introduced due to unique 

morphological features and striking differences to Cladosporium s. 

str. (Heuchert et al. 2005), but have as yet been excluded from 

DNA-based studies due to the absence of cultures. Schubert et al. 

(2007a – this volume) introduced a new genus, Dichocladosporium 

K. Schub., U. Braun & Crous (allied to the Davidiellaceae, Capnodiales) 

to accommodate a fungus with dimorphic fruiting that is 

pathogenic to Paeonia spp. The present study introduced yet several 

additional cladosporium-like genera, which could be distinguished 

based on their morphology and distinct DNA phylogeny, namely 

Ochrocladosporium (Pleosporales), Rhizocladosporium 

(incertae sedis), Rachicladosporium, Toxicocladosporium and 

Verrucocladosporium (Capnodiales). 

Although all these genera are cladosporium-like, and many 

have in the past been confused with Cladosporium s. str., the unique 


coronate scar type of Cladosporium s. str. allows a critical revision 

of cladosporioid hyphomycetes, based on reliable, distinctive 

morphological characters. In all cases where cladosporium-like 

(Cladosporium s. lat.) hyphomycetes clearly clustered apart 

from Cladosporium s. str. in the phylogenetic analyses, it could 

be demonstrated that the fungal groups concerned were also 

morphologically unambiguously distinguished, above all with regard 

to the structure of the conidiogenous hila. Hence, the excluded 

groups of species, belonging in other genera, sometimes even 

in new genera, are genetically as well as morphologically clearly 

distinct from Cladosporium s. str. 


We thank F. Hill, A. Aptroot, F.M. Dugan, R.F. Castañeda and W. Gams for providing 

collections and cultures of Cladosporium and cladosporium-like species over the 

past few years, without which this study would not have been possible. A research 

visit of K. Schubert to CBS was supported by a Synthesys grant (No. 2559), and 

the Odo van Vloten Stichting. We thank M. Vermaas for preparing the photographic 

plates, and A. van Iperen for preparing all the fungal cultures for examination. H.-J. 

Schroers is thanked for generating some of the sequence data used in this paper. 

A.J.L. Phillips is thanked for providing comments on an earlier draft of the paper. 

References 



Arzanlou M, Groenewald JZ, Gams W, Braun U, Shin H-D, Crous PW (2007). 

Phylogenetic and morphotaxonomic revision of Ramichloridium and allied 

genera. Studies in Mycology 58: 57–93. 

Arx JA von (1974). The Genera of Fungi Sporulating in Pure Culture. 2 nd edn. J. 

Cramer, Vaduz. 

Arx JA von (1981). The Genera of Fungi Sporulating in Pure Culture. 3 rd edn. J. 

Cramer, Vaduz. 

Arx JA von (1983). Mycosphaerella and its anamorphs. Proceedings of the 

Koninklijke Nederlandse Akademie van Wetenschappen, Ser. C, 86(1): 15–54. 

Beck A, Ritschel A, Schubert K, Braun U, Triebel D (2005). Phylogenetic relationship 

of the anamorphic genus Fusicladium s. lat. as inferred by ITS data. Mycological 

Progress 4: 111–116. 

Braun U (1995). A monograph of Cercosporella, Ramularia and allied genera 

(phytopathogenic hyphomycetes) Vol. 1. IHW-Verlag, Eching. 

Braun U (1996). Taxonomic notes on some species of the Cercospora complex (IV). 

Sydowia 48: 205–217. 


(phytopathogenic hyphomycetes) Vol. 2. IHW-Verlag, Eching. 




Braun U, Hill CF (2002). Some new micromycetes from New Zealand. Mycological 

Progress 1: 19–30. 




Castañeda RF, Kendrick B (1990). Conidial fungi from Cuba II. University of Waterloo 

Biological Series 33: 1–61. 



Crous PW, Aptroot A, Kang J-C, Braun U, Wingfield MJ (2000). The genus 


Crous PW, Braun U (2003). Mycosphaerella and its anamorphs: 1. Names published 

in Cercospora and Passalora. CBS Biodiversity Series 1: 1–569. 

Crous PW, Groenewald JZ, Mansilla JP, Hunter GC, Wingfield MJ (2004). 



55

Crous et al. 



1081–1101. 




Crous PW, Schubert K, Braun U, Hoog GS de, Hocking AD, Shin H-D, Groenewald 





A, Burgess T, Barber P, Groenewald JZ (2006d). Phylogenetic lineages in the 


Crous PW, Verkley GJM, Groenewald JZ (2006b). Eucalyptus microfungi known 

from culture. 1. Cladoriella and Fulvoflamma genera nova, with notes on some 

other poorly known taxa. Studies in Mycology 55: 53–63. 

Crous PW, Wingfield MJ, Mansilla JP, Alfenas AC, Groenewald JZ (2006c). 


occurring on Eucalyptus. II. Studies in Mycology 55: 99–131. 





Ellis MB (1971). Dematiaceous hyphomycetes. Commonwealth Mycological 


Ellis MB (1976). More dematiaceous hyphomycetes. Commonwealth Mycological 



from the Red Sea Coast of Egypt. Fungal Diversity 5: 43–54 

Gams W, Verkley GJM, Crous PW (2007). CBS Course of Mycology, 5 th ed. 



of the neurotropic species Cladophialophora bantiana. Studies in Mycology 43: 

151–162. 

Goh T-K, Hyde KD (1998). A synopsis of and key to Diplococcium species, based on 

the literature, with a description of a new species. Fungal Diversity 1: 65–83. 

Hawksworth DL (1979). The lichenicolous hyphomycetes. Bulletin of the British 

Museum (Natural History), Botany 6: 183–300. 





72: 115–157. 

Hoog GS de, Batenburg van der Vegte WH (1989). Retroconis, a new genus of 

ascomycetous hyphomycetes. Studies in Mycology 31: 99–105. 



Hoog GS de, Guarro, J, Gené J, Figueras MJ (2000). Atlas of clinical fungi, 2 nd ed. 

Centraalbureau voor Schimmelcultures, Utrecht and Universitat rovira I virgili, 

Reus. 

Inacio J, Pereira P, de Cavalho M, Fonseca A, Amaral-Collaco MT, Spencer-Martins 

I (2002). Estimation and diversity of phylloplane mycobiota on selected plants in 

a Mediterranean-type ecosystem in Portugal. Microbial Ecology 44: 344–353. 

Islam M, Hasin F (2000). Studies on phylloplane mycoflora of Amaranthus viridis L. 

National Academy Science Letters, India 23: 121–123. 

Jager ES de, Wehner FC, Karsten L (2001). Microbial ecology of the mango 

phylloplane. Microbial Ecology 42: 201–207. 

Kirk PM (1982). New or interesting microfungi. IV. Dematiaceous hyphomycetes 

from Devon. Transactions of the Britisch Mycological Society 78: 55–74. 

Levetin E, Dorsey K (2006). Contribution to leaf surface fungi to the air spora. 

Aerobiologia 22: 3–12. 

Masclaux F, Guého E, Hoog GS de, Christen R (1995). Phylogenetic relationship of 

human-pathogenic Cladosporium (Xylohypha) species. Journal of Medical and 

Veterinary Mycology 33: 327–338. 

Matsushima T (1975). Icones microfungorum a Matsushima lectorum. Kobe. 

McGinnis MR, Padhye AA (1978). Cladosporium castellanii is a synonym of Stenella 

araguata. Mycotaxon 7: 415–418. 




Minoura K (1966). Taxonomic studies on cladosporia (IV). Morphological properties 

(part II). Journal of Fermentation Technology 44: 137–149. 




Park HG, Managbanag JR, Stamenova EK, Jong SC (2004). Comparative analysis 

of common indoor Cladosporium species based on molecular data and conidial 

characters. Mycotaxon 89: 441–451. 





Kew. 



98: 625–634. 

Riesen T, Sieber T (1985). Endophytic fungi in winter wheat (Triticum aestivum L.). 

Swiss Federal Institute of Technology, Zürich. 

Roquebert MF (1981). Analyse des phénoménes pariétaux au cours de la 

conidiogenése chez quelques champignon microscopiques. Memoires du 

Museum d’Histoire Naturelle, Sér B, Botanique 28: 3–79. 



Mycologia 98: 1041–1052. 

Schubert K (2005a). Morphotaxonomic revision of foliicolous Cladosporium species 

(hyphomycetes). PhD thesis, Martin-Luther-University, Halle. 

Schubert K (2005b). Taxonomic revision of the genus Cladosporium s. lat. 3. A 

revision of Cladosporium species described by J.J. Davis and H.C. Greene 

(WIS). Mycotaxon 92: 55–76. 


Morphotaxonomic examination of Cladosporium species occurring on hosts of 

the families Bignoniaceae and Orchidaceae. Sydowia 56: 76–97. 

Schubert K, Braun U (2005a). Taxonomic revision of the genus Cladosporium 

s. lat. 1. Species reallocated to Fusicladium, Parastenella, Passalora, 

Pseudocercospora and Stenella. Mycological Progress 4: 101–109. 

Schubert K, Braun U (2005b). Taxonomic revision of the genus Cladosporium s. 

lat. 4. Species reallocated to Asperisporium, Dischloridium, Fusicladium, 

Passalora, Pseudasperisporium and Stenella. Fungal Diversity 20: 187–208. 

Schubert K, Braun U (2007). Taxonomic revision of the genus Cladosporium s. lat. 

6. New species, reallocations to and synonyms of Cercospora, Fusicladium, 

Passalora, Septonema and Stenella. Nova Hedwigia 84: 189–208. 

Schubert K, Braun U, Groenewald JZ, Crous PW (2007a). Cladosporium leaf-blotch 

and stem rot of Paeonia spp. caused by Dichocladosporium chlorocephalum 

gen. nov. Studies in Mycology 58: 95–104. 

Schubert K, Braun U, Mułenko W (2006). Taxonomic revision of the genus 

Cladosporium s. lat. 5. Validations and descriptions of new species. 



Hoog GS de, Crous PW (2007b). Biodiversity in the Cladosporium herbarum 





Seifert K, Nickerson NL, Corlett M, Jackson ED, Lois-Seize G, Davies RJ (2004). 

Devriesia, a new hyphomycete genus to accommodate heat-resistant, 


Stohr SN, Dighton J (2004). Effects of species diversity on establishment and 

coexistence: A phylloplane fungal community model system. Microbial Ecology 

48: 431–438. 


Capronia with notes on anamorph-teleomorph connections. Mycologia 89: 

120–131. 

Untereiner WA, Gerrits van den Ende AHG, Hoog GS de (1999). Nutritional 

physiology of species of Capronia. Studies in Mycology 43: 98–106. 

Untereiner WA, Naveau FA (1999). Molecular systematics of the Herpotrichiellaceae 

with an assessment of the phylogenetic positions of Exophiala dermatitidis and 


Varghese KIM, Rao VG (1979). Forest microfungi – I. Subramaniomyces, a new 

genus of Hyphomycetes. Kavaka 7: 83–85. 




Vries GA de (1952). Contribution to the Knowledge of the Genus Cladosporium Link 

ex Fr. Centraalbureau voor Schimmelcultures, Baarn. 





Wirsel SGR, Runge-Froböse C, Ahrén DG, Kemen E, Oliver RP, Mendgen 

KW (2002). Four or more species of Cladosporium sympatrically colonize 

Phragmites australis. Fungal Genetics and Biology 35: 99–113. 

56


doi:10.3114/sim.2007.58.03 


Phylogenetic and morphotaxonomic revision of Ramichloridium and allied 

genera 

M. Arzanlou 1,2 , J.Z. Groenewald 1 , W. Gams 1 , U. Braun 3 , H.-D Shin 4 and P.W. Crous 1,2* 

1 

CBS Fungal Biodiversity Centre, Uppsalalaan 8, 3584 CT Utrecht, The Netherlands; 2 Laboratory of Phytopathology, Wageningen University, Binnenhaven 5, 6709 PD 

Wageningen, The Netherlands; 3 Martin-Luther-Universität, Institut für Biologie, Geobotanik und Botanischer Garten, Neuwerk 21, D-06099 Halle (Saale), Germany; 4 Division of 

Environmental Science & Ecological Engineering, Korea University, Seoul 136-701, Korea 


Abstract: The phylogeny of the genera Periconiella, Ramichloridium, Rhinocladiella and Veronaea was explored by means of partial sequences of the 28S (LSU) rRNA 

gene and the ITS region (ITS1, 5.8S rDNA and ITS2). Based on the LSU sequence data, ramichloridium-like species segregate into eight distinct clusters. These include the 

Capnodiales (Mycosphaerellaceae and Teratosphaeriaceae), the Chaetothyriales (Herpotrichiellaceae), the Pleosporales, and five ascomycete clades with uncertain affinities. 

The type species of Ramichloridium, R. apiculatum, together with R. musae, R. biverticillatum, R. cerophilum, R. verrucosum, R. pini, and three new species isolated from 

Strelitzia, Musa and forest soil, respectively, reside in the Capnodiales clade. The human-pathogenic species R. mackenziei and R. basitonum, together with R. fasciculatum 

and R. anceps, cluster with Rhinocladiella (type species: Rh. atrovirens, Herpotrichiellaceae, Chaetothyriales), and are allocated to this genus. Veronaea botryosa, the type 

species of the genus Veronaea, also resides in the Chaetothyriales clade, whereas Veronaea simplex clusters as a sister taxon to the Venturiaceae (Pleosporales), and is placed 

in a new genus, Veronaeopsis. Ramichloridium obovoideum clusters with Carpoligna pleurothecii (anamorph: Pleurothecium sp., Chaetosphaeriales), and a new combination 

is proposed in Pleurothecium. Other ramichloridium-like clades include R. subulatum and R. epichloës (incertae sedis, Sordariomycetes), for which a new genus, Radulidium 

is erected. Ramichloridium schulzeri and its varieties are placed in a new genus, Myrmecridium (incertae sedis, Sordariomycetes). The genus Pseudovirgaria (incertae sedis) 

is introduced to accommodate ramichloridium-like isolates occurring on various species of rust fungi. A veronaea-like isolate from Bertia moriformis with phylogenetic affinity to 

the Annulatascaceae (Sordariomycetidae) is placed in a new genus, Rhodoveronaea. Besides Ramichloridium, Periconiella is also polyphyletic. Thysanorea is introduced to 

accommodate Periconiella papuana (Herpotrichiellaceae), which is unrelated to the type species, P. velutina (Mycosphaerellaceae). 

Taxonomic novelties: Myrmecridium Arzanlou, W. Gams & Crous, gen. nov., Myrmecridium flexuosum (de Hoog) Arzanlou, W. Gams & Crous, comb. et stat. nov., Myrmecridium 

schulzeri (Sacc.) Arzanlou, W. Gams & Crous var. schulzeri, comb. nov., Myrmecridium schulzeri var. tritici (M.B. Ellis) Arzanlou, W. Gams & Crous, comb. nov., Periconiella 

arcuata Arzanlou, S. Lee & Crous, sp. nov., Periconiella levispora Arzanlou, W. Gams & Crous, sp. nov., Pleurothecium obovoideum (Matsush.) Arzanlou & Crous, comb. nov., 

Pseudovirgaria H.D. Shin, U. Braun, Arzanlou & Crous, gen. nov., Pseudovirgaria hyperparasitica H.D. Shin, U. Braun, Arzanlou & Crous, sp. nov., Radulidium Arzanlou, W. 

Gams & Crous, gen. nov., Radulidium epichloës (Ellis & Dearn.) Arzanlou, W. Gams & Crous, comb. nov., Radulidium subulatum (de Hoog) Arzanlou, W. Gams & Crous, comb. 

nov., Ramichloridium australiense Arzanlou & Crous, sp. nov., Ramichloridium biverticillatum Arzanlou & Crous, nom. nov., Ramichloridium brasilianum Arzanlou & Crous, sp. 

nov., Ramichloridium strelitziae Arzanlou, W. Gams & Crous, sp. nov., Rhinocladiella basitona (de Hoog) Arzanlou & Crous, comb. nov., Rhinocladiella fasciculata (V. Rao & 

de Hoog) Arzanlou & Crous, comb. nov., Rhinocladiella mackenziei (C.K. Campb. & Al-Hedaithy) Arzanlou & Crous, comb. nov., Rhodoveronaea Arzanlou, W. Gams & Crous, 

gen. nov., Rhodoveronaea varioseptata Arzanlou, W. Gams & Crous, sp. nov., Thysanorea Arzanlou, W. Gams & Crous, gen. nov., Thysanorea papuana (Aptroot) Arzanlou, W. 

Gams & Crous, comb. nov., Veronaea japonica Arzanlou, W. Gams & Crous, sp. nov., Veronaeopsis Arzanlou & Crous, gen. nov., Veronaeopsis simplex (Papendorf) Arzanlou 

& Crous, comb.nov. 

Key words: Capnodiales, Chaetothyriales, Mycosphaerella, Periconiella, phylogeny, Rhinocladiella, Veronaea. 

Introduction 

The anamorph genus Ramichloridium Stahel ex de Hoog 1977 

presently accommodates a wide range of species with erect, dark, 

more or less differentiated, branched or unbranched conidiophores 

and predominantly aseptate conidia produced on a sympodially 

proliferating rachis (de Hoog 1977). This heterogeneous group 

of anamorphic fungi includes species with diverse life styles, viz. 

saprobes, human and plant pathogens, most of which were classified 

by Schol-Schwarz (1968) in Rhinocladiella Nannf. according to a 

very broad generic concept. Ramichloridium was originally erected 

by Stahel (1937) with R. musae Stahel as type species. However, 

because his publication lacked a Latin diagnosis, the genus was 

invalid. Stahel also invalidly described Chloridium musae Stahel for 

a fungus causing leaf spots (tropical speckle disease) on banana. 

Ellis (1976) validated Chloridium musae as Veronaea musae M.B. 

Ellis, and Ramichloridium musae as Periconiella musae Stahel ex 

M.B. Ellis. 

Periconiella Sacc. (1885) [type species P. velutina (G. Winter) 

Sacc.] differs from Veronaea Cif. & Montemart. chiefly based 

on its dark brown, apically branched conidiophores. However, 

de Hoog (1977) observed numerous specimens of V. musae to 

exhibit branched conidiophores in culture, as did Stahel (1937) 

for Ramichloridium musae. De Hoog (1977) subsequently reintroduced 

Ramichloridium, but typified it with R. apiculatum (J.H. 

Mill., Giddens & A.A. Foster) de Hoog. He regarded V. musae and 

P. musae to be conspecific, and applied the name R. musae (Stahel 

ex M.B. Ellis) de Hoog to both species, regarding Periconiella 

musae as basionym. The circumscription by de Hoog was based 

on their similar morphology and ecology. Central in his genus 

concept was the observed presence of more or less differentiated 

and pigmented conidiophores, with predominantly aseptate conidia 

produced on a sympodially proliferating rachis. De Hoog (1977) 

also used some ecological features as additional characters to 

discriminate Ramichloridium from other genera, noting, for instance, 

that species in Ramichloridium were non-pathogenic to humans 

(de Hoog 1977, Campbell & Al-Hedaithy 1993). This delimitation, 

however, was not commonly accepted (McGinnis & Schell 1980). 

De Hoog et al. (1983) further discussed the problematic separation 

of Ramichloridium from genera such as Rhinocladiella, Veronaea 

and Cladosporium Link. It was further noted that the main feature to 

distinguish Ramichloridium from Rhinocladiella, was the presence 

of exophiala-type budding cells in species of Rhinocladiella (de 

Hoog 1977, de Hoog et al. 1983, Veerkamp & Gams 1983). The 

separation of Veronaea from this complex is more problematic, as 

the circumscriptions provided by Ellis (1976) and Morgan-Jones 

(1979, 1982) overlap with that of Ramichloridium sensu de Hoog 

(1977). Cladosporium is more distinct, having very conspicuous, 

protuberant, darkened and thickened, coronate conidial scars, and 

catenate conidia (David 1997, Braun et al. 2003, Schubert et al. 

2007 – this volume). 

57

Arzanlou et al. 

To date 26 species have been named in Ramichloridium; they 

not only differ in morphology, but also in life style. Ramichloridium 

mackenziei C.K. Campb. & Al-Hedaithy is a serious human 

pathogen, causing cerebral phaeohyphomycosis (Al-Hedaithy et al. 

1988, Campbell & Al-Hedaithy 1993), whereas R. musae causes 

tropical speckle disease on members of the Musaceae (Stahel 1937, 

Jones 2000). Another plant-pathogenic species, R. pini de Hoog & 

Rahman, causes a needle disease on Pinus contorta (de Hoog et 

al. 1983). Other clinically relevant species of Ramichloridium are R. 

basitonum de Hoog and occasionally R. schulzeri (Sacc.) de Hoog, 

while the remaining species tend to be common soil saprobes. 

No teleomorph has thus far been linked to species of 

Ramichloridium. The main question that remains is whether shared 

morphology among the species in this genus reflects common 

ancestry (Seifert 1993, Untereiner & Naveau 1999). To delineate 

anamorphic genera adequately, morphology and conidial ontogeny 

alone are no longer satisfactory (Crous et al. 2006a, b), and DNA 

data provide additional characters to help delineate species and 

genera (Taylor et al. 2000, Mostert et al. 2006, Zipfel et al. 2006). 

The aim of the present study was to integrate morphological 

and cultural features with DNA sequence data to resolve the 

species concepts and generic limits of the taxa currently placed in 

Periconiella, Ramichloridium, Rhinocladiella and Veronaea, and to 

resolve the status of several new cultures that were isolated during 

the course of this study. 

Materials and Methods 

Isolates 

Species names, substrates, geographical origins and GenBank 

accession numbers of the isolates included in this study are listed 

in Table 1. Fungal isolates are maintained in the culture collection 

of the Centraalbureau voor Schimmelcultures (CBS) in Utrecht, the 

Netherlands. 

DNA extraction, amplification and sequence analysis 

Genomic DNA was extracted from colonies grown on 2 % malt 

extract agar (MEA, Difco) (Gams et al. 2007) using the FastDNA kit 

(BIO101, Carlsbad, CA, U.S.A.). The primers ITS1 and ITS4 (White 

et al. 1990) were used to amplify the internal transcribed spacer 

region (ITS) of the nuclear ribosomal RNA operon, including: the 

3’ end of the 18S rRNA gene, the first internal transcribed spacer 

region (ITS1), the 5.8S rRNA gene, the second internal transcribed 

spacer region (ITS2) and the 5’ end of 28S rRNA gene. Part of 

the large subunit 28S rRNA (LSU) gene was amplified with primers 

LR0R (Rehner & Samuels 1994) and LR5 (Vilgalys & Hester 1990). 

The ITS region was sequenced only for those isolates for which 

these data were not available. The ITS analyses confirmed the 

proposed classification based on LSU analysis for each major 

clade and are not presented here in detail; but the sequences are 

deposited in GenBank where applicable. The PCR reaction was 

performed in a mixture with 0.5 units Taq polymerase (Bioline, 

London, U.K.), 1× PCR buffer, 0.5 mM MgCl 2 

, 0.2 mM of each 

dNTP, 5 pmol of each primer, approximately 10–15 ng of fungal 

genomic DNA, with the total volume adjusted to 25 µL with 

sterile water. Reactions were performed on a GeneAmp PCR 

System 9700 (Applied Biosystems, Foster City, CA) with cycling 

conditions consisting of 5 min at 96 °C for primary denaturation, 

followed by 36 cycles at 96 °C (30 s), 52 °C (30 s), and 72 °C 

(60 s), with a final 7 min extension step at 72 °C to complete the 

reaction. The amplicons were sequenced using BigDye Terminator 

v. 3.1 (Applied Biosystems, Foster City, CA) or DYEnamicET 

Terminator (Amersham Biosciences, Freiburg, Germany) Cycle 

Sequencing Kits and analysed on an ABI Prism 3700 (Applied 

Biosystems, Foster City, CA) under conditions recommended by 

the manufacturer. Newly generated sequences were subjected 

to a Blast search of the NCBI databases, sequences with high 

similarity were downloaded from GenBank and comparisons 

were made based on the alignment of the obtained sequences. 

Sequences from GenBank were also selected for similar taxa. The 

LSU tree was rooted using sequences of Athelia epiphylla Pers. 

and Paullicorticium ansatum Liberta as outgroups. Phylogenetic 

analysis was performed with PAUP (Phylogenetic Analysis Using 

Parsimony) v. 4.0b10 (Swofford 2003), using the neighbour-joining 

algorithm with the uncorrected (“p”), the Kimura 2-parameter and 

the HKY85 substitution models. Alignment gaps longer than 10 


the remaining gaps were treated as missing data. Any ties were 

broken randomly when encountered. Phylogenetic relationships 

were also inferred with the parsimony algorithm using the heuristic 

search option with simple taxon additions and tree bisection and 

reconstruction (TBR) as the branch-swapping algorithm; alignment 

gaps were treated as a fifth character state and all characters were 

unordered and of equal weight. Branches of zero length were 

collapsed and all multiple, equally parsimonious trees were saved. 

Only the first 5 000 equally most parsimonious trees were saved. 

Other measures calculated included tree length, consistency 

index, retention index and rescaled consistency index (TL, CI, RI 

and RC, respectively). The robustness of the obtained trees was 

evaluated by 1 000 bootstrap replications. Bayesian analysis was 

performed following the methods of Crous et al. (2006c). The best 

nucleotide substitution model was determined using MrModeltest 

v. 2.2 (Nylander 2004). MrBayes v. 3.1.2 (Ronquist & Huelsenbeck 

2003) was used to perform phylogenetic analyses, using a general 

time-reversible (GTR) substitution model with inverse gamma 

rates, dirichlet base frequencies and the temp value set to 0.5. New 

sequences were lodged with NCBI’s GenBank (Table 1) and the 

alignment and trees with TreeBASE (www.treebase.org). 

Morphology 

Cultural growth rates and morphology were recorded from colonies 

grown on MEA for 2 wk at 24 ºC in the dark, and colony colours 

were determined by reference to the colour charts of Rayner (1970). 

Microscopic observations were made from colonies cultivated on 

MEA and OA (oatmeal agar, Gams et al. 2007), using a slide culture 

technique. Slide cultures were set up in Petri dishes containing 2 

mL of sterile water, into which a U-shaped glass rod was placed, 

extending above the water surface. A block of freshly growing 

fungal colony, approx. 1 cm square was placed onto a sterile 

microscope slide, covered with a somewhat larger, sterile glass 

cover slip, and incubated in the moist chamber. Fungal sporulation 

was monitored over time, and when optimal, images were captured 

by means of a Nikon camera system (Digital Sight DS-5M, Nikon 

Corporation, Japan). Structures were mounted in lactic acid, and 

30 measurements (× 1 000 magnification) determined wherever 

possible, with the extremes of spore measurements given in 

parentheses. 

58

Ramichloridium and allied genera 

Table 1. Isolates of Ramichloridium and similar genera used for DNA analysis and morphological studies. 

Species Accession number 1 Source Origin GenBank numbers 

(LSU, ITS) 

Myrmecridium flexuosum CBS 398.76*; IMI 203547 Soil Suriname EU041825, EU041768 

Myrmecridium schulzeri CBS 100.54; JCM 6974 Soil Zaire EU041826, EU041769 

CBS 134.68; ATCC 16310 Soil Germany EU041827, EU041770 

CBS 156.63 Homo sapiens Netherlands EU041828, EU041771 

CBS 188.96 Soil Papua New Guinea EU041829, EU041772 

CBS 304.73 Wheat straw South Africa EU041830, EU041773 

CBS 305.73; JCM 6967 Wheat straw South Africa EU041831, EU041774 

CBS 325.74; JCM 7234 Triticum aestivum Netherlands EU041832, EU041775 

CBS 381.87 — Australia EU041833, EU041776 

CBS 642.76 Malus sylvestris Switzerland EU041834, EU041777 

CBS 114996 Cannomois virgata South Africa EU041835, EU041778 

Periconiella arcuata CBS 113477* Ischyrolepsis subverticellata South Africa EU041836, EU041779 

Periconiella levispora CBS 873.73* Turpinia pomifera Sri Lanka EU041837, EU041780 

Periconiella velutina CBS 101948*; CPC 2262 Brabejum stellatifolium South Africa EU041838, EU041781 

CBS 101949; CPC 2263 Brabejum stellatifolium South Africa EU041839, EU041782 

CBS 101950; CPC 2264 Brabejum stellatifolium South Africa EU041840, EU041783 

Pleurothecium obovoideum CBS 209.95*; MFC 12477 Pasania edulis Japan EU041841, EU041784 

Pseudovirgaria hyperparasitica CBS 121735; CPC 10702 On Phragmidium sp. on Rubus coreanus Korea EU041822, EU041765 

CBS 121738; CPC 10704 On Phragmidium sp. on Rubus coreanus Korea EU041823, EU041766 

CBS 121739*; CPC 10753 

On Pucciniastrum agrimoniae on 

Korea 

EU041824, EU041767 

Agrimonia pilosa 

Radulidium epichloës CBS 361.63*; MUCL 3124 Epichloë typhina U.S.A. EU041842, EU041785 

Radulidium sp. CBS 115704 Poaceae Guyana EU041843, EU041786 

Radulidium subulatum CBS 287.84 Puccinia allii U.K. EU041844, EU041787 

Ramichloridium apiculatum 

CBS 405.76* Phragmites australis Czech Republic EU041845, EU041788 

CBS 912.96 Incubator for cell cultures Germany EU041846, EU041789 

CBS 101010 Lasioptera arundinis Czech Republic EU041847, EU041790 

CBS 156.59*; ATCC 13211; IMI Forest soil U.S.A. EU041848, EU041791 

100716; JCM 6972; MUCL 7991; 

MUCL 15753; QM 7716 

CBS 390.67 Cucumis sativus South Africa EU041849, EU041792 

CBS 391.67; JCM 6966 Aloe sp. South Africa EU041850, EU041793 

CBS 400.76; IMI 088021 Soil Pakistan EU041851, EU041794 

Ramichloridium australiense CBS 121710 Musa banksii Australia EU041852, EU041795 

Ramichloridium biverticillatum CBS 335.36 Musa sapientum — EU041853, EU041796 

Ramichloridium brasilianum CBS 283.92* Forest soil Brazil EU041854, EU041797 

Ramichloridium cerophilum CBS 103.59* Sasa sp. Japan EU041855, EU041798 

Ramichloridium indicum CBS 171.96 — — EU041856, EU041799 

Ramichloridium musae CBS 190.63; MUCL 9557 Musa sapientum — EU041857, EU041800 

CBS 365.36*; JCM 6973; MUCL 9556 Musa sapientum Surinam EU041858, EU041801 

Ramichloridium pini CBS 461.82*; MUCL 28942 Pinus contorta U.K. EU041859, EU041802 

Ramichloridium strelitziae CBS 121711 Strelitzia sp. South Africa EU041860, EU041803 

Rhinocladiella anceps 

CBS 157.54; ATCC 15680; MUCL Fagus sylvatica France EU041861, EU041804 

1081; MUCL 7992; MUCL 15756 

CBS 181.65*; ATCC 18655; DAOM Soil Canada EU041862, EU041805 

84422; IMI 134453; MUCL 8233; OAC 

10215 

Rhinocladiella basitona CBS 101460*; IFM 47593 Homo sapiens Japan EU041863, EU041806 

Rhinocladiella fasciculata CBS 132.86* Decayed wood India EU041864, EU041807 

Rhinocladiella mackenziei CBS 367.92; NCPF 2738; UTMB 3169 Homo sapiens Israel EU041865, EU041808 


59



Species Accession number 1 Source Origin GenBank numbers 

(LSU, ITS) 

CBS 368.92; UTMB 3170 Homo sapiens Israel EU041866, EU041809 

CBS 102590; NCPF 2853 Homo sapiens United Arab Emirates EU041867, EU041810 

Rhinocladiella phaeophora CBS 496.78*; IMI 287527 Soil Colombia EU041868, EU041811 

Rhinocladiella sp. CBS 264.49; MUCL 9904 Honey France EU041869, EU041812 

Rhodoveronaea varioseptata CBS 431.88* Bertia moriformis Germany EU041870, EU041813 

Thysanorea papuana CBS 212.96* — Papua New Guinea EU041871, EU041814 

Veronaea botryosa CBS 121.92 Xanthorrhoea preissii Australia EU041872, EU041815 

CBS 254.57*; IMI 070233; MUCL 9821 — Italy EU041873, EU041816 

CBS 350.65; IMI 115127; MUCL 7972 Goat dung India EU041874, EU041817 

Veronaea compacta CBS 268.75* — South Africa EU041876, EU041819 

Veronaea japonica CBS 776.83* On dead bamboo culm Japan EU041875, EU041818 

Veronaeopsis simplex CBS 588.66*; IMI 203547 Acacia karroo South Africa EU041877, EU041820 

Zasmidium cellare CBS 146.36 Wine cellar — EU041878, EU041821 

1 

ATCC: American Type Culture Collection, Virginia, U.S.A.; CBS: Centraalbureau voor Schimmelcultures, Utrecht, The Netherlands; CPC: Culture collection of Pedro Crous, 

housed at CBS; DAOM: Plant Research Institute, Department of Agriculture (Mycology), Ottawa, Canada; IFM: Research Center for Pathogenic Fungi and Microbial Toxicoses, 

Chiba University, Chiba, Japan; IMI: International Mycological Institute, CABI-Bioscience, U.K.; JCM: Japan Collection of Microorganism, RIKEN BioResource Center, Japan; 

MFC: Matsushima Fungus Collection, Kobe, Japan; MUCL: Mycotheque de l’ Université Catholique de Louvain, Louvain-la-Neuve, Belgium; NCPF: The National Collection of 

Pathogenic Fungi, Holborn, London, U.K.; OAC: Department of Botany and Genetics, University of Guelph, Ont., Canada; QM: Quartermaster Research and Developement 

Center, U.S. Army, MA, U.S.A.; UTMB: University of Texas Medical Branch, Texas, U.S.A. 


Results 

Phylogeny 

The manually adjusted alignment of the 28S rDNA data contained 

137 sequences (including the two outgroups) and 995 characters 

including alignment gaps. Of the 748 characters used in the 

phylogenetic analysis, 373 were parsimony-informative, 61 were 

variable and parsimony-uninformative, and 314 were constant. 

Neighbour-joining analysis using the three substitution models on 

the LSU alignment yielded trees with similar topology and bootstrap 

values. Parsimony analysis of the alignment yielded 5 000 equally 

most parsimonious trees, one of which is shown in Fig. 1 (TL = 

2 157, CI = 0.377, RI = 0.875, RC = 0.330). The Markov Chain 

Monte Carlo (MCMC) analysis of four chains started from a random 

tree topology and lasted 2 000 000 generations. Trees were saved 

each 1 000 generations, resulting in 2 000 trees. Burn-in was set 

at 500 000 generations after which the likelihood values were 

stationary, leaving 1 500 trees from which the consensus tree (Fig. 

2) and posterior probabilities (PP’s) were calculated. The average 

standard deviation of split frequencies was 0.043910 at the end 

of the run. Among the neighbour-joining, Bayesian and parsimony 

analyses, the trees differed in the hierarchical order of the main 

families and the support values (data not shown; e.g. the support 

within of the Capnodiales in Figs 1–2). 

The phylogenetic trees (Figs 1–2) show that the Ramichloridium 

species segregate into eight distinct clades, residing in the 

Capnodiales (Mycosphaerellaceae and Teratosphaeriaceae), the 

Chaetothyriales (Herpotrichiellaceae), the Pleosporales, and five 

other clades of which the relationships remain to be elucidated. The 

type species of Ramichloridium, R. apiculatum, together with R. 

musae, R. cerophilum (Tubaki) de Hoog, R. indicum (Subram.) de 

Hoog, R. pini and three new species respectively isolated from Musa 

banksii, Strelitzia nicolai, and forest soil, reside in different parts of 

the Capnodiales clade (all in the Mycosphaerellaceae, except for the 

species from forest soil which clusters in the Teratosphaeriaceae). 

The second clade (in the Chaetothyriomycetes clade), including 

the human-pathogenic species R. mackenziei and R. basitonum, 

together with R. fasciculatum V. Rao & de Hoog and R. anceps 

(Sacc. & Ellis) de Hoog, groups together with Rhinocladiella in the 

Herpotrichiellaceae. The third clade (in the Sordariomycetes clade) 

includes R. obovoideum (Matsush.) de Hoog, which in a Blast 

search was found to have affinity with Carpoligna pleurothecii F.A. 

Fernández & Huhndorf (Chaetosphaeriales). The fourth clade (in 

the Sordariomycetes clade) includes a veronaea-like isolate from 

Bertia moriformis, with phylogenetic affinity to the Annulatascaceae 

(Sordariomycetidae). The fifth clade (in the Sordariomycetes clade) 

includes R. schulzeri var. schulzeri and R. schulzeri var. flexuosum 

de Hoog, the closest relatives being Thyridium vestitum (Fr.) 

Fuckel in the Thyridiaceae and Magnaporthe grisea (T.T. Hebert) 

M.E. Barr in the Magnaporthaceae. The sixth clade (in the Incertae 

sedis clade) includes R. subulatum de Hoog, R. epichloës (Ellis & 

Dearn.) de Hoog and a species isolated from the Poaceae. Three 

ramichloridium-like isolates from Rubus coreanus and Agrimonia 

pilosa form another unique clade (in the Incertae sedis clade) 

with uncertain affinity. Veronaea simplex Papendorf clusters as 

sister taxon to the Venturiaceae representing the eighth clade 

(Dothideomycetes). The type species of Periconiella, P. velutina, 

clusters within the Mycosphaerellaceae (Capnodiales clade), 

whereas P. papuana Aptroot resides in the Herpotrichiellaceae 

(Chaetothyriales clade). Veronaea botryosa Cif. & Montemart., the 

type species of Veronaea, also resides in the Herpotrichiellaceae. 

Taxonomy 

The species previously described in Ramichloridium share 

some morphological features, including erect, pigmented, more 

or less differentiated conidiophores, sympodially proliferating 

conidiogenous cells and predominantly aseptate conidia. Other than 

conidial morphology, features of the conidiogenous apparatus that 

60


appear to be more phylogenetically informative include pigmentation 

of vegetative hyphae, conidiophores and conidia, denticle density 

on the rachis, and structure of the scars. By integrating these data 

with the molecular data set, more natural genera are delineated, 

which are discussed below. 

Key to ramichloridium-like genera 

1. Conidiogenous cells integrated, terminal and lateral on creeping or ascending hyphae (differentiation between branched vegetative 

hyphae and conidiophores barely possible); conidiogenous loci bulging, more or less umbonate, apex rounded; occurring on rust 

pustules ......................................................................................................................................................................... Pseudovirgaria 

1. Conidiogenous cells integrated in distinct conidiophores; conidiogenous loci non-umbonate (flat, not prominent; subcylindrical or conical 

denticles; or terminally flat-tipped; or thickened and darkened); rarely occurring on rust pustules, but if so, with a raduliform rachis and 

distinct denticles......................................................................................................................................................................................... 2 

2. Conidia 0–2(–3)-septate, conidial base truncate, retaining a marginal frill after liberation [anamorphs of Sordariomycetes] 

........................................................................................................................................................................................... Rhodoveronaea 

2. Conidial base without marginal frill ............................................................................................................................................................ 3 

3. Conidiophores composed of a well-developed erect stalk and a terminal branched head ........................................................................ 4 

3. Conidiophores unbranched or, if branched, branches loose, irregular or dichotomous, but not distinctly separated into stalk and branched 

head ........................................................................................................................................................................................................... 5 

4. Conidiophores dimorphic, either macronematous, dark brown with a dense apical cluster of branches or micronematous, undifferentiated, 

resembling vegetative hyphae; both kinds with a denticulate rachis; conidia predominantly 1-septate [anamorph of Chaetothyriales] 

.................................................................................................................................................................................................. Thysanorea 

4. Conidiophores monomorphic; branched head with fewer branches and looser; conidiogenous loci usually flat, non-prominent, less 

denticle-like; conidia aseptate to pluriseptate [anamorphs of Capnodiales] ............................................................................ Periconiella 

5. Rachis with denticles 1–1.5 µm long, denticles almost cylindrical; conidia at least partly in short chains ......................... Pleurothecium 

5. Rachis with denticles less than 1 µm long, denticles not cylindrical or denticles lacking, rachis with flat, barely prominent scars ........... 6 

6. Conidia predominantly septate .................................................................................................................................................................. 7 

6. Conidia predominantly aseptate ................................................................................................................................................................ 8 

7. Conidiophores up to 200 µm long; rachis straight, not to slightly geniculate; conidiogenous loci more or less flat, barely prominent, 

unthickened, slightly darkened [anamorphs of Chaetothyriales, Herpotrichiellaceae] ................................................................. Veronaea 

7. Conidiophores up to 60 µm long; rachis distinctly geniculate; conidiogenous loci denticle-like, prominent, up to 0.5 µm high, slightly 

thickened and darkened [anamorph of Pleosporales, Venturiaceae] ................................................................................... Veronaeopsis 

8. Vegetative mycelium entirely hyaline; rachis long, hyaline, with widely scattered pimple-shaped, terminally pointed, unpigmented 

denticles .............................................................................................................................................................................. Myrmecridium 

8. Vegetative mycelium at least partly pigmented; conidiogenous loci distinct, non-denticulate, somewhat darkened-refractive, or denticles, 

if present, neither pimple-shaped nor pointed ............................................................................................................................................ 9 

9. Rachis distinctly raduliform, with distinct, prominent blunt denticles, 0.5–1 µm long; scars and hila unthickened, but pigmented 

.................................................................................................................................................................................................. Radulidium 

9. Rachis not distinctly raduliform, at most subdenticulate; scars flat or only slightly prominent (subdenticulate), shorter ......................... 10 

10. Conidiophores usually poorly differentiated from the vegetative hyphae; conidial apparatus often loosely branched; exophiala-like 

budding cells usually present in culture [anamorphs of Chaetothyriales, Herpotrichiellaceae] .......................................... Rhinocladiella 

10. Conidiophores usually well differentiated from the vegetative mycelium (macronematous), usually unbranched; without exophiala-like 

states [anamorphs of Capnodiales] .................................................................................................................................. Ramichloridium 

Capnodiales (Mycosphaerellaceae, Teratosphaeriaceae) 

The type species of Ramichloridium, R. apiculatum, together 

with R. indicum cluster as a sister group to the Dissoconium de 

Hoog, Oorschot & Hijwegen clade in the Mycosphaerellaceae. 

Some other Ramichloridium species, including R. musae, R. 

biverticillatum Arzanlou & Crous, R. pini and R. cerophilum, are 

also allied with members of the Mycosphaerellaceae. Three 

additional new species are introduced for Ramichloridium isolates 


from Musa banksii, Strelitzia nicolai, and forest soil. Periconiella 

velutina, the type species of Periconiella, which also resides in the 

Mycosphaerellaceae, is morphologically sufficiently distinct to retain 

its generic status. Two new species of Periconiella are introduced 

for isolates obtained from Turpinia pomifera and Ischyrolepis 

subverticellata in South Africa. Zasmidium cellare (Pers.) Fr., the 

type species of Zasmidium (Pers.) Fr., is also shown to cluster 

within the Mycosphaerellaceae. 

61


10 changes 



100 Conioscyphascus varius AY484512 

100 

Conioscypha lignicola AY484513 

100 Carpoligna pleurothecii AF064645 

78 

Carpoligna pleurothecii AF064646 

Carpoligna pleurothecii AY544685 

98 

74 Pleurothecium obovoideum CBS 209.95 

Ascotaiwania hughesii AY316357 

100 Ascotaiwania hughesii AY094189 

Ophiostoma stenoceras DQ836904 

96 

Magnaporthe grisea AB026819 

100 Cryptadelphia polyseptata AY281102 

Cryptadelphia groenendalensis AY281103 

100 

100 82 

86 

96 

89 

Rhodoveronaea varioseptata CBS 431.88 

Annulatascus triseptatus AY780049 


Thyridium vestitum AY544671 

Capronia pulcherrima AF050256 

Exophiala dermatitidis AF050270 


Rhinocladiella sp. CBS 264.49 

100 

100 

Myrmecridium schulzeri CBS 114996 

Myrmecridium flexuosum CBS 398.76 

Myrmecridium schulzeri CBS 188.96 









Rhinocladiella mackenziei CBS 368.92 

Rhinocladiella mackenziei AF050288 


Rhinocladiella mackenziei CBS102590 

Capronia coronata AF050242 

Rhinocladiella fasciculata CBS 132.86 

Rhinocladiella phaeophora CBS 496.78 

Rhinocladiella anceps AF050284 

Rhinocladiella anceps CBS 181.65 




Veronaea botryosa CBS 350.65 



Thysanorea papuana CBS 212.96 

Veronaea japonica CBS 776.83 

Veronaea compacta CBS 268.75 

99 

80 60 

100 

95 

88 

Rhinocladiella basitona CBS 101460 

Rhinocladiella atrovirens AF050289 


Veronaeopsis simplex CBS 588.66 

Repetophragma goidanichii DQ408574 

Venturia pyrina EF114715 

Metacoleroa dickiei DQ384100 

Venturia chlorospora DQ384101 

Venturia inaequalis EF114713 

Venturia hanliniana AF050290 

54 

Venturia asperata EF114711 

Venturia carpophila AY849967 

Sordariomycetes 




Venturiaceae, 

Pleosporales, 

Dothideomycetes 

Fig. 1. (Page 62–63). One of 5 000 equally most parsimonious trees obtained from a heuristic search with simple taxon additions of the LSU sequence alignment using PAUP v. 

4.0b10. The scale bar shows 10 changes; bootstrap support values from 1 000 replicates are shown at the nodes. Thickened lines indicate the strict consensus branches and extype 

sequences are printed in bold face. The tree was rooted to two sequences obtained from GenBank (Athelia epiphylla AY586633 and Paullicorticium ansatum AY586693). 

Periconiella Sacc., in Sacc. & Berlese, Atti Ist. Veneto Sci., Ser. 6, 

3: 727. 1885. 

In vitro: Colonies with entire margin; aerial mycelium rather 

compact, raised, velvety, olivaceous-grey; reverse olivaceousblack. 

Submerged hyphae verrucose, hyaline, thin-walled, 1–3 µm 

wide; aerial hyphae subhyaline, later becoming dark brown, thickwalled, 

smooth. Conidiophores arising vertically from creeping 

hyphae, straight or flexuose, up to 260 µm long, dark brown at 

the base, paler towards the apex, thick-walled; in the upper part 

bearing short branches. Conidiogenous cells terminally integrated, 

polyblastic, smooth or verrucose, subcylindrical, mostly not or barely 

geniculate-sinuous, variable in length, subhyaline, later becoming 

pale brown, fertile part as wide as the basal part, proliferating 

sympodially, sometimes becoming septate and forming a short, 

straight rachis with pigmented, slightly thickened and hardly 

prominent, more or less flat scars. Conidia solitary, occasionally in 

short chains, 0–multi-septate, subhyaline to rather pale olivaceous 

or olivaceous-brown, smooth to verrucose, globose, ellipsoidal to 

obovoid or obclavate, with a slightly darkened and thickened hilum; 

conidial secession schizolytic. 

Type species: P. velutina (G. Winter) Sacc., Miscell. mycol. 2: 17. 

1884. 

62


54 

10 changes 

Fig. 1. (Continued). 

52 

76 

100 100 

98 

94 

71 

100 

77 

99 

90 

98 

Radulidium sp. CBS 115704 

Pseudovirgaria hyperparasitica CPC 10702 



96 Radulidium subulatum CBS 287.84 

99 Radulidium subulatum CBS 912.96 

Radulidium epichloës CBS 361.63 

78 

Radulidium epichloës AF050287 

64 

Radulidium subulatum CBS 405.76 

Radulidium subulatum CBS 101010 

Sporidesmium pachyanthicola DQ408557 


Ramichloridium brasilianum CBS 283.92 





Teratosphaeria molleriana EU019292 

Teratosphaeria fibrillosa EU019282 

Catenulostroma macowanii EU019254 

67 

58 100 

100 

87 

100 

56 

52 

99 54 

99 

87 


Cibiessia dimorphospora EU019258 


Xenomeris juniperi EF114709 

Catenulostroma abietis EU019249 



69 

78 

64 


Cladosporium cladosporioides EU019262 

Cladosporium uredinicola EU019264 

Cladosporium bruhnei EU019261 

Cladosporium sphaerospermum EU019263 

Ramichloridium indicum CBS 171.96 

Dissoconium aciculare EU019266 

“Mycosphaerella” communis EU019267 

“Mycosphaerella” lateralis EU019268 

Ramichloridium apiculatum CBS 390.67 




100 

100 

95 

99 

56 

55 

80 

Ramichloridium pini CBS 461.82 

Mycosphaerella endophytica DQ246252 


Mycosphaerella gregaria DQ246251 

Mycosphaerella graminicola EU019297 

Septoria tritici EU019298 

Cercosporella centaureicola EU019257 


Ramularia sp. EU019285 


Ramularia pratensis var. pratensis EU019284 

Mycosphaerella walkeri DQ267574 

Mycosphaerella parkii DQ246245 

Mycosphaerella madeirae DQ204756 

Periconiella levispora CBS 873.73 

Mycosphaerella marksii DQ246249 

Pseudocercospora epispermogonia DQ204758 


Mycosphaerella intermedia DQ246248 


Periconiella arcuata CBS 113477 

Rasutoria pseudotsugae EF114704 

Rasutoria tsugae EF114705 

Periconiella velutina CBS 101950 



Ramichloridium biverticillatum CBS 335.36 

Ramichloridium musae CBS 365.36 


56 

56 

77 

66 

97 

Ramichloridium australiense CBS 121710 

Ramichloridium strelitziae CBS 121711 

Zasmidium cellare CBS 146.36 

Ramichloridium cerophilum CBS 103.59 

Ramichloridium cerophilum AF050286 






Notes: Periconiella is distinct from other ramichloridium-like genera 

by its conidiophores that are prominently branched in the upper 

part, and by its darkened, thickened conidial scars, that are more 

or less flat and non-prominent. Although conidiophores are also 

branched in the upper part in Thysanorea Arzanlou, W. Gams & 

Crous, the branching pattern in the latter genus is different from 

that of Periconiella. Thysanorea has a complex head consisting of 

up to six levels of branches, while in Periconiella the branching is 

limited, with mainly primary and secondary branches. Furthermore, 

Thysanorea is characterised by having dimorphic conidiophores 

and more or less prominent denticle-like conidiogenous loci. 


63




Veronaeopsis simplex CBS 588.66 

0.98 Repetophragma goidanichii DQ408574 


0.79 Venturia pyrina EF114715 

1.00 Venturia hanliniana AF050290 

Venturia inaequalis EF114713 

0.84 Venturia chlorospora DQ384101 

Venturia asperata EF114711 

1.00 


0.99 

0.52 

0.74 

1.00 

1.00 

0.97 

1.00 

0.80 

0.63 

0.99 

0.50 

0.68 

0.69 

1.00 

0.58 




Rhinocladiella phaeophora CBS 496.78 

Rhinocladiella fasciculata CBS 132.86 

1.00 




Rhinocladiella mackenziei AF050288 


Rhinocladiella mackenziei CBS 102590 

1.00 





Rhinocladiella sp. CBS 264.49 


Rhinocladiella atrovirens AF050289 

Rhinocladiella basitona CBS 101460 


Thysanorea papuana CBS 212.96 

Veronaea japonica CBS 776.83 

Veronaea compacta CBS 268.75 




0.84 

0.99 

0.96 

0.99 

0.91 

1.00 

1.00 Conioscyphascus varius AY484512 

Conioscypha lignicola AY484513 

1.00 Carpoligna pleurothecii AF064646 

1.00 

Carpoligna pleurothecii AF064645 

Carpoligna pleurothecii AY544685 

0.99 1.00 Pleurothecium obovoideum CBS 209.95 

1.00 Ascotaiwania hughesii AY316357 

Ascotaiwania hughesii AY094189 

Magnaporthe grisea AB026819 

Ophiostoma stenoceras DQ836904 

Cryptadelphia polyseptata AY281102 

Cryptadelphia groenendalensis AY281103 

1.00 

1.00 

1.00 

0.61 

0.87 

0.71 

0.92 

1.00 

Venturiaceae, 

Pleosporales, 

Dothideomycetes 

Rhodoveronaea varioseptata CBS 431.88 



Thyridium vestitum AY544671 

Myrmecridium schulzeri CBS 114996 





Myrmecridium flexuosum CBS 398.76 









Sordariomycetes 

Fig. 2. (Page 64–65). Consensus phylogram (50 % majority rule) of 1 500 trees resulting from a Bayesian analysis of the LSU sequence alignment using MrBayes v. 3.1.2. 

Bayesian posterior probabilities are indicated at the nodes. Ex-type sequences are printed in bold face. The tree was rooted to two sequences obtained from GenBank (Athelia 

epiphylla AY586633 and Paullicorticium ansatum AY586693). 

Periconiella velutina (G. Winter) Sacc., Miscell. mycol. 2: 17. 

1884. Fig. 3. 

Basionym: Periconia velutina G. Winter, Hedwigia 23: 174. 1884. 

In vitro: Submerged hyphae verrucose, hyaline, thin-walled, 1–3 

µm wide; aerial hyphae subhyaline, later becoming dark brown, 

thick-walled, smooth. Conidiophores arising vertically from creeping 

hyphae, straight or flexuose, up to 260 µm long, dark brown at the 

base, paler towards the apex, thick-walled; in the upper part bearing 

short branches, 10–35 µm long. Conidiogenous cells mostly 

terminally integrated, sometimes discrete, smooth or verrucose, 

cylindrical, variable in length, subhyaline, later becoming pale 

brown, fertile part as wide as the basal part, proliferating sympodially, 

sometimes becoming septate and forming a short, straight rachis 

with pigmented, slightly thickened and hardly prominent, more 

or less flat scars, less than 1 µm diam. Conidia 0(–1)-septate, 

subhyaline, thin-walled, verrucose or smooth, globose, ellipsoidal 

to obovoid, (7–)8–9(–11) × (2.5–)3(–4) µm, with a slightly darkened 

and thickened hilum, 1.5–2 µm diam. 

Cultural characteristics: Colonies on MEA slow-growing, reaching 

4 mm diam after 14 d at 24 °C, with entire margin; aerial mycelium 

rather compact, raised, velvety, olivaceous-grey; reverse 

olivaceous-black. 

Specimens examined: South Africa, Cape Town, on Brabejum stellatifolium, P. 

MacOwan, herb. G. Winter (B), lectotype selected here; Cape Town, on leaves 

of Brabejum stellatifolium (= B. stellatum), P. Mac-Owan, PAD, F42165, F462166, 

isolectotypes; Stellenbosch, Jonkershoek Nature Reserve, on Brabejum 

stellatifolium, 21 Jan. 1999, J.E. Taylor, epitype designated here CBS H-15612, 

cultures ex-epitype CBS 101948–101950. 

64




0.64 

0.61 

0.96 

0.99 

0.87 

1.00 

1.00 

1.00 

0.68 

0.99 

1.00 

1.00 

1.00 

1.00 

0.99 

0.86 




Radulidium sp. CBS 115704 

0.95 Radulidium subulatum CBS 287.84 


Radulidium epichloës CBS 361.63 

Radulidium epichloës AF050287 


Radulidium subulatum CBS 101010 

0.60 

0.57 

Cladosporium cladosporioides EU019262 

0.91 

Cladosporium uredinicola EU019264 

1.00 Cladosporium bruhnei EU019261 

0.81 Cladosporium sphaerospermum EU019263 

1.00 Sporidesmium pachyanthicola DQ408557 


1.00 Ramichloridium brasilianum CBS 283.92 

1.00 Batcheloromyces proteae EU019247 




Teratosphaeria molleriana EU019292 

Teratosphaeria fibrillosa EU019282 

Catenulostroma macowanii EU019254 

0.63 

0.73 

0.72 

0.95 

1.00 

1.00 

1.00 

1.00 

0.64 

1.00 

1.00 


Cibiessia dimorphospora EU019258 





Xenomeris juniperi EF114709 

Catenulostroma abietis EU019249 

Ramichloridium indicum CBS 171.96 

Dissoconium aciculare EU019266 

“Mycosphaerella” communis EU019267 

“Mycosphaerella” lateralis EU019268 





0.99 

0.72 

1.00 



Mycosphaerella gregaria DQ246251 

Cercosporella centaureicola EU019257 


Ramularia sp. EU019285 


Ramularia pratensis var. pratensis EU019284 

Mycosphaerella graminicola EU019297 

Septoria tritici EU019298 

Ramichloridium pini CBS 461.82 

1.00 Mycosphaerella walkeri DQ267574 

Mycosphaerella parkii DQ246245 

Mycosphaerella madeirae DQ204756 

0.95 Periconiella levispora CBS 873.73 

0.76 

0.60 

0.97 

0.64 

0.80 

1.00 

0.90 

0.95 

0.97 

1.00 

0.64 

0.99 

0.94 

0.97 

0.57 




Mycosphaerella intermedia DQ246248 


Periconiella arcuata CBS 113477 

Rasutoria pseudotsugae EF114704 

Rasutoria tsugae EF114705 




0.96 

0.97 

0.97 

0.96 

Ramichloridium biverticillatum CBS 335.36 



Ramichloridium australiense CBS 121710 

Ramichloridium strelitziae CBS 121711 

Zasmidium cellare CBS 146.36 

Ramichloridium cerophilum CBS 103.59 







Periconiella arcuata Arzanlou, S. Lee & Crous, sp. nov. MycoBank 

MB504547. Figs 4, 7A. 

Etymology: Named after its curved conidia. 

Ab aliis speciebus Periconiellae conidiis obclavatis, rectis vel curvatis, (30–)53–61(– 

79) × (3–)5(–7) µm, distinguenda. 

Submerged hyphae smooth, hyaline, thin-walled, 2 µm wide; 

aerial hyphae pale brown, smooth or verrucose, slightly narrower. 

Conidiophores arising vertically from creeping hyphae, straight 

or flexuose, up to 300 µm long, dark brown at the base, paler 


towards the apex, thick-walled; loosely branched in the upper part, 

bearing short branches. Conidiogenous cells integrated, cylindrical, 

variable in length, 20–50 µm long, subhyaline, later becoming 

pale brown, fertile part as wide as the basal part, proliferating 

sympodially, forming a geniculate conidium-bearing rachis with 

pigmented and thickened, prominent, cone-shaped scars, 1 µm 

diam. Conidia formed singly, obclavate, straight or mostly curved, 

0(–4)-septate, coarsely verrucose, pale olive, thin-walled, tapering 

towards the apex, (30–)53–61(–79) × (3–)5(–7) µm, with a narrowly 

truncate base and a darkened, hardly thickened hilum, 2 µm diam; 

microcyclic conidiation observed in culture. 

65


Fig. 3. Periconiella velutina (CBS 101948). A–B. Macronematous conidiophores with short branches in the upper part. C. Sympodially proliferating conidiogenous cell with 

darkened and slightly thickened scars. D. Conidia. Scale bar = 10 µm. 

Fig. 4. Periconiella arcuata (CBS 113477). A–B. Sympodially proliferating conidiogenous cells with darkened, thickened and cone-shaped scars. C–E. Macronematous 

conidiophores with loose branches in the upper part. F–I. Conidia. Scale bar = 10 µm. 

66


Fig. 5. Periconiella levispora (CBS 873.73). A–C. Conidial apparatus at different stages of development, which gives rise to macronematous conidiophores with dense branches 

in the upper part. D. Sympodially proliferating conidiogenous cells with darkened and somewhat protruding scars. E–F. Conidia with truncate base and darkened hilum. Scale 

bar = 10 µm. 

Fig. 6. A. Pseudovirgaria hyperparasitica (CBS 121739 = CPC 10753). B. Periconiella levispora (CBS 873.73). Scale bar = 10 µm. 


67



after 14 d at 24 °C, with entire, smooth, sharp margin; mycelium 

compacted, becoming hairy, colonies up to 1 mm high; surface 

olivaceous to olivaceous-grey, reverse dark grey-olivaceous to 

olivaceous-black. 

Specimen examined: South Africa, Western Cape Province, Kogelberg, on dead 

culms of Ischyrolepis subverticillata, May 2001, S. Lee, holotype CBS H-19927, 

culture ex-type CBS 113477. 

Periconiella levispora Arzanlou, W. Gams & Crous, sp. nov. 

MycoBank MB504546. Figs 5–6B. 

Etymology: (Latin) levis = smooth. 

A simili Periconiella velutina conidiis levibus et maioribus, ad 23 μm longis 

distinguenda. 

In vitro: Submerged hyphae smooth, hyaline, thin-walled, 2–2.5 µm 

wide; aerial hyphae subhyaline, later becoming dark brown, thickwalled, 

smooth. Conidiophores arising vertically from creeping 

aerial hyphae, dark brown at the base, paler towards the apex, 

thick-walled; in the upper part bearing several short branches, 

up to 120 µm long. Conidiogenous cells integrated, occasionally 

discrete, cylindrical, variable in length, 10–20 µm long, subhyaline, 

later becoming pale brown, fertile part as wide as the basal part, 

proliferating sympodially, forming a short rachis with pigmented 

and slightly thickened, somewhat protruding scars, less than 1 

µm diam. Conidia solitary, 0(–2)-septate, smooth, pale olivaceous, 

cylindrical, ellipsoidal, pyriform to clavate, (7–)11–14(–23) × (3–)4– 

5(–6) µm, with a truncate base and a darkened, slightly thickened 

hilum, 2 µm diam. 



compact, raised, velvety, olivaceous-grey; reverse olivaceousblack. 

Basionym: Chloridium apiculatum J.H. Mill., Giddens & A.A. Foster, 

Mycologia 49: 789. 1957. 

≡ Veronaea apiculata (J.H. Mill., Giddens & A.A. Foster) M.B. Ellis, in Ellis, 

More Dematiaceous Hyphomycetes: 209. 1976. 

[non Rhinocladiella apiculata Matsush., in Matsushima, Icon. 

Microfung. Mats. lect.: 122. 1975]. 

= Rhinocladiella indica Agarwal, Lloydia 32: 388. 1969. 

[non Chloridium indicum Subram., Proc. Indian Acad. Sci., Sect. B, 

42: 286. 1955]. 

In vitro: Submerged hyphae hyaline to subhyaline, thin-walled, 

1–2.5 µm wide; aerial hyphae slightly darker, smooth-walled. 

Conidiophores generally arising at right angles from creeping aerial 

hyphae, straight, unbranched, thick-walled, dark brown, continuous 

or with 1–2(–3) additional thin septa, up to 100 µm long; intercalary 

cells 10–28 µm long. Conidiogenous cells integrated, terminal, 

smooth, thick-walled, golden-brown, straight, cylindrical, 25–37(– 

47) × 2–3.5 µm; proliferating sympodially, resulting in a straight 

rachis with conspicuous conidiogenous loci; scars prominent, 

crowded, slightly pigmented, less than 1 µm diam. Conidia solitary, 

obovate to obconical, pale brown, finely verrucose, (3–)5–5.5(–7.5) 

× (2–)2.5–3(–4) µm, hilum conspicuous, slightly pigmented, about 

1 µm diam. 


after 14 d at 24 °C; minimum temperature for growth above 6 °C, 

optimum 24 °C, maximum 30 °C. Colonies raised, velvety, dense, 

with entire margin; surface olivaceous-green, reverse olivaceousblack, 

often with a diffusing citron-yellow pigment. 

Specimens examined: Pakistan, Lahore, from soil, A. Kamal, CBS 400.76 = IMI 

088021. South Africa, from preserved Cucumis sativus in 8-oxyquinoline sulphate, 

M.C. Papendorf, CBS 390.67; Potchefstroom, from Aloe sp., M.C. Papendorf, CBS 

391.67. U.S.A., Georgia, isolated from forest soil, CBS 156.59 = ATCC 13211 = IMI 

100716 = QM 7716, ex-type culture. 

Specimen examined: Sri Lanka, Hakgala Botanic Gardens, on dead leaves of 

Turpinia pomifera, Jan. 1973, W. Gams, holotype CBS H-15611, culture ex-type 

CBS 873.73. 

Ramichloridium Stahel ex de Hoog, Stud. Mycol. 15: 59. 1977. 

In vitro: Colonies flat to raised, with entire margin; surface 

olivaceous-green to olivaceous-black. Mycelium consisting of 

submerged and aerial hyphae; submerged hyphae hyaline to 

subhyaline, thin-walled, aerial hyphae smooth or verrucose. 

Conidiophores straight, unbranched, rarely branched, thick-walled, 

dark brown (darker than the subtending hyphae), continuous or 

with several additional thin septa. Conidiogenous cells integrated, 

terminal, polyblastic, smooth, thick-walled, golden-brown, apical 

part subhyaline, with sympodial proliferation, straight or flexuose, 

geniculate or nodose, with conspicuous conidiogenous loci; 

scars crowded or scattered, unthickened, unpigmented to faintly 

pigmented, or slightly prominent denticles. Conidia solitary, 0–1- 

septate, subhyaline to pale brown, smooth to coarsely verrucose, 

rather thin-walled, obovate, obconical or globose to ellipsoidal, 

fusiform, with a somewhat prominent, slightly pigmented hilum; 


Type species: R. apiculatum (J.H. Mill., Giddens & A.A. Foster) de 

Hoog, Stud. Mycol. 15: 69. 1977. 

Ramichloridium apiculatum (J.H. Mill., Giddens & A.A. Foster) de 

Hoog, Stud. Mycol. 15: 69. 1977. Fig. 8. 

Fig. 7. A. Periconiella arcuata (CBS 113477). B. Myrmecridium schulzeri (CBS 

325.74). C. Thysanorea papuana (CBS 212.96). Scale bars = 10 µm. 

68


Fig. 8. Ramichloridium apiculatum (CBS 156.59). A–C. Macronematous conidiophores with sympodially proliferating conidiogenous cells, which give rise to a conidium-bearing 

rachis with crowded and prominent scars. D. Conidia. Scale bar = 10 µm. 

Fig. 9. Ramichloridium australiense (CBS 121710). A–C. Macronematous conidiophores with thick-walled and warted subtending hyphae. D. Sympodially proliferating 

conidiogenous cell, which results in a short rachis with darkened and slightly thickened scars. E. Conidia. Scale bar = 10 µm. 

Ramichloridium australiense Arzanlou & Crous, sp. nov. 

MycoBank MB504548. Figs 9–10A. 

Etymology: Named after its country of origin, Australia. 

Ab aliis speciebus Ramichloridii conidiophoris ex hyphis verrucosis, crassitunicatis 

ortis distinguendum. 

In vitro: Submerged hyphae hyaline, smooth, thin-walled, 1–2 µm 

wide; aerial hyphae pale brown, warted. Conidiophores arising 

vertically and clearly differentiated from creeping aerial hyphae, up 

to 400 µm tall, with several additional thin septa; intercalary cells, 

8–40 × 2–5 µm, from the broadest part at the base tapering towards 

the apex, subhyaline, later becoming pale brown and warted in the 

lower part. Subtending hyphae thick-walled, warted. Conidiogenous 

cells integrated, terminal, 10–18 µm long, proliferating sympodially, 


giving rise to a short rachis with conspicuous conidiogenous loci; 

scars slightly thickened and darkened, about 1 µm diam. Conidia 

solitary, aseptate, thin-walled, smooth, subhyaline, subcylindrical 

to obclavate, (10–)12–15(–23) × 2.5–3 µm, with a truncate base 

and a slightly darkened and thickened hilum,1.5–2 µm diam, rarely 

fusing at the basal part. 

Cultural characteristics: Colonies on MEA rather slow growing, 

reaching 8 mm diam after 14 d at 24 °C, with entire, smooth 

margin; mycelium flat, olivaceous-grey, becoming granular, with 

gelatinous droplets at the margin developing with aging; reverse 

pale olivaceous-grey. 

Specimen examined: Australia, Queensland, Mount Lewis, Mount Lewis Road, 

16°34’47.2” S, 145°19’7” E, 538 m alt., on Musa banksii leaf, Aug. 2006, P.W. Crous 

and B. Summerell, holotype CBS H-19928, culture ex-type CBS 121710. 

69


Fig. 10. A. Ramichloridium australiense (CBS 121710). B. Ramichloridium brasilianum (CBS 283.92). C. Radulidium subulatum (CBS 405.76). D. Rhodoveronaea varioseptata 

(CBS 431.88). Scale bar = 10 µm. 

Fig. 11. Ramichloridium musae (CBS 365.36). A. Conidiophores with loose branches. B–D. Sympodially proliferating conidiogenous cells, resulting in a long conidium-bearing 

rachis. E. Rachis with hardly prominent, slightly darkened scars. F. Conidia. Scale bars = 10 µm. 

70


Fig. 12. Ramichloridium biverticillatum (CBS 335.36). A–B. Profusely branched and biverticillate conidiophores. C. Sympodially proliferating conidiogenous cells, 

which give rise to a conidium-bearing rachis with crowded, slightly pigmented and thickened scars. D. Conidia. Scale bar = 10 µm. 

Fig. 13. Ramichloridium brasilianum (CBS 283.92). A–B. Macronematous conidiophores with sympodially proliferating conidiogenous cells, resulting in a conidiumbearing 

rachis. C. Rachis with crowded and slightly pigmented scars. D. Conidia. Scale bar = 10 µm. 

Fig. 14. Ramichloridium cerophilum (CBS 103.59). A–C. Conidial apparatus at different stages of development, resulting in macronematous conidiophores and 

sympodially proliferating conidiogenous cells. D–E. Formation of secondary conidia. F. Conidia. Scale bar = 10 µm. 


71


Ramichloridium musae (Stahel ex M.B. Ellis) de Hoog, Stud. 

Mycol. 15: 62. 1977. Fig. 11. 

Basionym: Veronaea musae Stahel ex M.B. Ellis, in Ellis, More 

Dematiaceous Hyphomycetes: 209. 1976. 

≡ Chloridium musae Stahel, Trop. Agric., Trinidad 14: 43. 1937 (nom. 

inval. Art. 36). 

Misapplied name: Chloridium indicum Subram., sensu Batista & 

Vital, Anais Soc. Biol. Pernambuco 15: 379. 1957. 

In vitro: Submerged hyphae smooth, hyaline, thin-walled, 1–2 µm 

wide; aerial hyphae subhyaline, smooth. Conidiophores arising 

vertically and mostly sharply differentiated from creeping aerial 

hyphae, golden-brown; unbranched, rarely branched in the upper 

part, up to 250 µm tall, with up to 6 additional thin septa, cells 

23–40 × 2–2.5 µm, basal cell occasionally inflated. Conidiogenous 

cells terminally integrated, cylindrical, variable in length, 10–40 µm 

long, golden-brown near the base, subhyaline to pale brown near 

the end, fertile part as wide as the basal part, later also becoming 

septate; rachis elongating sympodially, 2–2.5 µm wide, with hardly 

prominent, scattered, slightly pigmented scars, about 0.5 µm diam. 

Conidia solitary, aseptate, hyaline to subhyaline, ellipsoidal, (4–)7– 

8(–12) × 2–3 µm, smooth or verruculose, thin-walled, with slightly 

darkened hilum, about 1 µm diam. 


27 mm diam after 14 d at 24 °C, with entire, smooth, sharp margin; 

mycelium mostly submerged, some floccose to lanose aerial 

mycelium in the olivaceous-grey centre, becoming pale pinkish 

olivaceous towards the margin; reverse pale orange. 

Specimens examined: Cameroon, from Musa sapientum, J.E. Heron, CBS 169.61 

= ATCC 15681 = IMI 079492 = DAOM 84655 = MUCL 2689; from Musa sapientum, 

J. Brun, CBS 190.63 = MUCL 9557. Surinam, Paramaribo, from Musa sapientum 

leaf, G. Stahel, CBS 365.36 = JCM 6973 = MUCL 9556, ex-type strain of Chloridium 

musae; from Musa sapientum, G. Stahel, CBS 365.36; dried culture preserved as 

CBS H-19933. 

Ramichloridium biverticillatum Arzanlou & Crous, nom. nov. 


Basionym: Periconiella musae Stahel ex M.B. Ellis, Mycol. Pap. 

111: 5. 1967. 

[non Ramichloridium musae (Stahel ex M.B. Ellis) de Hoog, 1977]. 

≡ Ramichloridium musae Stahel, Trop. Agric., Trinidad 14: 43. 1937 (nom. 

inval. Art. 36). 

= Ramichloridium musae (Stahel ex M.B. Ellis) de Hoog, Stud. Mycol. 15: 62. 

1977, sensu de Hoog, p.p. 

Etymology: Named after its biverticillate conidiophores. 

In vitro: Submerged hyphae smooth, hyaline, thin-walled, 1–2 

µm wide; aerial hyphae subhyaline, smooth, slightly darker. 

Conidiophores arising vertically from creeping aerial hyphae, pale 

brown, profusely branched, biverticillate, with up to three levels 

of main branches; branches tapering distally, 2–3 µm wide at the 

base, approx. 2 µm wide in the upper part, up to 250 µm long. 

Conidiogenous cells terminally integrated, cylindrical, variable in 

length, 15–50 µm long, rachis straight or geniculate, pale brown, 

as wide as the basal part; elongating sympodially, forming a rachis 

with crowded, slightly darkened and thickened minute scars, less 

than 0.5 µm wide. Conidia solitary, aseptate, hyaline to subhyaline, 

dacryoid to pyriform, (2–)3–4(–6) × (1.5–)2(–2.5) µm, smooth, thinwalled, 

with an inconspicuous hilum. 


16 mm diam after 14 d at 24 °C, with entire, smooth, sharp margin, 

rather compact, velvety; surface vinaceous-buff to olivaceous-buff; 

reverse buff. 

Specimen examined: Surinam, from Musa sapientum, Aug. 1936, G. Stahel, CBS 

335.36. 

Notes: Ramichloridium biverticillatum is a new name based on 

Periconiella musae. The species is distinct from R. musae because 

of its profusely branched conidiophores, and conidia that are smaller 

(2–5 × 1.5–2.5 µm) than those of R. musae (5–11 × 2–3 µm). 

Ramichloridium brasilianum Arzanlou & Crous, sp. nov. 

MycoBank MB504550. Figs 10B, 13. 

Etymology: Named after its country of origin, Brazil. 

A simili Ramichloridio cerophilo conidiis minoribus, ad 8 μm longis, et conidiis 

secundariis absentibus distinguendum. 

In vitro: Submerged hyphae pale olivaceous, smooth or slightly 

rough, 1.5–2 µm wide; aerial hyphae olivaceous, smooth or rough, 

narrower and darker than the submerged hyphae. Conidiophores 

unbranched, arising vertically from creeping aerial hyphae, straight 

or flexuose, dark brown, with up to 10 additional septa, thick-walled, 

cylindrical, 2–2.5 µm wide and up to 70 µm long. Conidiogenous 

cells integrated, terminal, 10–30 µm long, proliferating sympodially, 

giving rise to a long, straight rachis with crowded, slightly darkened 

minute scars, about 0.5 µm diam. Conidia solitary, obovoid 

to fusiform with the widest part below the middle, thin-walled, 

verruculose, aseptate, pale brown, slightly rounded at the apex, 

truncate at the base, (4–)5–6(–8.5) × 2–2.5(–3) µm, with a slightly 

thickened and darkened hilum, 1–1.5 µm diam. 


6 mm diam after 14 d at 24 °C, velvety to hairy, colonies with entire 

margin, surface dark olivaceous-grey; black gelatinous exudate 

droplets produced on OA. 

Specimen examined: Brazil, São Paulo, Peruibe, Jureia Ecological Reserve, forest 

soil, Jan. 1991, D. Attili, holotype CBS H-19929, culture ex-type CBS 283.92. 

Ramichloridium cerophilum (Tubaki) de Hoog, Stud. Mycol. 15: 

74. 1977. Fig. 14. 

Basionym: Acrotheca cerophila Tubaki, J. Hattori Bot. Lab. 20: 143. 

1958. 

≡ Cladosporium cerophilum (Tubaki) Matsush., in Matsushima, Icon. 

Microfung. Matsush. lect. (Kobe): 34. 1975. 

In vitro: Submerged hyphae pale olivaceous-brown, smooth or 

slightly rough, 1.5–3 µm wide; aerial hyphae olivaceous-brown, 

smooth or slightly rough, somewhat narrower and darker than the 

submerged hyphae. Conidiophores unbranched, arising vertically 

from creeping aerial hyphae, dark brown, thick-walled, smooth or 

verruculose, hardly tapering towards the apex, 2–3 µm wide, up 

to 50 µm long, with up to 3 additional septa. Conidiogenous cells 

integrated, terminal, proliferating sympodially, rachis short and 

straight, with crowded, prominent, pigmented unthickened scars, 

minute, approx. 0.5 µm diam. Conidia solitary, fusiform to clavate, 

thin-walled, smooth, 0(–1)-septate, subhyaline, (4–)6–7(–11) × (2–) 

2.5(–3) µm, with a conspicuous hilum, about 0.5 µm diam, slightly 

raised with an inconspicuous marginal frill, somehow resembling 

those of Cladosporium. Conidia sometimes producing 1–3(–4) 

short secondary conidia. 

Cultural characteristics: Colonies on MEA rather slow-growing, 

reaching 12 mm diam after 14 d at 24 °C, velvety to hairy, with 

entire margin; surface dark olivaceous-grey, with black gelatinous 

exudate droplets on OA. 

Specimen examined: Japan, isolated from Sasa sp., K. Tubaki, CBS 103.59, extype. 

72


Fig. 15. Ramichloridium indicum (CBS 171.96). A–B. Macronematous conidiophores. C–E. Sympodially proliferating conidiogenous cells, resulting in a conidium-bearing rachis 

with pigmented and thickened scars. F. Conidia. Scale bar = 10 µm. 

Fig. 16. Ramichloridium strelitziae (CBS 121711). A–C. Conidial apparatus at different stages of development, resulting in macronematous conidiophores and sympodially 

proliferating conidiogenous cells. D–E. Rachis with crowded, slightly pigmented, thickened, circular scars. F. Conidia. Scale bars = 10 µm. 

Notes: Phylogenetically, this species together with Ramichloridium 

apiculatum and R. musae cluster within the Mycosphaerellaceae 

clade. Ramichloridium cerophilum can be distinguished from its 

relatives by the production of secondary conidia and its distinct 

conidial hila. 

Ramichloridium indicum (Subram.) de Hoog, Stud. Mycol. 15: 

70. 1977. Fig. 15. 

Basionym: Chloridium indicum Subram., Proc. Indian Acad. Sci., 

Sect. B, 42: 286. 1955 [non Rhinocladiella indica Agarwal, Lloydia 

32: 388. 1969]. 

≡ Veronaea indica (Subram.) M.B. Ellis, in Ellis, More Dematiaceous 


= Veronaea verrucosa Geeson, Trans. Brit. Mycol. Soc. 64: 349. 1975. 

In vitro: Submerged hyphae smooth, thin-walled, hyaline, 1–2.5 

µm wide, with thin septa; aerial hyphae coarsely verrucose, 

olivaceous-green, rather thick-walled, 2–2.5 µm wide, with thin 

septa. Conidiophores arising vertically from creeping hyphae at right 

angles, straight, unbranched, thick-walled, smooth, dark brown, 

with up to 10 thin septa, up to 250 µm long, 2–4 µm wide, often 

with inflated basal cells. Conidiogenous cells terminally integrated, 

up to 165 µm long, smooth, dark brown, sympodially proliferating, 

rachis straight or flexuose, geniculate or nodose, subhyaline; scars 

thickened and darkened, clustered at nodes, approx. 0.5 µm diam. 

Microcyclic conidiation observed in culture. Conidia solitary, (0–)1- 

septate, not constricted at the septum, subhyaline to pale brown, 

smooth or coarsely verrucose, rather thin-walled, broadly ellipsoidal 

to globose, (5–)7–8(–10) × (4–)6–6.5(–9) µm, with truncate base; 

hilum conspicuous, slightly darkened, not thickened, about 1 µm 

diam. 


73



after 14 d at 24 °C. Colonies velvety, rather compact, slightly 

elevated, with entire, smooth, whitish margin, dark olivaceousgreen 

in the central part. 

Specimen examined: Living culture, Feb. 1996, L. Marvanová, CBS 171.96. 

Ramichloridium pini de Hoog & Rahman, Trans. Brit. Mycol. Soc. 

81: 485. 1983. 

Specimen examined: U.K., Scotland, Old Aberdeen, branch of Pinus contorta 

(Pinaceae), 1982, M.A. Rahman, ex-type strain, CBS 461.82 = MUCL 28942. 

Note: The culture examined (CBS 461.82) was sterile. For a full 

description see de Hoog et al. (1983). 

Ramichloridium strelitziae Arzanlou, W. Gams & Crous, sp. nov. 

MycoBank MB504551. Figs 16–17A. 

Etymology: Named after its host, Strelitzia. 

Ab aliis speciebus Ramichloridii conidiophoris brevibus, ad 40 μm longis, et 

cicatricibus rotundis, paulo protrudentibus distinguendum. 

In vitro: Submerged hyphae smooth, hyaline, thin-walled, 2–2.5 µm 

wide; aerial hyphae pale brown, verrucose. Conidiophores arising 

vertically from creeping aerial hyphae, clearly differentiated from the 

vegetative hyphae, subhyaline, later becoming pale brown, thickwalled, 

smooth or verruculose, with 1–3 additional septa; up to 40 

µm long and 2 µm wide. Conidiogenous cells integrated, terminal, 

cylindrical, variable in length, 10–35 µm long, subhyaline, later 

turning pale brown, fertile part as wide as the basal part, proliferating 

sympodially, forming a straight rachis with slightly thickened and 

darkened, circular, somewhat protruding scars, approx. 0.5 µm 

diam. Conidia solitary, aseptate, smooth or verruculose, subhyaline, 

oblong, ellipsoidal to clavate, (3–)4–5(–5.5) × (1–)2(–2.5) µm, with 

truncate base and unthickened, non-pigmented hilum. 



rather compact, raised, dense, olivaceous-grey; reverse olivaceousblack. 

Specimen examined: South Africa, KwaZulu-Natal, Durban, near Réunion, 

on leaves of Strelitzia nicolai, 5 Feb. 2005, W. Gams & H. Glen, CBS-H 19776, 

holotype, culture ex-type CBS 121711. 

Zasmidium Fr., Summa Veg. Scand. 2: 407. 1849. 

Fig. 17. A. Ramichloridium strelitziae (CBS 121711). B. Veronaea japonica (CBS 

776.83). C. Veronaeopsis simplex (CBS 588.66). Scale bar = 10 µm. 

In vitro: Submerged hyphae smooth, thin-walled, hyaline, with 

thin septa; aerial hyphae coarsely verrucose, olivaceous-green, 

thick-walled, with thin septa. Conidiophores not differentiated 

from vegetative hyphae, often reduced to conidiogenous 

Fig. 18. Zasmidium cellare (CBS 146.36). A–D. Micronematous conidiophores with terminal, integrated conidiogenous cells. E. Conidiogenous cell with pigmented, thickened 

and refractive scars. F–G. Primary and secondary conidia. Scale bar = 10 µm. 

74


Fig. 19. Rhinocladiella anceps (CBS 181.65). A. Macronematous conidiophores. B–D. Conidial apparatus at different stages of development, resulting in semi-micronematous 

conidiophores and sympodially proliferating conidiogenous cells. E. Conidiogenous loci. F. Conidia. Scale bars = 10 µm. 

cells. Conidiogenous cells integrated, predominantly terminal, 

sometimes lateral, arising from aerial hyphae, cylindrical, pale 

brown; polyblastic, proliferating sympodially producing crowded, 

conspicuously pigmented, almost flat, darkened, somewhat 

refractive scars. Conidia in short chains, cylindrical to fusiform, 

verrucose, obovate to obconical, pale brown, base truncate, 

with a conspicuous, slightly pigmented, thickened and refractive 

hilum. Primary conidia sometimes larger, subhyaline, verrucose 

or smooth-walled, 0–4-septate, variable in length, fusiform to 

cylindrical; conidial secession schizolytic. 

Type species: Zasmidium cellare (Pers. : Fr.) Fr., Summa Veg. 

Scand. 2: 407. 1849. 

Zasmidium cellare (Pers. : Fr.) Fr., Summa Veg. Scand. 2: 407. 

1849. Fig. 18. 

Basionym: Racodium cellare Pers., Neues Mag. Bot. 1: 123. 1794. 

≡ Antennaria cellaris (Pers. : Fr.) Fr., Syst. Mycol. 3: 229. 1829. 

≡ Cladosporium cellare (Pers. : Fr.) Schanderl, Zentralbl. Bakteriol., 2. 

Abt., 94: 117. 1936. 

≡ Rhinocladiella cellaris (Pers. : Fr.) M.B. Ellis, in Ellis, Dematiaceous 


In vitro: Submerged hyphae smooth, thin-walled, hyaline, 2–3 

µm wide, with thin septa; aerial hyphae coarsely verrucose, 

olivaceous-green, rather thick-walled, 2–2.5 µm wide, with thin 

septa. Conidiophores not differentiated from vegetative hyphae, 

often reduced to conidiogenous cells. Conidiogenous cells 

integrated, predominantly terminal, sometimes lateral, arising from 

aerial hyphae, cylindrical, 20–60 µm long and 2–2.5 µm wide, pale 

brown, proliferating sympodially producing crowded, conspicuously 

pigmented scars that are thickened and refractive, about 1 µm diam. 

Conidia cylindrical to fusiform, verrucose, obovate to obconical, 

pale brown, with truncate base, (6–)9–14(–27) × 2–2.5 µm, with 

a conspicuous, slightly pigmented, refractive hilum, approx. 1 µm 

diam. Primary conidia sometimes subhyaline, verrucose or smoothwalled, 

thin-walled, 0–1(–4)-septate, variable in length, fusiform to 

cylindrical. 


Cultural characteristics: Colonies reaching 7 mm diam after 14 d 

at 24 °C. Colonies velvety, rather compact, slightly elevated with 

entire margin; surface dark olivaceous-green in the central part, 

margin smooth, whitish. 

Specimen examined: Wall in wine cellar, Jun. 1936, H. Schanderl, ATCC 36951 

= IFO 4862 = IMI 044943 = LCP 52.402 = LSHB BB274 = MUCL 10089 = CBS 

146.36. 

Notes: The name Racodium Fr., typified by Ra. rupestre Pers. : Fr., 

has been conserved over the older one by Persoon, with Ra. cellare 

as type species. De Hoog (1979) defended the use of Zasmidium in 

its place for the well-known wine-cellar fungus. 

Morphologically Zasmidium resembles Stenella Syd., and 

both reside in the Capnodiales, though the type of Stenella, S. 

araguata Syd., clusters in the Teratosphaeriaceae, and the type of 

Zasmidium, Z. cellare, in the Mycosphaerellaceae. When accepting 

anamorph genera as polyphyletic within an order, preference would 

be given to the well-known name Stenella over the less known 

Zasmidium, even though the latter name is older. Further studies 

are required, however, to clarify if all stenella-like taxa should be 

accommodated in a single genus, Stenella. If this is indeed the 

case, a new combination for Zasmidium cellare will be proposed 

in Stenella, and the latter genus will have to be conserved over 

Zasmidium. 

Chaetothyriales (Herpotrichiellaceae) 

The four “Ramichloridium” species residing in the Chaetothyriales 

clade do not differ sufficiently in morphology to separate them from 

Rhinocladiella (type Rh. atrovirens). Because of the pale brown 

conidiophores, conidiogenous cells with crowded, slightly prominent 

scars and the occasional presence of an Exophiala J.W. Carmich. 

synanamorph, Rhinocladiella is a suitable genus to accommodate 

them. These four species chiefly differ from Ramichloridium in the 

morphology of their conidial apparatus, which is clearly differentiated 

from the vegetative hyphae. The appropriate combinations are 

therefore introduced for Ramichloridium anceps, R. mackenziei, R. 

fasciculatum and R. basitonum. 

75


Fig. 20. Rhinocladiella basitona (CBS 101460). A–B. Semi-micronematous conidiophores with verticillate branching pattern. C–D. Sympodially proliferating conidiogenous cells, 

giving rise to a long rachis with slightly prominent, truncate conidium-bearing denticles. E. Intercalary conidiogenous cell. F. Conidia. Scale bars = 10 µm. 

The genus Veronaea (type species: V. botryosa) also resides 

in the Chaetothyriales clade. Veronaea can be distinguished from 

Rhinocladiella by the absence of exophiala-type budding cells and 

its predominantly 1-septate conidia. Furthermore, the conidiogenous 

loci in Veronaea are rather flat, barely prominent. 

Rhinocladiella Nannf., Svensk Skogsvårdsfören. Tidskr., Häfte 32: 

461. 1934. 

In vitro: Colonies dark olivaceous-brown, slow-growing, almost 

moist. Submerged hyphae hyaline to pale olivaceous, smooth; 

aerial hyphae, if present, more darkly pigmented. Exophialatype 

budding cells usually present in culture. Conidial apparatus 

usually branched, olivaceous-brown, consisting of either slightly 

differentiated tips of ascending hyphae or septate, markedly 

differentiated conidiophores. Conidiogenous cells intercalary or 

terminal, polyblastic, cylindrical to acicular, with a sympodially 

proliferating, subdenticulate rachis; scars unthickened, nonpigmented 

to somewhat darkened-refractive. Conidia solitary, 

hyaline to subhyaline, aseptate, thin-walled, smooth, subglobose, 

with a slightly pigmented hilum; conidial secession schizolytic. 

Type species: Rh. atrovirens Nannf., Svenska Skogsvårdsfören. 

Tidskr. 32: 461. 1934. 

Rhinocladiella anceps (Sacc. & Ellis) S. Hughes, Canad. J. Bot. 

36: 801. 1958. Fig. 19. 

Basionym: Sporotrichum anceps Sacc. & Ellis, Michelia 2: 576. 

1882. 

= Veronaea parvispora M.B. Ellis, in Ellis, More Dematiaceous Hyphomycetes: 

210. 1976. 

Misapplied name: Chloridium minus Corda sensu Mangenot, Rev. 

Mycol. (Paris) 18: 137. 1953. 

In vitro: Submerged hyphae subhyaline, smooth, thick-walled, 2– 

2.5 µm wide; aerial hyphae pale brown. Swollen germinating cells 

often present on MEA, giving rise to an Exophiala synanamorph. 

Conidiophores slightly differentiated from vegetative hyphae, arising 

from prostrate aerial hyphae, consisting of either unbranched or 

loosely branched stalks, thick-walled, golden to dark-brown, up 

to 350 µm tall, which may have up to 15 thin, additional septa, 

intercalary cells 9–14 µm long. Conidiogenous cells terminal, 

rarely lateral, cylindrical, occasionally intercalary, variable in length, 

smooth, golden to dark brown at the base, paler toward the apex, 

later becoming inconspicuously septate, fertile part as wide as the 

basal part, 15–40 × 1.5–2 µm; with crowded, slightly prominent, 

unpigmented, conidium-bearing denticles, about 0.5 µm diam. 

Conidia solitary, subhyaline, thin-walled, smooth, subglobose to 

ellipsoidal, 2.5–4 × 2–2.5 µm, with a less conspicuous, slightly 

darkened hilum, less than 0.5 µm diam. 

Cultural characteristics: Colonies on MEA reaching 6–12 mm diam 


powdery, becoming hairy at centre; olivaceous-green to brown, 

reverse dark-olivaceous. 

Specimens examined: Canada, Ontario, Campbellville, from soil under Thuja plicata, 

Apr. 1965, G. L. Barron, CBS H-7715 (isoneotype); CBS H-7716 (isoneotype); CBS 

H-7717 (isoneotype); CBS H-7718 (isoneotype); CBS H-7719 (isoneotype), ex-type 

strain, CBS 181.65 = ATCC 18655 = DAOM 84422 = IMI 134453 = MUCL 8233 = 

OAC 10215. France, from stem of Fagus sylvatica, 1953, F. Mangenot, CBS 157.54 

= ATCC 15680= MUCL 1081= MUCL 7992 = MUCL 15756. 

Notes: Rhinocladiella anceps (conidia 2.5–4 µm long) resembles 

Rh. phaeophora Veerkamp & W. Gams (1983) (conidia 5.5–6 µm 

long), but has shorter conidia. 

76


Fig. 21. Rhinocladiella fasciculata (CBS 132.86). A. Conidiophores. B. Sympodially proliferating conidiogenous cells, which give rise to a long rachis with slightly prominent, 

unthickened scars. C. Conidia. D–E. Synanamorph consisting of conidiogenous cells with percurrent proliferation. F. Conidia. Scale bars = 10 µm. 

Rhinocladiella basitona (de Hoog) Arzanlou & Crous, comb. nov. 


Basionym: Ramichloridium basitonum de Hoog, J. Clin. Microbiol. 

41: 4774. 2003. 

In vitro: Submerged hyphae hyaline, smooth, thin-walled, 2 µm 

wide; aerial hyphae rather thick-walled, pale brown. Conidiophores 

slightly differentiated from vegetative hyphae, profusely and mostly 

verticillately branched, straight or flexuose, pale-brown, 2–2.5 µm 

wide. Conidiogenous cells terminal, variable in length, 10–100 µm 

long, pale brown, straight or geniculate, proliferating sympodially, 

giving rise to a long, 2–2.5 µm wide rachis, with slightly prominent, 

truncate conidium-bearing denticles, slightly darkened. Conidia 

solitary, hyaline, thin-walled, smooth, pyriform to clavate, with a 

round apex, and slightly truncate base, (1–)3–4(–5) × 1–2 µm, 

hilum conspicuous, slightly darkened and thickened, less than 0.5 

µm diam. 



rather flat and slightly elevated in the centre, pale olivaceous-grey 

to olivaceous-grey; reverse olivaceous-black. 

Specimen examined: Japan, Hamamatsu, from subcutaneous lesion with fistula on 

knee of 70-year-old male, Y. Suzuki, ex-type culture CBS 101460 = IFM 47593. 

Rhinocladiella fasciculata (V. Rao & de Hoog) Arzanlou & Crous, 


Basionym: Ramichloridium fasciculatum V. Rao & de Hoog, Stud. 

Mycol. 28: 39. 1986. 


In vitro: Submerged hyphae subhyaline, smooth, thick-walled, 

2–2.5 µm wide; aerial hyphae pale brown. Conidiophores arising 

vertically from ascending hyphae in loose fascicles, unbranched or 

loosely branched at acute angles, cylindrical, smooth, brown and 

thick-walled at the base, up to 220 µm long and 2–3 µm wide, with 

0–5 thin additional septa. Conidiogenous cells terminal, cylindrical, 

30–100 µm long, thin-walled, smooth, pale brown, fertile part as 

wide as the basal part, up to 2 µm wide, proliferating sympodially, 

giving rise to a rachis with hardly prominent, slightly pigmented, 

not thickened scars, less than 0.5 µm diam. Conidia solitary, 

smooth, thin-walled, subhyaline, ellipsoidal, (2.5–)4–5(–6) × 2–3 

µm, with truncate, slightly pigmented hilum, about 0.5 µm diam. 

Synanamorph forming on torulose hyphae originating from giant 

cells; compact heads of densely branched hyphae forming thinwalled, 

lateral, subglobose cells, on which conidiogenous cells are 

formed; conidiogenous cells proliferating percurrently, giving rise to 

tubular annellated zones with inconspicuous annellations, up to 12 

µm long, 1–1.5 µm wide. Conidia smooth, thin-walled, aseptate, 

subhyaline, globose, 2–2.5 µm diam. 

Cultural characteristics: Colonies on MEA reaching 8 mm diam after 

14 d at 24 °C, with entire, smooth, sharp margin; mycelium velvety, 

becoming farinose in the centre due to abundant sporulation, 

olivaceous-green to brown, reverse dark olivaceous. Blackish 

droplets often produced at the centre, which contain masses of 

Exophiala conidia. 

Specimen examined: India, Karnataka, Thirathahalli, isolated by V. Rao from 

decayed wood, holotype CBS-H 3866, culture ex-type CBS 132.86. 

77


Fig. 22. Rhinocladiella mackenziei (CBS 368.92). A. Intercalary conidiogenous cell. B–E. Semi-micronematous conidiophores and sympodially proliferating conidiogenous cells, 

resulting in a rachis with slightly prominent, unthickened scars. F. Conidia. Scale bar = 10 µm. 

78


Fig. 24. Thysanorea papuana (CBS 212.96), periconiella-like synanamorph. A. Macronematous conidiophores. B–C. Conidiophores with dense apical branches. D. Branches 

with different levels of branchlets. E–I. Conidiogenous cells at different stages of development; sympodially proliferating conidiogenous cells give rise to a denticulate rachis. 

J–K. Conidia. Scale bars = 10 µm. 

Fig. 23. (Page 78). Thysanorea papuana (CBS 212.96). A. Intercalary conidiogenous cell. B–I. Semi-micronematous conidiophores and sympodially proliferating conidiogenous 

cells, resulting in a rachis with prominent conidium bearing denticles. J–K. Microcyclic conidiation observed in slide cultures. L. Conidia. Scale bar = 10 µm. 


79


Rhinocladiella mackenziei (C.K. Campb. & Al-Hedaithy) Arzanlou 

& Crous, comb. nov. MycoBank MB504554. Fig. 22. 

Basionym: Ramichloridium mackenziei C.K. Campb. & Al-Hedaithy, 

J. Med. Veterin. Mycol. 31: 330. 1993. 

In vitro: Submerged hyphae subhyaline, smooth, thin-walled, 2–3 

µm wide; aerial hyphae pale brown, slightly narrower. Conidiophores 

slightly or not differentiated from vegetative hyphae, arising 

laterally from aerial hyphae, with one or two additional septa, often 

reduced to a discrete or intercalary conidiogenous cell, pale-brown, 

10–25 × 2.5–3.5 µm. Conidiogenous cells terminal or intercalary, 

variable in length, 5–15 µm long and 3–5 µm wide, occasionally 

slightly wider than the basal part, pale brown, rachis with slightly 

prominent, unpigmented, non-thickened scars, about 0.5 µm diam. 

Conidia golden-brown, thin-walled, smooth, ellipsoidal to obovate, 

subcylindical, (5–)8–9(–12) × (2–)3–3.5(–5) µm, with darkened, 

inconspicously thickened, protuberant or truncate hilum, less than 

1 µm diam. 



densely lanose and elevated in the centre, olivaceous-green to 

brown; reverse dark olivaceous. 

Specimens examined: Israel, Haifa, isolated from brain abscess, CBS 368.92 = 

UTMB 3170; human brain abscess, E. Lefler, CBS 367.92 = NCPF 2738 = UTMB 

3169. Saudi Arabia, from phaeohyphomycosis of the brain, S.S.A. Al-Hedaithy, 

ex-type strain, CBS 650.93 = MUCL 40057 = NCPF 2808; from brain abscess, 

Pakistani male who travelled to Saudi Arabia, CBS 102592 = NCPF 7460. United 

Arab Emirates, from fatal brain abscess, CBS 102590 = NCPF 2853. 

Notes: Morphologically Rhinocladiella mackenziei is somewhat 

similar to Pleurothecium obovoideum (Matsush.) Arzanlou & 

Crous, which was originally isolated from dead wood. However, P. 

obovoideum has distinct conidiophores, and the ascending hyphae 

are thick-walled, and the denticles cylindrical, up to 1.5 µm long. 

In contrast, Rh. mackenziei has only slightly prominent denticles. 

Rhinocladiella mackenziei is a member of the Chaetothyriales, 

while P. obovoideum clusters in the Chaetosphaeriales. 

Thysanorea Arzanlou, W. Gams & Crous, gen. nov. MycoBank 

MB504555. 

Etymology: (Greek) thysano = brush, referring to the brush-like 

branching pattern, suffix derived from Veronaea. 

Veronaeae similis sed conidiophoris partim Periconiae similibus dense ramosis 

distinguenda. 

In vitro: Submerged hyphae subhyaline, smooth, thin-walled; aerial 

hyphae pale brown, smooth or verrucose. Conidiophores dimorphic; 

micronematous conidiophores slightly differentiated from vegetative 

hyphae, branched or simple, multiseptate. Conidiogenous cells 

terminal, polyblastic, variable in length, smooth, golden- to dark 

brown at the base, paler towards the apex, later sometimes 

inconspicuously septate; fertile part often wider than the basal 

part, clavate to doliiform, with crowded, more or less prominent 

conidium-bearing denticles, unpigmented, but slightly thickened. 

Macronematous conidiophores consisting of well-differentiated, 

thick-walled, dark brown stalks; apically repeatedly densely 

branched, forming a complex head, each branchlet giving rise 

to a conidium-bearing denticulate rachis with slightly pigmented, 

thickened scars. Conidia of both kinds of conidiophore formed 

singly, smooth, pale brown, obovoidal to pyriform, (0–)1-septate, 

with a truncate base and darkened hilum; conidial secession 

schizolytic. 

Type species: Thysanorea papuana (Aptroot) Arzanlou, W. Gams 

& Crous, comb. nov. 

Thysanorea papuana (Aptroot) Arzanlou, W. Gams & Crous, 

comb. nov. MycoBank MB504556. Figs 7C, 23–24. 

Basionym: Periconiella papuana Aptroot, Nova Hedwigia 67: 491. 

1998. 

In vitro: Submerged hyphae subhyaline, smooth, thin-walled, 1.5–3 

µm wide; aerial hyphae pale brown, smooth to verrucose, 1.5–2 

µm wide. Conidiophores dimorphic; micronematous conidiophores 

slightly differentiated from vegetative hyphae, branched or simple, 

up to 6-septate. Conidiogenous cells terminal or intercalary, variable 

in length, 5–20 µm long, thin-walled, smooth, golden- to dark brown 

at the base, paler toward the apex, later sometimes becoming 

inconspicuously septate, fertile part wider than basal part, often 

clavate, with crowded, more or less prominent conidium-bearing 

denticles, about 1 µm diam, unpigmented but slightly thickened. 

Conidia solitary, subhyaline, thin-walled, smooth, cylindrical to 

pyriform, rounded at the apex and truncate at the base, pale brown, 

(0–)1-septate, (5–)7–8(–11) × (2–)3(–4) µm, with a truncate base 

and darkened hilum, 1 µm diam. Macronematous conidiophores 

present in old cultures after 1 mo of incubation, consisting of welldifferentiated, 

thick-walled, dark brown stalks, up to 220 µm long, 

(4–)5–6(–7) µm wide, with up to 15 additional septa, often with 

inflated basal cells; apically densely branched, forming a complex 

head, with up to five levels of branchlets, 20–50 µm long, each 

branchlet giving rise to a denticulate conidium-bearing rachis; scars 

slightly pigmented, thickened, about 1 µm diam. Conidia solitary, 

thin-walled, smooth, pale brown, obovoidal to pyriform, (0–)1- 

septate, (4–)5–6(–8) × (2–)3(–4) µm, with a truncate base and 

darkened hilum, 1–2 µm diam. 


after 14 d at 24 °C, with entire, sharp margin; mycelium velvety, 

elevated, with colonies up to 2 mm high, surface olivaceous-grey to 

iron-grey; reverse greenish black. 

Specimen examined: Papua New Guinea, Madang Province, foothill of Finisterre 

range, 40.8 km along road Madang-Lae, alt. 200 m, isolated from unknown stipe, 2 

Nov. 1995, A. Aptroot, holotype CBS-H 6351, culture ex-type CBS 212.96. 

Veronaea Cif. & Montemart., Atti Ist. Bot. Lab. Crittog. Univ. Pavia, 

sér. 5, 15: 68. 1957. 

In vitro: Colonies velvety, pale olivaceous-brown, moderately 

fast-growing. Submerged hyphae hyaline to pale olivaceous, 

smooth; aerial hyphae, more darkly pigmented. Exophiala-type 

budding cells absent in culture. Conidiophores erect, straight or 

flexuose, unbranched or occasionally loosely branched, sometimes 

geniculate, smooth-walled, pale to medium- or olivaceous-brown. 

Conidiogenous cells terminally integrated, polyblastic, occasionally 

intercalary, cylindrical, pale brown, later often becoming septate, 

fertile part subhyaline, often as wide as the basal part, rachis with 

crowded, flat to slightly prominent, faintly pigmented, unthickened 

scars. Conidia solitary, smooth, cylindrical to pyriform, rounded 

at the apex and truncate at the base, pale brown, 1(–2)-septate; 


Type species: Veronaea botryosa Cif. & Montemart., Atti Ist. Bot. 

Lab. Crittog. Univ. Pavia, sér. 5, 15: 68. 1957. 

Veronaea botryosa Cif. & Montemart., Atti Ist. Bot. Lab. Crittog. 

Univ. Pavia, sér. 5, 15: 68. 1957. Fig. 25. 

80


Fig. 25. Veronaea botryosa (CBS 254.57). A–C. Semi-micronematous conidiophores and sympodially proliferating conidiogenous cells. D–E. Rachis with crowded and flat 

scars. F–G. Microcyclic conidiation. H. Conidia. Scale bars = 10 µm. 

In vitro: Submerged hyphae hyaline to pale olivaceous, smooth; 

aerial hyphae more darkly pigmented. Conidiophores erect, 

straight or flexuose, unbranched or occasionally loosely branched, 

sometimes geniculate, smooth-walled, pale brown to olivaceousbrown, 

2–3 µm wide and up to 200 µm long. Conidiogenous cells 

terminal, occasionally intercalary, cylindrical, 10–100 µm long, 

pale brown, later often becoming septate, fertile part subhyaline, 

often as wide as the basal part, rachis with crowded, flat to slightly 

prominent, faintly pigmented, unthickened scars. Conidia solitary, 

smooth, cylindrical to pyriform, (3–)6.5–8.5(–12) × (1.5–)2–2.5(–3) 

µm, rounded at the apex and truncate at the base, pale brown, 

1(–2)-septate, with a faintly darkened, unthickened hilum, about 0.5 

µm diam. 


after 14 d at 24 °C, with entire, sharp margin; mycelium velvety, 

slightly elevated in the centre, surface olivaceous-grey to greyishbrown; 

reverse greenish black. 

Specimens examined: India, Ramgarh, about 38 km from Jaipur, isolated from goat 

dung, 1 Sep. 1963, B.C. Lodha, CBS 350.65 = IMI 115127 = MUCL 7972. Italy, 

Tuscany, Pisa, isolated from Sansa olive slag, 1954, O. Verona, ex-type strain, CBS 

254.57 = IMI 070233 = MUCL 9821. 

Veronaea compacta Papendorf, Bothalia 12: 119. 1976. Fig. 26. 

In vitro: Submerged hyphae subhyaline, smooth, thin-walled, 

1.5–3 µm wide; aerial hyphae rather thick-walled, pale brown. 

Conidiophores slightly differentiated from vegetative hyphae, 

lateral or occasionally terminal, often wider than the supporting 

hypha, up to 4 µm wide, unbranched or branched at acute angles, 

with 1–3 adititional septa, cells often inflated and flask-shaped, 

pale-brown, up to 60 µm long. Conidiogenous cells terminal, 

occasionally intercalary, variable in length, up to 10 µm long, 

pale brown, cylindrical to doliiform or flask-shaped, with hardly 

prominent denticles; scars flat, slightly pigmented, not thickened, 

about 0.5 µm diam. Conidia solitary, pale brown, smooth, thinwalled, 

ellipsoidal to ovoid, 0–1(–2)-septate, often constricted at 

the septa, (4–)6–7(–9) × 2–3 µm, with a round apex and truncate 

base; hilum prominent, slightly darkened, unthickened, about 0.5 

µm diam. 

Cultural characteristics: Colonies rather slow growing, reaching 15 

mm diam on MEA after 14 d at 24 °C; surface velvety to lanose, 

slightly raised in the centre, pale grey to pale brownish grey; reverse 

dark grey. 

Specimen examined: South Africa, soil, M.C. Papendorf, ex-type culture CBS 

268.75. 


81


Fig. 26. Veronaea compacta (CBS 268.75). A–B. Semi-micronematous conidiophores and sympodially proliferating conidiogenous cells. C–D. Rachis with hardly prominent 

denticles. E. Conidia. Scale bar = 10 µm. 

Fig. 27. Veronaea japonica (CBS 776.83). A. Intercalary conidiogenous cells. B–D. Semi-micronematous conidiophores and sympodially proliferating conidiogenous cells. E. 

Conidia. F. Thick-walled, dark brown hyphal cells. Scale bar = 10 µm. 

Veronaea japonica Arzanlou, W. Gams & Crous, sp. nov. 

MycoBank MB504557. Figs 17B, 27. 

Etymology: Named after the country of origin, Japan. 

Veronaeae compactae similis, sed cellulis inflatis, aggregatis, crassitunicatis, fuscis 

in vitro formatis distinguenda. 

In vitro: Submerged hyphae subhyaline, smooth, thin-walled, 1.5–3 

µm wide; aerial hyphae slightly narrower, pale brown; hyphal cells 

later becoming swollen, thick-walled, dark brown, often aggregated. 

Conidiophores slightly differentiated from aerial vegetative hyphae, 

lateral, or terminal, often wider than the supporting hypha, 2–3 µm 

wide, up to 65 µm long, unbranched or occasionally branched, 

82


Fig. 28. Pleurothecium obovoideum (CBS 209.95). A–C. Conidial apparatus 

consisting of conidiophores with sympodially proliferating conidiogenous cells as 

seen in slide cultures of ca. 14 d. D. Short chain of conidia. E–G. Sympodially 

proliferating conidiogenous cells, resulting in a short rachis with subcylindrical to 

cylindrical denticles. H. Conidia. Scale bar = 10 µm. 

pale brown, thin-walled, smooth, with 1–3 additional septa. 

Conidiogenous cells terminal, occasionally intercalary, variable in 

length, up to 15 µm long, pale brown, cylindrical to clavate, with 

hardly prominent denticles; scars flat, slightly pigmented, not 

thickened, about 0.5 µm diam. Conidia solitary, pale brown, smooth, 

thin-walled, ellipsoidal to ovoid, (0–)1-septate, often constricted at 

the septum, (6–)7–8(–10) × 2–2.5(–4) µm, with a round apex and 

truncate base; hilum unthickened but slightly darkened, about 1 µm 

diam. 

Cultural characteristics: Colonies rather slow growing, reaching 7.5 

mm diam on MEA after 14 d at 24 °C; surface velvety to lanose, 

slightly raised in the centre, olivaceous-brown, with entire margin; 

reverse dark-olivaceous. 

Specimen examined: Japan, Kyoto, Daitokuji Temple, Kyoto, inside dead bamboo 

culm, Dec. 1983, W. Gams, holotype CBS-H 3490, ex-type culture CBS 776.83. 

Note: This species is morphologically similar to V. compacta 

(Papendorf 1976), but can be distinguished based on the presence 

of dark brown, swollen hyphal cells in culture, which are absent in 

V. compacta. 

Pleurothecium obovoideum clade (Chaetosphaeriales) 

Ramichloridium obovoideum was regarded as similar to 

“Ramichloridium” (Rhinocladiella) mackenziei by some authors, 

and subsequently reduced to synonymy (Ur-Rahman et al. 1988). 

However, R. obovoideum clusters with Carpoligna pleurothecii, 

the teleomorph of Pleurothecium Höhn. Because it is also 

morphologically similar to other species of Pleurothecium, we 

herewith combine it into that genus. 

Pleurothecium obovoideum (Matsush.) Arzanlou & Crous, comb. 


Basionym: Rhinocladiella obovoidea Matsush., Icones Microfung. 

Mats. lect.: 123. 1975. 

≡ Ramichloridium obovoideum (Matsush.) de Hoog, Stud. Mycol. 15: 73. 

1977. 

In vitro: Submerged hyphae smooth, hyaline, thin-walled, 1–2 µm 

wide; aerial hyphae hyaline to subhyaline, smooth. Conidiophores 

arising vertically from creeping hyphae, ascending hyphae thickwalled 

and dark brown; conidiophores 10–35 µm long, 1–2-septate, 

often reduced to a conidiogenous cell, unbranched, thick-walled, 

smooth, tapering towards the apex, pale brown. Conidiogenous 

cells integrated, cylindrical to ampulliform, 5–20 µm long, pale 

brown, elongating sympodially, with a short rachis giving rise to 

denticles, 1 µm long, slightly pigmented. Conidia aseptate, solitary 

or in short chains of up to 3, smooth, pale brown, ellipsoidal to 

obovate, (9–)11–12(–14.5) × (3–)4(–5) µm, smooth, thin-walled, 

with a more or less rounded apex, a truncate base and a slightly 

darkened, unthickened hilum, 1.5 µm diam. 


83


Cultural characteristics: Colonies slow-growing, reaching 15 diam 

after 14 d at 24 °C, with entire, smooth margin; surface rather 

compact, mycelium mainly flat, submerged, some floccose to 

lanose aerial mycelium in the centre, buff; reverse honey. 

Specimen examined: Japan, Kobe Municipal Arboretum, T. Matsushima, from dead 

leaf of Pasania edulis, CBS 209.95 = MFC 12477. 

Incertae sedis (Sordariomycetes) 

Ramichloridium schulzeri clade 

Ramichloridium schulzeri, including its varieties, clusters 

near Thyridium Nitschke and the Magnaporthaceae, and is 

phylogenetically as well as morphologically distinct from the other 

genera in the Ramichloridium complex. To accommodate these 

taxa, a new genus is introduced below. 

Myrmecridium Arzanlou, W. Gams & Crous, gen. nov. MycoBank 

MB504559. 

Etymology: (Greek) myrmekia = wart, referring to the wart-like 

denticles on the rachis, suffix -ridium from Chloridium. 

Genus ab allis generibus Ramichloridii similibus rachide recta longa, subhyalina, 

denticulis distantibus, verruciformibus praedita distinguendum. 

In vitro: Colonies moderately fast-growing, flat, with mainly 

submerged mycelium, and entire margin, later becoming powdery 

to velvety, pale orange to orange. Mycelium rather compact, mainly 

submerged, in the centre velvety with fertile bundles of hyphae. 

Conidiophores arising vertically and clearly distinct from creeping 

hyphae, unbranched, straight or flexuose, brown, thick-walled. 

Conidiogenous cells terminally integrated, polyblastic, cylindrical, 

straight or flexuose, pale brown, sometimes secondarily septate, 

fertile part subhyaline, as wide as the basal part, with scattered 

pimple-shaped, apically pointed, unpigmented, conidium-bearing 

denticles. Conidia solitary, subhyaline, smooth or finely verrucose, 

rather thin-walled, with a wing-like gelatinous sheath, obovoidal or 

fusiform, tapering towards a narrowly truncate base with a slightly 

prominent, unpigmented hilum; conidial secession schizolytic. 

Type species: Myrmecridium schulzeri (Sacc.) Arzanlou, W. Gams 


Notes: Myrmecridium schulzeri was fully described as Acrotheca 

acuta Grove by Hughes (1951). The author discussed several 

genera, none of which is suitable for the present fungus for various 

reasons as analysed by de Hoog (1977). Only Gomphinaria Preuss 

is not yet sufficiently documented. Our examination of G. amoena 

Preuss (B!) showed that this is an entirely different fungus, of which 

no fresh material is available to ascertain its position. 

Myrmecridium can be distinguished from other ramichloridiumlike 

fungi by having entirely hyaline vegetative hyphae, and widely 

scattered, pimple-shaped denticles on the long hyaline rachis. 

The conidial sheath is visible in lactic acid mounts with brightfield 

microscopy. The Myrmecridium clade consists of several 

subclusters, which are insufficiently resolved based on the ITS 

sequence data. However, two morphologically distinct varieties of 

Myrmecridium are treated here. The status of the other isolates in 

this clade will be dealt with in a future study incorporating more 

strains, and using a multi-gene phylogenetic approach. 

Myrmecridium schulzeri (Sacc.) Arzanlou, W. Gams & Crous, 

comb. nov. MycoBank MB504560. var. schulzeri Figs 7B, 29. 

Basionym: Psilobotrys schulzeri Sacc., Hedwigia 23: 126. 1884. 

≡ Chloridium schulzerii (Sacc.) Sacc., Syll. Fung. 4: 322. 1886. 

≡ Rhinocladiella schulzeri (Sacc.) Matsush., Icon. Microfung. Mats. lect. 

(Kobe): 124. 1975. 

≡ Ramichloridium schulzeri (Sacc.) de Hoog, Stud. Mycol. 15: 64. 1977 

var. schulzeri. 

= Acrotheca acuta Grove, J. Bot., Lond. 54: 222. 1916. 

≡ Pleurophragmium acutum (Grove) M.B. Ellis in Ellis, More Dematiaceous 


= Rhinotrichum multisporum Doguet, Rev. Mycol., Suppl. Colon. 17: 78. 1953 

(nom. inval. Art. 36) [non Acrotheca multispora (Preuss) Sacc., Syll. Fung. 4: 

277. 1886]. 

[non Acrothecium (?) multisporum G. Arnaud, Bull. Trimestriel Soc. 

Mycol. France 69: 288. 1953 (nom. inval. Art. 36)]. 

[non Acrothecium multisporum G. Arnaud sensu Tubaki, J. Hattori 

Bot. Lab. 20: 145. 1958]. 

In vitro: Submerged hyphae hyaline, thin-walled, 1–2 µm wide; 

aerial hyphae, if present, pale olivaceous-brown. Conidiophores 

arising vertically from creeping aerial hyphae, unbranched, straight, 

reddish brown, thick-walled, septate, up to 250 µm tall, 2.5–3.5 

µm wide, with 2–7 additional septa, basal cell often inflated, 3.5–5 

µm wide. Conidiogenous cells integrated, cylindrical, variable in 

length, 15–110 µm long, subhyaline to pale brown, later becoming 

inconspicuously septate, fertile part subhyaline, as wide as the 

basal part, forming a straight rachis with scattered, pimple-shaped 

denticles less than 1 µm long and approx. 0.5 µm wide, apically 

pointed, unpigmented, slightly thickened scars. Conidia solitary, 

subhyaline, thin-walled, smooth or finely verrucose, surrounded 

by a wing-like, gelatinous conidial sheath, up to 0.5 µm thick, 

ellipsoid, obovoid or fusiform, (6–)9–10(–12) × 3–4 µm, tapering to 

a subtruncate base; hilum unpigmented, inconspicuous. 

Cultural characteristics: Colonies reaching 29 mm diam after 14 

d at 24 °C, pale orange to orange, with entire margin; mycelium 

flat, rather compact, later becoming farinose or powdery due to 

sporulation, which occurs in concentric zones when incubated on 

the laboratory bench. 

Specimens examined: Germany, Kiel-Kitzeberg, from wheat-field soil, W. Gams, 

CBS 134.68 = ATCC 16310. The Netherlands, isolated from a man, bronchial 

secretion, A. Visser, CBS 156.63 = MUCL 1079; Lienden, isolated from Triticum 

aestivum root, C.L. de Graaff, CBS 325.74 = JCM 7234. 

Myrmecridium schulzeri var. tritici (M.B. Ellis) Arzanlou, W. 

Gams & Crous, comb. nov. MycoBank MB504562. 

Basionym: Pleurophragmium tritici M.B. Ellis, in Ellis, More 


≡ Ramichloridium schulzeri var. tritici (M.B. Ellis) de Hoog, Stud. Mycol. 

15: 68. 1977. 

Specimen examined: Ireland, Dublin, on wheat stem, Oct. 1960, J.J. Brady, 

holotype IMI 83291. 

Notes: No reliable living culture is available of this variety. Based 

on a re-examination of the type specimen in this study, the variety 

appears sufficiently distinct from Myrmecridium schulzeri var. 

schulzeri based on the frequent production of septate conidia. 

Myrmecridium flexuosum (de Hoog) Arzanlou, W. Gams & Crous, 

comb. et stat. nov. MycoBank MB504563. Fig. 30. 

Basionym: Ramichloridium schulzeri var. flexuosum de Hoog, Stud. 

Mycol. 15: 67. 1977. 

In vitro: Submerged hyphae hyaline, thin-walled, 1–2 µm wide. 

Conidiophores unbranched, flexuose, arising from creeping aerial 

84


Fig. 29. Myrmecridium schulzeri (CBS 325.74). A. Macronematous conidiophores. B. Inflated basal cells visible in some conidiophores. C–E. Conidial apparatus at different 

stages of development, resulting in macronematous conidiophores and sympodially proliferating conidiogenous cells. F–G. Rachis with scattered, pimple-shaped denticles. 

H. Conidia. Scale bars: A =100 µm, B–H = 10 µm. 

hyphae, pale brown, up to 250 µm tall, 3–3.5 µm wide, thick-walled, 

smooth, with up to 24 thin septa, delimiting 8–12 µm long cells. 

Conidiogenous cells integrated, elongating sympodially, cylindrical, 

20–150 µm long, flexuose, brown at the base, subhyaline in the 

upper part, later becoming inconspicuously septate; rachis slightly 

flexuose, subhyaline, as wide as the basal part, thick-walled near 

the base, hyaline and thin-walled in the apical part, with scattered 

pimple-shaped, unpigmented, approx. 0.5 µm long denticles. 

Conidia solitary, subhyaline, thin-walled, finely verrucose, with 

a wing-like gelatinous sheath, approx. 0.5 µm wide, ellipsoid 

to obovoid, (5–)6–7(–9) × 3–4 µm; hilum slightly prominent, 

unpigmented, approx. 0.5 µm diam. 


at 24 °C; mycelium submerged, flat, smooth; centrally orange, later 

becoming powdery to velvety and greyish brown due to sporulation, 

with sharp, smooth, entire margin; reverse yellowish orange. 

Specimen examined: Surinam, isolated from soil, J.H. van Emden, ex-type culture 

CBS 398.76 = JCM 6968. 


Note: This former variety is sufficiently distinguished from M. 

schulzeri s. str. by its flexuose conidiophores and conidia which 

lack an acuminate base, to be regarded as a separate species. 

Ramichloridium torvi (Ellis & Everh.) de Hoog, Stud. Mycol. 15: 

79. 1977. 

≡ Ramularia torvi Ellis & Everh., Rep. Missouri Bot. Gard. 9: 119. 1898. 

≡ Hansfordia torvi (Ellis & Everh.) Deighton & Piroz., Mycol. Pap. 101: 

39. 1965. 

= Acladium biophilum Cif., Sydowia 10: 164. 1956. 

≡ Hansfordia biophila (Cif.) M.B. Ellis, in Ellis, More Dematiaceous 


Specimen: Jamaica, Port Marant, Dec. 1890, on leaves of Solanum torvum, 

holotype of Ramularia torvi (NY) (specimen not examined). 

Notes: According to the description and illustration of R. torvi 

provided by de Hoog (1977), this appears to be an additional 

species of Myrmecridium. Although it is morphologically similar to 

M. flexuosum in having a flexuose rachis, it differs from the other 

species of the genus by having smooth, clavate conidia. Fresh 

collections and cultures would be required to resolve its status. 

85


Fig. 30. Myrmecridium flexuosum (CBS 398.76). A–C. Conidial apparatus at different stages of development, resulting in macronematous conidiophores with sympodially 

proliferating conidiogenous cells. D–H. Sympodially proliferating conidiogenous cells giving rise to a flexuose conidium-bearing rachis with pimple-shaped denticles. I. Conidia. 

Scale bar = 10 µm. 

Fig. 31. Pseudovirgaria hyperparasitica (CBS 121739). A–D. Conidial apparatus at different stages of development; conidiogenous cells with geniculate proliferation. E. Conidia. 

Scale bar = 10 µm. 

86


Pseudovirgaria H.D. Shin, U. Braun, Arzanlou & Crous, gen. nov. 


Etymology: Named after its morphological similarity to Virgaria. 

Hyphomycetes. Uredinicola. Coloniae in vivo pallide vel modice brunneae, 

ferrugineae vel cinnamomeae, in vitro lentissime crescentes, murinae. Mycelium 

immersum et praecipue externum, ex hyphis ramosis et cellulis conidiogenis 

integratis compositum, conidiophoris ab hyphis vegetativis vix distinguendis. 

Hyphae ramosae, septatae, leves, tenuitunicatae, hyalinae vel pallide brunneae. 

Cellulae conidiogenae integratae in hyphis repentibus, terminales et intercalares, 

polyblasticae, sympodialiter proliferentes, subcylindricae vel geniculatae, cicatricibus 

conspicuis, solitariis vel numerosis, dispersis vel aggregatis, subdenticulatis, 

prominentibus, umbonatis vel apicem versus paulo attenuatis, non inspissatis, 

non vel parce fuscatis-refringentibus. Conidia solitaria, holoblastica, plus minusve 

obovoidea, recta vel leniter curvata, asymmetrica, continua, hyalina, subhyalina 

vel pallidissime olivaceo-brunnea, hilo subconspicuo vel conspicuo, truncato 

vel rotundato, non inspissato, non vel lenissime fuscato-refringente; secessio 

schizolytica. 

Hyperparasitic on uredosori of rust fungi. Colonies in vivo pale 

to medium brown, rusty or cinnamom, in vitro slow-growing, 

pale to dark mouse-grey. Mycelium immersed and mainly aerial, 

composed of branched hyphae with integrated conidiogenous 

cells, differentiation between vegetative hyphae and conidiophores 

barely possible. Hyphae branched, septate, smooth, thin-walled, 

hyaline to pale brown. Conidiogenous cells similarly hyaline to 

pale brown, integrated in creeping threads (hyphae), terminal and 

intercalary, polyblastic, proliferation sympodial, rachis subcylindrical 

to geniculate, conidiogenous loci (scars) conspicuous, solitary to 

numerous, scattered to aggregated, subdenticulate, bulging out, 

umbonate or slightly attenuated towards a rounded apex, wall 

unthickened, not to slightly darkened-refractive. Conidia solitary, 

formation holoblastic, more or less obovoid, straight to somewhat 

curved, asymmetrical, aseptate, hyaline, subhyaline to very pale 

olivaceous-brown, with more or less conspicuous hilum, truncate to 

rounded, unthickened, not or slightly darkened-refractive; conidial 


Type species: Pseudovirgaria hyperparasitica H.D. Shin, U. Braun, 

Arzanlou & Crous, sp.nov. 

Notes: Other ramichloridium-like isolates from various rust species 

form another unique clade, sister to Radulidium subulatum (de 

Hoog) Arzanlou, W. Gams & Crous and Ra. epichloës (Ellis & 

Dearn.) Arzanlou, W. Gams & Crous in the Sordariomycetidae. 

Although Pseudovirgaria is morphologically similar to Virgaria Nees, 

it has hyaline to pale brown hyphae, conidia and conidiogenous 

cells. The conidiogenous cells are integrated in creeping threads 

(hyphae), terminal and intercalary, and the proliferation is distinctly 

sympodial. The subdenticulate conidiogenous loci are scattered, 

solitary, at small shoulders of geniculate conidiogenous cells, 

caused by sympodial proliferation, or aggregated, forming slight 

swellings of the rachis, i.e., a typical raduliform rachis as in Virgaria 

is lacking. Furthermore, the conidiogenous loci of Pseudovirgaria 

are bulging, convex, slightly attenuated towards the rouded apex, 

in contrast to more cylindrical denticles in Virgaria (Ellis 1971). 

The scar type of Pseudovirgaria is peculiar due to its convex, 

papilla-like shape and reminiscent of conidiogenous loci in plantpathogenic 

genera like Neoovularia U. Braun and Pseudodidymaria 

U. Braun (Braun 1998). The superficially similar genus Veronaea is 

quite distinct from Pseudovirgaria by having erect conidiophores 

with a typical rachis and crowded conidiogenous loci which are 

flat or only slightly prominent and darkened. Pseudovirgaria is 

characterised by its mycelium which is composed of branched 

hyphae with integrated, terminal and intercalary conidiogenous 

cells. A differentiation between branched hyphae and “branched 


conidiophores” is difficult and barely possible. It remains unclear 

if the “creeping threads” and terminal branches of hyphae are 

to be interpreted as “creeping conidiophores”. In any case, the 

mycelium forms complex fertile branched hyphal structures in which 

individual conidiophores are barely discernable. These structures 

and difficulties in discerning individual conidiophores remind one of 

some species of Pseudocercospora Speg. and other cercosporoid 

genera with abundant superficial mycelium in vivo. 

Pseudovirgaria hyperparasitica H.D. Shin, U. Braun, Arzanlou & 

Crous, sp. nov. MycoBank MB504565. Figs 6A, 31. 

Etymology: Named after its hyperparasitic habit on rust fungi. 

Hyphae 1.5–4 µm latae, tenuitunicatae, ≤ 0.5 µm crassae. Cellulae conidiogenae 

15–50 × 2–5 µm, tenuitunicatae (≤ 0.5 µm), cicatricibus (0.5–)1.0(–1.5) µm diam, 

0.5–1 µm altis. Conidia saepe obovoidea, interdum subclavata, 10–20 × 5–9 µm, 

apice rotundato vel paulo attenuato, basi truncata vel rotundata, hilo ca 1 µm 

diam. 

In vivo: Colonies on rust sori, thin to moderately thick, loose, 

cobwebby, to dense, tomentose, pale to medium brown, rusty 

or cinnamon. Mycelium partly immersed in the sori, but mainly 

superficial, composed of a system of branched hyphae with 

integrated conidiogenous cells (fertile threads), distinction between 

conidiophores and vegetative hyphae difficult and barely possible. 

Hyphae 1.5–4 µm wide, hyaline, subhyaline to pale yellowish, 

greenish or very pale olivaceous, light brownish in mass, thin-walled 

(≤ 0.5 µm), smooth, pluriseptate, occasionally slightly constricted 

at the septa. Conidiogenous cells integrated in creeping fertile 

threads, terminal or intercalary, 15–50 µm long, 2–5 µm wide, 

subcylindrical to geniculate, subhyaline to very pale brownish, wall 

thin, ≤ 0.5 µm, smooth, proliferation sympodial, with a single to 

usually several conidiogenous loci per cell, often crowded, causing 

slight swellings, up to 6 µm wide, subdenticulate loci, formed by the 

slightly bulging wall, convex, slightly narrowed towards the rounded 

apex, (0.5–)1.0(–1.5) µm diam and 0.5–1 µm high, wall of the loci 

unthickened, not or slightly darkened-refractive, in surface view 

visible as minute circle (only rim visible and dark). Conidia solitary, 

obovoid, often slightly curved with ± unequal sides, 10–20 × 5–9 

µm, aseptate, subhyaline, pale yellowish greenish to very pale 

olivaceous, wall ≤ 0.5 µm thick, smooth, apex slightly attenuated 

to usually broadly rounded, base rounded to somewhat attenuated 

towards a more or less conspicuous hilum, (0.5–)1(–1.5) µm 

diam, convex to truncate, unthickened, not to slightly darkenedrefractive. 

In vitro: Submerged hyphae hyaline to subhyaline, smooth; aerial 

hyphae smooth, subhyaline, up to 4 µm wide. Conidiogenous cells 

arising imperceptably from aerial vegetative hyphae, terminal, 

occasionally intercalary, holoblastic, proliferating sympodially in a 

geniculate pattern, with more or less long intervals between groups 

of scars; loci slightly darkened, unthickened, approx. 0.5 µm diam. 

Conidia hyaline to subhyaline, aseptate, ovoid, often somewhat 

curved, (10–)13–15(–17) × (5–)6–7(–8) µm, with truncate base 

and acutely rounded apex; hila unthickened, slightly darkenedrefractive. 

Cultural characteristics: Colonies on MEA rather slow-growing, 

reaching 11 mm diam after 14 d at 24 °C, pale to dark mouse-grey, 

velvety, compacted, with colonies being up to 1 mm high. 

Specimens examined: Korea, Seoul, on uredosori of Frommeëlla sp., on Duchesnea 

chrysantha, 17 Sep. 2003, H.D. Shin, paratype, 4/10, CPC 10702–10703 = CBS 

121735–121736, HAL 2053 F; Chunchon, on Phragmidium griseum on Rubus 

crataegifolius, 20 Jul. 2004, H.D. Shin, paratype, 2/8, HAL 2057 F; Suwon, 

87


Fig. 32. Radulidium subulatum (CBS 405.76). A–B. Macronematous conidiophores with sympodially proliferating conidiogenous cells, resulting in a conidium-bearing rachis. 

C–D. Rachis with crowded, blunt conidium-bearing denticles. E. Conidia. Scale bar = 10 µm. 

Fig. 33. Radulidium epichloës (CBS 361.63). A–C. Conidial apparatus at different stages of development, resulting in semi-micronematous conidiophores and sympodially 

proliferating conidiogenous cells. D. Rachis with crowded, blunt conidium-bearing denticles. E–F. Conidiophores with acute branches in the lower part. G. Conidia. Scale bar 

= 10 µm. 

88


on Phragmidium pauciloculare on Rubus parvifolius, 14 Oct. 2003, H.D. Shin, 

paratype, 23/10, HAL 2055 F; Hongchon, on Phragmidium rosae-multiflorae on 

Rosa multiflora, 11 Aug. 2004, H.D. Shin, paratype, 23/8, HAL 2056 F; Yangpyong, 

on Phragmidium sp. on Rubus coreanus, 30 Sep. 2003, H.D. Shin, paratype, 11/10- 

1, CPC 10704–10705 = CBS 121737–121738, HAL 2052 F, and the same locality, 

23 Jul. 2004, HAL 2058 F; Chunchon, on Pucciniastrum agrimoniae on Agrimonia 

pilosa, 7 Oct. 2002, H.D. Shin, holotype, HAL 2054 F, culture ex-type CPC 10753– 

10755 = CBS 121739–121741. 

Radulidium subulatum and Ra. epichloës clade 

Ramichloridium subulatum and R. epichloës form a distinct, wellsupported 

clade with uncertain affinity. This clade is morphologically 

distinct and a new genus is introduced below to accommodate it. 

Radulidium Arzanlou, W. Gams & Crous, gen. nov. MycoBank 

MB504566. 

Etymology: Latin radula = A flexible tongue-like organ in gastropods, 

referring to the radula-like denticles on the rachis. 

Genus ab aliis generibus Ramichloridii similibus denticulis densissimis, prominentibus, 

hebetibus in rachide e cellula conidiogena aculeata orta distinguendum. 

Type species: Radulidium subulatum (de Hoog) Arzanlou, W. Gams 


In vitro: Colonies fast-growing, velvety, floccose near the margin, 

centrally with fertile hyphal bundles up to 10 mm high, about 2 

mm diam, with entire but vague margin; mycelium whitish, later 

becoming greyish brown. Submerged hyphae smooth, thin-walled. 

Conidiophores usually reduced to polyblastic conidiogenous cells 

arising from undifferentiated or slightly differentiated aerial hyphae, 

terminally integrated or lateral, rarely a branched conidiophore 

present, smooth, slightly thick-walled, pale brown, cylindrical to 

acicular, widest at the base and tapering towards the apex; apical 

part forming a pale brown, generally straight rachis, with crowded, 

prominent, blunt denticles, suggesting a gastropod radula; denticles 

0.5–1 µm long, apically pale brown. Conidia solitary, subhyaline, 

thin- or slightly thick-walled, smooth or verrucose, obovoidal, 

fusiform to subcylindrical, base subtruncate and with a slightly 

prominent, conspicuously pigmented hilum; conidial secession 

schizolytic. 

Notes: Radulidium can be distinguished from other ramichloridiumlike 

fungi by its slightly differentiated conidiophores and prominent, 

blunt, very dense conidium-bearing denticles. Although the 

Radulidium clade consists of several subclusters that correlate 

with differences in morphology, the ITS sequence data appear 

insufficient to resolve this species complex. Therefore, only two 

species of Radulidium with clear morphological and molecular 

differences are treated here. The phylogenetic situation of other 

taxa in this clade will be treated in a further study employing a multigene 

approach. 

Radulidium subulatum (de Hoog) Arzanlou, W. Gams & Crous, 

comb. nov. MycoBank MB504567. Figs 10C, 32. 

Basionym: Ramichloridium subulatum de Hoog, Stud. Mycol. 15: 

83. 1977. 

Misapplied name: Rhinocladiella elatior Mangenot sensu dal Vesco 

& B. Peyronel, Allionia 14: 38. 1968. 

In vitro: Submerged hyphae hyaline, thin-walled, 1–2.5 µm wide; 

aerial hyphae brownish. Conidiogenous cells arising laterally from 

vegetative hyphae, pale brown, smooth, thick-walled, sometimes 

without a basal septum, cylindrical to aculeate, tapering gradually 

towards the apex, widest at the base, 25–40 × 2–3 µm; proliferating 

sympodially, forming a pale brown rachis, with densely crowded, 

prominent, blunt conidium-bearing denticles, with pale brown apex. 

Conidia solitary, subhyaline, thin-walled, smooth, ellipsoidal to 

almost clavate, 5–7 × 1.5–2 µm, with a slightly pigmented, nonrefractive 

hilum, about 1 µm diam. 

Cultural characteristics: Colonies on MEA rather fast growing, 

reaching 50 mm diam after 14 d at 24 °C, with entire but vague 

margin, velvety, floccose near the margin, centrally with fertile 

hyphal bundles up to 10 mm high, about 2 mm diam; mycelium 

whitish, later becoming greyish brown; reverse grey, zonate. 

Specimens examined: Czech Republic, on Phragmites australis, A. Samšiňáková, 

ex-type culture CBS 405.76; Opatovicky pond, from Lasioptera arundinis (gall 

midge) mycangia on Phragmites australis, M. Skuhravá, CBS 101010. 

Radulidium epichloës (Ellis & Dearn.) Arzanlou, W. Gams & 

Crous, comb. nov. MycoBank MB504568. Fig. 33. 

Basionym: Botrytis epichloës Ellis & Dearn., Canad. Record Sci. 

9: 272. 1893. 

≡ Ramichloridium epichloës (Ellis & Dearn.) de Hoog, Stud. Mycol. 15: 

81. 1977. 

In vitro: Submerged hyphae hyaline, thin-walled, 1–2.5 µm wide; 

aerial hyphae somewhat darker. Conidiogenous cells arising 

laterally or terminally from undifferentiated or slightly differentiated 

aerial hyphae, occasionally acutely branched in the lower part, 

smooth, thick-walled, pale brown, more or less cylindrical, later 

with thin septa, 25–47 µm long; proliferating sympodially, forming 

a rather short, pale brown, straight or somewhat geniculate 

rachis, with crowded, prominent, blunt denticles with pale brown 

apex. Conidia solitary, subhyaline, rather thin-walled, verruculose, 

obovoidal to fusiform, (4.5–)7–8(–11) × 2–3 µm, with a pigmented 

hilum, 1–1.5 µm diam. 


at 24 °C, with smooth, rather vague, entire margin; velvety, centrally 

floccose and elevated up to 2 mm high; surface mycelium whitish, 

later becoming greyish brown; reverse pale ochraceous. 

Specimen examined: U.S.A., Cranberry Lake, Michigan, isolated from Epichloë 

typhina on Glyceria striata, G.L. Hennebert, CBS 361.63 = MUCL 3124; specimen 

in MUCL designated here as epitype. 

Veronaea-like clade, allied to the Annulatascaceae 

A veronaea-like isolate from Bertia moriformis clusters near the 

Annulatascaceae, and is morphologically distinct from other known 

anamorph genera in the Ramichloridium complex, and therefore a 

new genus is introduced to accommodate it. 

Rhodoveronaea Arzanlou, W. Gams & Crous, gen. nov. MycoBank 

MB504569. 

Etymology: (Greek) rhodon = the rose, referring to the red-brown 

conidiophores, suffix -veronaea from Veronaea. 

Genus ab aliis generibus Ramichloridii similibus basi condiorum late truncata et 

marginata distinguenda. 

In vitro: Colonies slow-growing, velvety, floccose; surface 

olivaceous-grey to dark olivaceous-green; reverse olivaceousblack. 

Hyphae smooth, thin-walled, pale olivaceous. Conidiophores 

arising vertically from creeping hyphae, straight or flexuose, simple, 

thick-walled, red-brown, with inflated basal cell. Conidiogenous 

cells terminally integrated, polyblastic, sympodial, smooth, thick- 


89


Fig. 34. Rhodoveronaea varioseptata (CBS 431.88). A–D. Macronematous conidiophores with sympodially proliferating conidiogenous cells, resulting in conidium bearing rachis 

with slightly prominent conidium-bearing denticles. E–F. Conidia with minute marginal frill. Scale bar = 10 µm. 

Fig. 35. Veronaeopsis simplex (CBS 588.66). A–C. Conidial apparatus at different stages of development, resulting in semi-micronematous conidiophores and sympodially 

proliferating conidiogenous cells. D–E. Rachis with crowded, prominent denticles. F. Intercalary conidiogenous cells. G. Conidia. Scale bar = 10 µm. 

walled, pale brown, rachis straight, occasionally geniculate, with 

crowded, slightly prominent conidium-bearing denticles; denticles 

flat-tipped, slightly pigmented. Conidia solitary, pale brown, thinor 

slightly thick-walled, smooth, ellipsoidal to obovoidal, 0–multiseptate, 

with a protruding base and a marginal basal frill; conidial 


Type species: Rhodoveronaea varioseptata Arzanlou, W. Gams & 

Crous, sp. nov. 

Notes: Rhodoveronaea differs from other ramichloridium-like fungi 

by the presence of a basal, marginal conidial frill, and variably 

septate conidia. 

90


Rhodoveronaea varioseptata Arzanlou, W. Gams & Crous, sp. 

nov. MycoBank MB504570. Figs 10D, 34. 

Etymology: Named for its variably septate conidia. 

Hyphae 2–3 µm latae. Conidiophora ad 125 µm longa et 3–5 µm lata. Cellulae 

conidiogenae 30–70 µm longae et 3–5 µm latae. Conidia 0–2(–3)-septata, (8–)11– 

13(–15) × (2–)3–4(–6) µm. 

In vitro: Submerged hyphae smooth, thin-walled, pale olivaceous, 

2–3 µm wide; aerial hyphae smooth, brownish and slightly narrower. 

Conidiophores arising vertically from creeping hyphae, straight or 

flexuose, simple, smooth, thick-walled, red-brown, up to 125 µm 

long, 3–5 µm wide, often with inflated basal cell. Conidiogenous 

cells terminally integrated, smooth, thick-walled, pale brown at the 

base, paler towards the apex, straight, variable in length, 30–70 

µm long and 3–5 µm wide, rachis straight, occasionally geniculate; 

slightly prominent conidium-bearing denticles, crowded, with slightly 

pigmented apex, about 1 µm diam. Conidia solitary, pale brown, 

thin- or slightly thick-walled, smooth, ellipsoid to obovoid, 0–2(–3)- 

septate, (8–)11–13(–15) × (2–)3–4(–6) µm with a protruding base, 

1.5 µm wide, and marginal frill. 


d at 24 °C, velvety, floccose; surface olivaceous-grey to dark 

olivaceous-green; reverse olivaceous-black. 

Specimen examined: Germany, Eifel, Berndorf, on Bertia moriformis, Sep. 1987, W. 

Gams, holotype CBS-H 19932, culture ex-type CBS 431.88. 

Venturiaceae (Pleosporales) 

The ex-type strain of Veronaea simplex (Papendorf 1969) did not 

cluster with the genus Veronaea (Herpotrichiellaceae), but is allied 

to the Venturiaceae. Veronaea simplex is distinct from species 

of Fusicladium Bonord. by having a well-developed rachis with 

densely aggregated scars. A new genus is thus introduced to 

accommodate this taxon. 

Veronaeopsis Arzanlou & Crous, gen. nov. MycoBank 

MB504571. 

Etymology: The suffix -opsis refers to its similarity with Veronaea. 

Genus Veronaeae simile sed conidiophoris brevioribus (ad 60 μm longis) et rachide 

dense denticulata distinguendum. 

In vitro: Colonies moderately fast-growing; surface velvety, floccose, 

greyish sepia to hazel, with smooth margin; reverse mouse-grey 

to dark mouse-grey. Conidiophores arising vertically from aerial 

hyphae, lateral or intercalary, simple or branched, occasionally 

reduced to conidiogenous cells, pale brown. Conidiogenous 

cells terminally integrated on simple or branched conidiophores, 

polyblastic, smooth, thin-walled, pale brown; rachis commonly 

straight, geniculate, with densely crowded, prominent denticles, 

and slightly pigmented scars. Conidia solitary, subhyaline to pale 

brown, thin- or slightly thick-walled, smooth, oblong-ellipsoidal to 

subcylindrical, (0–)1-septate, with a slightly darkened, thickened, 

hilum; conidial secession schizolytic. 

Type species: Veronaeopsis simplex (Papendorf) Arzanlou & 

Crous, comb. nov. 

Veronaeopsis simplex (Papendorf) Arzanlou & Crous, comb. 

nov. MycoBank MB504572. Figs 17C, 35. 

Basionym: Veronaea simplex Papendorf, Trans. Brit. Mycol. Soc. 

52: 486. 1969. 


In vitro: Submerged hyphae smooth, thin-walled, pale brown; aerial 

hyphae aggregated in bundles. Conidiophores arising vertically 

from aerial hyphae, lateral or intercalary, simple or branched, 

occasionally reduced to conidiogenous cells, pale brown, rather 

short, up to 60 µm long, 1.5–2 µm wide. Conidiogenous cells 

terminally integrated in the conidiophores, smooth, thin-walled, pale 

brown, variable in length, 5–25 µm long, rachis generally straight 

or irregularly geniculate, with crowded, prominent denticles, about 

0.5 µm long, flat-tipped, with slightly pigmented apex. Conidia 

solitary, subhyaline to pale brown, thin- or slightly thick-walled, 

smooth, oblong-ellipsoidal to subcylindrical, (0–)1-septate, slightly 

constricted at the septum, (6–)10–12(–15) × (2–)2.5–3(–4) µm; 

hilum slightly darkened and thickened, not refractive, about 1 µm 

diam. 


d at 24 °C; surface velvety, floccose, greyish sepia to hazel, with 

smooth margin; reverse mouse-grey to dark mouse-grey. 

Specimen examined: South Africa, Potchefstroom, on leaf litter of Acacia karroo, 

1966, M.C. Papendorf, holotype, CBS H-7810; culture ex-type CBS 588.66 = IMI 

203547. 

Notes: The presence of 1-septate conidia in Veronaeopsis overlaps 

with Veronaea. However, Veronaeopsis differs from Veronaea 

based on its conidiophore and conidiogenous cell morphology. 

Veronaea has much longer, macronematous conidiophores than 

Veronaeopsis. Furthermore, Veronaea has a more or less straight 

rachis, whereas in Veronaeopsis the rachis is often geniculate. 

The conidiogenous loci in Veronaea are less prominent, i.e., less 

denticle-like. 

Discussion 

The present study was initiated chiefly to clarify the status of 

Ramichloridium musae, the causal organism of tropical speckle 

disease of banana (Jones 2000). Much confusion surrounded this 

name in the past, relating, respectively, to its validation, species 

and generic status. As was revealed in the present study, however, 

two species are involved in banana speckle disease, namely R. 

musae and R. biverticillatum. Even more surprising was the fact 

that Ramichloridium comprises anamorphs of Mycosphaerella 

Johanson (Mycosphaerellaceae), though no teleomorphs have 

thus far been conclusively linked to any species of Ramichloridium. 

By investigating the Ramichloridium generic complex as outlined 

by de Hoog (1977), another genus associated with leaf spots, 

namely Periconiella, was also shown to represent an anamorph 

of Mycosphaerella. Although no teleomorph connections have 

been proven for ramichloridium-like taxa, de Hoog et al. (1983) 

refer to the type specimen of Wentiomyces javanicus Koord. 

(Pseudoperisporiaceae), on the type specimen of which (PC) 

some ramichloridium-like conidiophores were seen. Without 

fresh material and an anamorph-teleomorph connection proven 

in culture, however, this matter cannot be investigated further. It 

is interesting to note, however, that Wentiomyces Koord. shows 

a strong resemblance to Mycosphaerella, except for the external 

perithecial appendages. 

The genus Mycosphaerella is presently one of the largest genera 

of ascomycetes, containing close to 3 000 names (Aptroot 2006), to 

which approximately 30 anamorph genera have already been linked 

(Crous et al. 2006a, b, 2007). By adding two additional anamorph 

genera, the Mycosphaerella complex appears to be expanding 

91


even further, though some taxa have been shown to reside in other 

families in the Capnodiales, such as Davidiella Crous & U. Braun 

(Davidiellaceae) and Teratosphaeria (Teratosphaeriaceae) (Braun 

et al. 2003, Crous et al. 2007, Schubert et al. 2007 – this volume). 

Another family, which proved to accommodate several 

ramichloridium-like taxa, is the Herpotrichiellaceae (Chaetothyriales). 

Members of the Chaetothyriales are regularly encountered as 

causal agents of human mycoses (Haase et al. 1999, de Hoog 

et al. 2003), whereas species of the Capnodiales are common 

plant pathogens, or chiefly associated with plants. Species in the 

Chaetothyriales have consistently melanized thalli, which is a factor 

enabling them to invade humans, and cause a wide diversity of 

mycoses, such as chromoblastomycosis, mycetoma, brain infection 

and subcutaneous phaeohyphomycosis (de Hoog et al. 2003). The 

only known teleomorph connection in this genus is Capronia Sacc. 

(Untereiner & Naveau 1999). 

Rhinocladiella and Veronaea were in the past frequently 

confused with the genus Ramichloridium. However, Rhinocladiella, 

as well as Veronaea and Thysanorea, were shown to cluster in the 

Chaetothyriales, while Ramichloridium clusters in the Capnodiales. 

Rhinocladiella mackenziei, which causes severe cerebral 

phaeohyphomycosis in humans (Sutton et al. 1998), has in the 

past been confused with Pleurothecium obovoideum (Ur-Rahman 

et al. 1988). Data presented here reveal, however, that although 

morphologically similar, these species are phylogenetically 

separate, with P. obovoideum belonging to the Sordariales, where 

it clusters with sexual species of Carpoligna F.A. Fernández & 

Huhndorf that have Pleurothecium anamorphs (Fernández et al. 

1999). 

In addition to the genera clustering in the Capnodiales and 

Chaetothyriales, several ramichloridium-like genera are newly 

introduced to accommodate species that cluster elsewhere 

in the ascomycetes, namely Pseudovirgaria, Radulidium and 

Myrmecridium, Veronaeopsis, and Rhodoveronaea. Although the 

ecological role of these taxa is much less known than that of taxa in 

the Capnodiales and Chaetothyriales, some exhibit an interesting 

ecology. For instance, the fungicolous habit of Pseudovirgaria, as 

well as some species in Radulidium, which are found on various 

rust species, suggests that these genera should be screened 

further to establish if they have any potential biocontrol properties. 

Furthermore, these two genera share a common ancestor, and 

further work is required to determine whether speciation was shaped 

by co-evolution with the rusts. A further species of “Veronaea” that 

might belong to Pseudovirgaria is Veronaea harunganae (Hansf.) 

M.B. Ellis, which is known to occur on Hemileia harunganae 

Cummins on Harungana in Tanzania and Uganda (Ellis 1976). The 

latter species, however, is presently not known from culture, and 

needs to be recollected to facilitate further study. 

The genera distinguished here represent homogeneous clades 

in the phylogenetic analysis. Only the species of Rhinocladiella are 

dispersed among others morphologically classified in Exophiala or 

other genera. 

By integrating the phylogenetic data generated here with 

the various morphological data sets, we were able to resolve 

eight clades for taxa formerly regarded as representative of the 

Ramichloridium complex. According to the phylogeny inferred 

from 28S rDNA sequence data, the genera Ramichloridium and 

Periconiella were heterogeneous, requiring the introduction of 

several novel genera. Although the present 11 odd genera can 

still be distinguished based on their morphology, it is unlikely that 

morphological identifications without the supplement of molecular 

data would in the future be able to accurately identify all the novel 

isolates that undoubtably await description. The integration of 

morphology with phylogenetic data not only helps to resolve generic 

affinities, but it also assists in discriminating between the various 

cryptic species that surround many of these well-known names that 

are presently freely used in the literature. To that end it is interesting 

to note that for the majority of the taxa studied here, the ITS domain 

(Table 1) provided good species resolution. However, more genes 

will have to be screened in future studies aimed at characterising 

some of the species complexes where the ITS domain provided 

insufficient phylogenetic signal (data not shown) to resolve all of the 

observed morphological species. 

ACKNOWLEDGEMENTS 

The work of Mahdi Arzanlou was funded by the Ministry of Science, Research and 

Technology of Iran, which we gratefully acknowledge. Several colleagues from 

different countries provided material without which this work would not have been 

possible. We thank Marjan Vermaas for preparing the photographic plates, and 

Arien van Iperen for taking care of the cultures. 

REFERENCES 

Al-Hedaithy SSA, Jamjoom ZAB, Saeed ES (1988). Cerebral phaeohyphomycosis 

caused by Fonsecaea pedrosoi in Saudi-Arabia. Acta Pathologica Microbiologica 

Scandinaviae 96 Suppl. 3: 94–100. 




(phytopathogenic hypomycetes). Vol. 2. IHW-Verlag, Eching. 



the teleomorph of Cladosporium s. str. Mycological Progress 2: 3–18. 

Campbell CK, Al-Hedaithy SSA (1993). Phaeohyphomycosis of the brain caused 

by Ramichloridium mackenziei sp. nov. in middle eastern countries. Journal of 

Medical and Veterinary Mycology 31: 325–332. 

Crous PW, Groenewald JZ, Groenewald M, Caldwell P, Braun U, Harrington TC 

(2006a). Species of Cercospora associated with grey leaf spot of maize. 


Crous PW, Groenewald JZ, Risède J-M, Simoneau P, Hyde KD (2006b). Calonectria 

species and their Cylindrocladium anamorphs: species with clavate vesicles. 



A, Burgess T, Barber P, Groenewald JZ (2006c). Phylogenetic lineages in the 


Crous PW, Summerell BA, Carnegie AJ, Mohammed C, Himaman W, Groenewald 

JZ (2007). Foliicolous Mycosphaerella spp. and their anamorphs on Corymbia 

and Eucalyptus. Fungal Diversity 26: 143–185. 







Fernández FA, Lutzoni FM, Huhndorf SM (1999). Teleomorph-anamorph 

connections: the new pyrenomycetous genus Carpoligna and its Pleurothecium 

anamorph. Mycologia 91: 251–262. 

Gams W, Verkleij GJM, Crous PW (eds) (2007). CBS Course of Mycology, 5 th ed. 





Hoog GS de (1977). Rhinocladiella and allied genera. Studies in Mycology 15: 

1–140. 

Hoog GS de (1979). Nomenclatural notes on some black yeast-like hyphomycetes. 

Taxon 28: 347–348. 

Hoog GS de, Rahman MA, Boekhout T (1983). Ramichloridium, Veronaea and 

Stenella: generic delimitation, new combinations and two new species. 

Transactions of the British Mycological Society 81: 485–490. 

Hoog GS de, Vicente V, Caligiorne RB, Kantarcioglu S, Tintelnot KA, Gerrits van 

den Ende AHG, Haase G (2003). Species diversity and polymorphism in 

92


the Exophiala spinifera clade containing opportunistic black yeast-like fungi. 

Journal of Clinical Microbiology 41: 4767–4778. 

Hughes SJ (1951). Studies on Microfungi. 5. Acrotheca. Mycological Papers 38: 

1–8. 

Jones RD (2000). Tropical speckle. In: Disease of Banana, Abaca and Enset (Jones 

RD, ed.). CABI Publishing, Wallingford: 116–120. 

McGinnis MR, Schell WA (1980). The genus Fonsecaea and its relationship to the 

genera Cladosporium, Phialophora, Ramichloridium, and Rhinocladiella. Pan 

American Health Organisation Scientific Publication 396: 215–234. 

Morgan-Jones G (1979). Notes on hyphomycetes. 28. Veronaea bambusae sp. nov. 

Mycotaxon 8: 149–151. 

Morgan-Jones G (1982). Notes on Hyphomycetes. 40. New species of Codinaea 

and Veronaea. Mycotaxon 14: 175–180. 

Mostert L, Groenewald JZ, Summerbell RC, Gams W, Crous PW (2006). Taxonomy 

and pathology of Togninia (Diaporthales) and its Phaeoacremonium anamorphs. 


Nylander JAA (2004). MrModeltest v2.2. Program distributed by the author. 

Evolutionary Biology Centre, Uppsala University. 

Papendorf MC (1969). New South African soil Fungi. Transactions of the British 

Mycological Society 52: 483–489. 

Papendorf MC (1976). Notes on Veronaea including V. compacta sp. nov. Bothalia 

12: 119–121. 


Kew. 



98: 625–634. 

Ronquist F, Huelsenbeck JP (2003). MrBayes 3: Bayesian phylogenetic inference 

under mixed models. Bioinformatics 19: 1572–1574. 

Saccardo PA, Berlese AN (1885). Miscellanea mycologica. Ser. II. Atti dell’Istituto 

Veneto di Scienze, Lettere ed Arti Ser. 6 3: 711–742. 

Schol-Schwarz MB (1968). Rhinocladiella, its synonym Fonsecaea and its relation 

to Phialophora. Antonie van Leeuwenhoek 34: 119–152. 





Seifert KA (1993). Integrating anamorphic fungi into the fungal system. In: The 

Fungal Holomorph: Mitotic, Meiotic and Pleomorphic Speciation in Fungal 

Systematics (Reynolds DR, Taylor JW, eds). International Mycological Institute, 

Egham: 79–85. 

Stahel G (1937). The banana leaf speckle in Surinam caused by Chloridium musae 

nov. spec. and another related banana disease. Tropical Agriculture 14: 42– 

44. 

Sutton DA, Slifkin M, Yakulis R, Rinaldi MG (1998). U.S. case report of cerebral 

phaeohyphomycosis caused by Ramichloridium obovoideum (R. mackenziei): 

Criteria for identification, therapy, and review of other known dematiaceous 

neurotropic taxa. Journal of Clinical Microbiology 36: 708–715. 

Swofford DL (2003). PAUP*. Phylogenetic Analysis Using Parsimony (*and other 

Methods). Version 4. Sinauer Associates, Sunderland, Massachusetts. 

Taylor JW, Jacobson DJ, Kroken S, Kasuga T, Geiser DM, Hibbett DS, Fisher MC 

(2000). Phylogenetic species recognition and species concepts in fungi. Fungal 

Genetics and Biology 31: 21–32. 

Untereiner WA, Naveau FA (1999). Molecular systematics of the Herpotrichiellaceae 

with an assessment of the phylogenetic positions of Exophiala dermatitidis and 


Ur-Rahman N, Mahgoub E, Chagla AH (1988). Fatal brain abscesses caused by 

Ramichloridium obovoideum – Report of three cases. Acta Neurochirurgica 93: 

92–95. 

Veerkamp J, Gams W (1983). Los hongos de Colombia – VIII. Some new species of 

soil fungi from Colombia. Caldasia 13: 709–717. 








Zipfel RD, Beer W de, Jacobs K, Wingfield BD, Wingfield MJ (2006). Multi-gene 

phylogenies define Ceratocystiopsis and Grosmannia distinct from Ophiostoma. 



93


doi:10.3114/sim.2007.58.04 


Cladosporium leaf-blotch and stem rot of Paeonia spp. caused by Dichocladosporium 

chlorocephalum gen. nov. 

K. Schubert 1* , U. Braun 2 , J.Z. Groenewald 3 and P.W. Crous 3 

1 

Botanische Staatssammlung München, Menzinger Strasse 67, D-80638 München, Germany; 2 Martin-Luther-Universität, Institut für Biologie, Geobotanik und Botanischer Garten, 

Herbarium, Neuwerk 21, D-06099 Halle (Saale), Germany; 3 CBS Fungal Biodiversity Centre, P.O. Box 85167, 3508 AD Utrecht, The Netherlands 

*Correspondence: Konstanze Schubert, konstanze.schubert@gmx.de 

Abstract: Cladosporium chlorocephalum (= C. paeoniae) is a common, widespread leaf-spotting hyphomycete of peony (Paeonia spp.), characterised by having dimorphic 

conidiophores. During the season, one stage of this fungus causes distinct, necrotic leaf-blotch symptoms on living leaves of Paeonia spp. In late autumn, winter or after 

overwintering, a second morphologically distinct conidiophore type occurs on dead, blackish, rotting stems. Conspecificity of the two morphs, previously proposed on the basis 

of observations in culture, was supported by DNA sequence data from the ITS and LSU gene regions, using cultures obtained from leaf-blotch symptoms on living leaves, as 

well as from dead stems of Paeonia spp. Sequence data were identical, indicating a single species with two morphs. On account of its distinct conidiogenous loci and conidial 

hila, as well as its sequence-based phylogenetic position separate from the Davidiella/Cladosporium clade, the peony fungus has to be excluded from Cladosporium s. str., but 

still belongs to the Davidiellaceae (Capnodiales). The leaf-blotching (cladosporioid) morph of this fungus morphologically resembles species of Fusicladium, but differs in having 

dimorphic fruiting, and is phylogenetically distant from the Venturiaceae. The macronematous (periconioid) morph resembles Metulocladosporiella (Chaetothyriales), but lacks 

rhizoid conidiophore hyphae, and has 0–5-septate conidia. Hence, C. chlorocephalum is assigned to the new genus Dichocladosporium. 

Taxonomic novelties: Dichocladosporium K. Schub., U. Braun & Crous, gen. nov., Dichocladosporium chlorocephalum (Fresen.) K. Schub., U. Braun & Crous, comb. nov. 

Key words: Anamorphic fungi, Cladosporium chlorocephalum, C. paeoniae, hyphomycetes, new genus, phylogeny, taxonomy. 

Introduction 

Fresenius (1850) described Periconia chlorocephala Fresen. from 

Germany on dead stems of Paeonia sp. Mason & Ellis (1953) 

examined this species in vitro and in vivo and stated that it only 

occurred on dead stems of Paeonia spp. They described, illustrated 

and discussed this species in detail, and reallocated it to the genus 

Cladosporium Link. 

A second, cladosporioid hyphomycete on Paeonia spp., 

Cladosporium paeoniae Pass., was collected by Passerini on living 

leaves of P. albiflora (as P. edulis) in Italy, and distributed in Thümen, 

Herbarium mycologicum oeconomicum, Fasc. IX, No. 416 (1876), 

together with the first valid description, which was repeated by 

Passerini (1876). Later, Passerini collected this fungus on Paeonia 

officinalis at Parma in Italy and distributed it in Thümen, Mycotheca 

universalis, No. 670 (1877). Saccardo (1882) listed a collection of 

this species on Paeonia anomala from Russia, Siberia, which he 

later described as Cladosporium paeoniae var. paeoniae-anomalae 

Sacc. (Saccardo 1886). A first examination of C. paeoniae in culture 

was accomplished by Meuli (1937), followed by a treatment in vitro 

by de Vries (1952). Mason & Ellis (1953) described and illustrated in 

their treatment of C. chlorocephalum macroconidiophores, agreeing 

with those of the original diagnosis and illustration of Periconia 

chlorocephala, as well as semi-macronematous conidiophores 

concurring with those of C. paeoniae, although no mention was 

made of the latter name. McKemy & Morgan-Jones (1991) carried 

out comprehensive studies on Cladosporium on Paeonia spp. in 

vitro and in vivo, including detailed discussions of the history of 

the taxa concerned, taxonomic implications and comprehensive 

descriptions and illustrations. They concluded that Cladosporium 

paeoniae, found in culture together with C. chlorocephalum, was 

a semi-macronematous form (synanamorph) of the latter species, 

and reduced C. paeoniae to synonymy with the latter species. 

In the present study, re-examination and reassessment of 

morphological characters, conidiogenesis, and DNA sequence 

data of the ITS and 28S nrDNA were used to confirm the identity 

of Cladosporium chlorocephalum (the periconioid morph) and C. 

paeoniae (the cladosporioid morph), and clarify their relation to 

Cladosporium s. str. (Davidiellaceae) (emend. David 1997, Braun 

et al. 2003). 


Isolates 

Single-conidial isolates were obtained from symptomatic leaves 

and dead stems, and cultured as detailed in Crous (1998). Cultural 

characteristics and morphology of isolates (Table 1) were recorded 

from plates containing either 2 % potato-dextrose agar (PDA) or 

synthetic nutrient-poor agar (SNA) (Gams et al. 2007). Plates were 

incubated at 25 °C under continuous near-UV light to promote 

sporulation. 

DNA isolation, amplification and sequencing 


DNA was isolated following the protocol of Lee & Taylor (1990). 

The primers V9G (de Hoog & Gerrits van den Ende 1998) and LR5 

(Vilgalys & Hester 1990) were used to amplify part (ITS) of the nuclear 

rDNA operon spanning the 3’ end of the 18S rRNA gene (SSU), the 

first internal transcribed spacer (ITS1), the 5.8S rRNA gene, the 

second ITS region and the 5’ end of the 28S rRNA gene (LSU). 

The primers ITS4 (White et al. 1990), LR0R (Rehner & Samuels 

1994), LR3R (www.biology.duke.edu/fungi/mycolab/primers.htm) 

and LR16 (Moncalvo et al. 1993) were used as internal sequence 

primers to ensure good quality sequences over the entire length 

of the amplicon. The PCR conditions, sequence alignment and 

subsequent phylogenetic analysis followed the methods of Crous 

et al. (2006b). Gaps longer than 10 bases were coded as single 

events for the phylogenetic analyses; the remaining gaps were 

treated as new character states. Sequence data were deposited 

in GenBank (Table 1) and alignments in TreeBASE (www.treebase. 

org). 

95

Schubert et al. 

Morphology 

Morphological examinations were made from herbarium samples, 

fresh symptomatic leaves and stems, as well as cultures sporulating 

on SNA. Structures were mounted in water or Shear’s solution 

(Dhingra & Sinclair 1985), and 30 measurements at × 1 000 

magnification were made of each structure under an Olympus BX 

50 microscope (Hamburg, Germany). The 95 % confidence levels 

were determined and the extremes of spore measurements given 

in parentheses. Scanning electron microscopic examinations were 

conducted at the Institute of Zoology, Martin-Luther-University, 

Halle (Saale), Germany, using a Hitachi S-2400. Samples were 

coated with a thin layer of gold applied with a sputter coater SCD 

004 (200 s in an argon atmosphere of 20 mA, 30 mm distant 

from the electrode). Colony colours were noted after 2 wk growth 

on PDA at 25 °C in the dark, using the colour charts of Rayner 

(1970). All cultures studied were deposited in the culture collection 

of the Centraalbureau voor Schimmelcultures (CBS), Utrecht, 

the Netherlands (Table 1). Taxonomic novelties were lodged with 

MycoBank (www.MycoBank.org). 

Results 

DNA phylogeny 

Amplification products of approximately 1 700 bases were obtained 

for the isolates listed in Table 1. The ITS region of the sequences 

was used to obtain additional sequences from GenBank which were 

added to the alignment. The manually adjusted alignment contained 

26 sequences (including the two outgroup sequences) and 518 

characters including alignment gaps. Of the 518 characters used 

in the phylogenetic analysis, 226 were parsimony-informative, 33 

were variable and parsimony-uninformative, and 259 were constant. 

Neighbour-joining analysis using three substitution models on the 

sequence data yielded trees supporting the same clades but with 

a different arrangement at the deeper nodes. These nodes were 

supported poorly in the bootstrap analyses (the highest value 

observed for one of these nodes was 64 %; data not shown). 

Two equally most parsimonious trees (TL = 585 steps; CI = 

0.761; RI = 0.902; RC = 0.686), one of which is shown in Fig. 1, 

was obtained from the parsimony analysis of the ITS region. The 

Dichocladosporium K. Schub., U. Braun & Crous isolates formed a 

well-supported clade (100 % bootstrap support), distinct from clades 

containing species of Davidiella Crous & U. Braun, Mycosphaerella 

Johanson and Teratosphaeria Syd. & P. Syd. This placement was 

also supported by analyses of the first part of the 28S rRNA gene 

(see Crous et al. 2007 – this volume). 

Taxonomy 

Because conidia formed holoblastically in simple or branched 

acropetal chains, Cladosporium chlorocephalum and C. paeoniae 

coincided with previous concepts of Cladosporium s. lat. (Braun 

et al. 2003, Schubert 2005), belonging to a wide assemblage of 

genera classified by Kiffer & Morelet (1999) as “Acroblastosporae”. 

Previous studies conducted in vitro concluded that Cladosporium 

chlorocephalum and C. paeoniae represent two developmental 

stages (morphs) of a single species, a result confirmed here by 

DNA sequence analyses. A detailed analysis of conidiogenesis, 

structure of the conidiogenous loci and conidial hila, and a 

comparison with Cladosporium s. str., typified by C. herbarum 

(Pers. : Fr.) Link, revealed obvious differences: The conidiogenous 

loci and conidial hila of C. chlorocephalum are quite distinct from 

those of Cladosporium s. str. by being denticulate or subdenticulate, 

apically broadly truncate, unthickened or slightly thickened, but 

somewhat darkened-refractive. The scars in Cladosporium s. 

str. are, however, characteristically coronate, i.e., with a central 

convex dome surrounded by a raised periclinal rim (David 1997, 

Braun et al. 2003, Schubert 2005). Hence, the peony fungus 

has to be excluded from Cladosporium s. str. A comparison with 

phaeoblastic hyphomycetous genera revealed a close similarity of 

this fungus with the genus Metulocladosporiella Crous, Schroers, 

J.Z. Groenew., U. Braun & K. Schub. recently introduced for the 

Cladosporium speckle disease of banana (Crous et al. 2006a). 

Both fungi have dimorphic fruiting, pigmented macronematous 

conidiophores often with distinct basal swellings and densely 

branched terminal heads composed of short branchlets and 

ramoconidia, denticulate or subdenticulate unthickened, but 

somewhat darkened-refractive conidiogenous loci, as well as 

phaeoblastic conidia, formed in simple or branched acropetal 

chains. The semi-macronematous leaf-blotching morph is close 

to and barely distinguishable from Fusicladium Bonord. However, 

unlike Metulocladosporiella, the peony fungus does not form rhizoid 

hyphae at the base of conidiophore swellings and the conidia are 

amero- to phragmosporous [0–5-septate versus 0(–1)-septate 

in Metulocladosporiella]. Furthermore, the peony fungus neither 

clusters within the Chaetothyriales (with Metulocladosporiella) 

nor within the Venturiaceae (with Fusicladium), but clusters basal 

to the Davidiellaceae (see also Crous et al. 2007 – this volume). 

Hence, we propose to place C. chlorocephalum in the new genus 

Dichocladosporium. 

Dichocladosporium K. Schub., U. Braun & Crous, gen. nov. 

MycoBank MB504428. Figs 2–5. 

Etymology: dicha in Greek = twofold. 

Differt a Metulocladosporiella conidiophoris cum cellulis basalibus saepe inflatis, 

sed sine hyphis rhizoidibus, conidiis amero- ad phragmosporis (0–5-septatis). 

Type species: Dichocladosporium chlorocephalum (Fresen.) K. 

Schub., U. Braun & Crous, comb. nov. 

Dichocladosporium chlorocephalum (Fresen.) K. Schub., U. 

Braun & Crous, comb. nov. MycoBank MB504429. Figs 2–5. 

Basionym: Periconia chlorocephala Fresen., Beiträge zur Mykologie 

1: 21. 1850. 

≡ Haplographium chlorocephalum (Fresen.) Grove, Sci. Gossip 21: 198. 

1885. 

≡ Graphiopsis chlorocephala (Fresen.) Trail, Scott. Naturalist (Perth) 10: 

75. 1889. 

≡ Cladosporium chlorocephalum (Fresen.) E.W. Mason & M.B. Ellis, 

Mycol. Pap. 56: 123. 1953. 

= Cladosporium paeoniae Pass., in Thümen, Herb. Mycol. Oecon., Fasc. IX, 

No. 416. 1876, and in Just´s Bot. Jahresber. 4: 235. 1876. 

= Periconia ellipsospora Penz. & Sacc., Atti Reale Ist. Veneto Sci. Lett. Arti, 

Ser. 6, 2: 596. 1884. 

= Cladosporium paeoniae var. paeoniae-anomalae Sacc., Syll. Fung. 4: 351. 

1886. 

= Haplographium chlorocephalum var. ovalisporum Ferraris, Fl. Ital. Cryptog., 

Hyphales: 875. 1914. 

Descriptions: Mason & Ellis (1953: 123–126), McKemy & Morgan- 

Jones (1991: 140–144), Schubert (2005: 216). 

Illustrations: Fresenius (1850: Pl. IV, figs 10–15), Mason & Ellis 

(1953: 124–125, figs 42–43), McKemy & Morgan-Jones (1991: 

137, fig. 1; 141, fig. 2; 139, pl. 1; 143, pl. 2), Schubert (2005: 217, 

fig. 113; 275, pl. 34). 

96

Dichocladosporium gen. nov. 

10 changes 

Botryosphaeria ribis DQ316076 

Botryosphaeria stevensii AY343484 

100 

97 

70 

100 

100 

100 

97 

100 

100 

69 

Cladosporium tenellum CPC 12053 

Cladosporium variabile CPC 12753 

Cladosporium herbarum CPC 11600 

Cladosporium macrocarpum CBS 299.67 

Cladosporium bruhnei CPC 12211 

Cladosporium ossifragi CBS 842.91 

Cladosporium iridis CBS 138.40 

CPC 11383 

CPC 11969 

CBS 213.73 

CBS 100405 

100 

91 


Passalora arachidicola AF297224 

Mycosphaerella aurantia AY509744 

Passalora fulva AF393701 

Cercospora apii AY840512 

Cercospora beticola AY840527 

Readeriella mirabilis AY725529 


Teratosphaeria readeriellophora AY725577 

Teratosphaeria microspora AY260098 

Trimmatostroma abietis AY559362 

Trimmatostroma abietina AJ244267 

Trimmatostroma salicis AJ244264 

Dichocladosporium chlorocephalum 

Fig. 1. One of two equally most parsimonious trees obtained from a heuristic search with 100 random taxon additions of the ITS sequence alignment. The scale bar shows 

10 changes, and bootstrap support values from 1 000 replicates are shown at the nodes. Thickened lines indicate the strict consensus branches. The tree was rooted to two 

Botryosphaeria species. 

Characters of the cladosporioid morph: Leaf-blotch symptoms on 

living leaves amphigenous, variable in shape and size, subcircularoval 

to irregular, broad, oblong to expanded, up to 30 mm long 

and 20 mm wide, at times covering the entire leaf surface, forming 

olivaceous-brown to blackish brown patches, rarely violet-brown, 

margin usually indefinite, attacked areas turning dry with age, 

also occurring on young, green stems. Colonies amphigenous, 

punctiform to effuse, loose to dense, caespitose, brown, villose. 

Mycelium immersed, subcuticular to intraepidermal; hyphae 

sparingly branched, 4–7(–10) µm wide, septate, sometimes with 

swellings and constrictions, swollen cells up to 13 µm diam, 

subhyaline to pale brown, smooth, walls thickened, hyphae 

sometimes aggregated; in vitro mycelium at first mainly immersed, 

later also superficial, branched, 1–5(–7) µm wide, pluriseptate, 

often constricted at septa and with swellings and constrictions, 

therefore irregular in outline, smooth to verruculose or irregularly 

rough-walled, loosely verruculose with distinct large warts. Semimacronematous 

conidiophores formed on leaf-blotches solitary 

or in small, loose groups, arising from internal hyphae or swollen 

hyphal cells, erumpent through the cuticle, occasionally emerging 

through stomata, erect, straight to somewhat flexuous, oblongcylindrical, 

usually unbranched or occasionally branched, 13–80 

(–120) × (4–)5–8(–10) µm, slightly attenuated towards the apex, 

septate, septa often dense, unconstricted, pale to medium brown, 

sometimes paler towards the apex, smooth, thick-walled, wall often 

with two distinct layers, often somewhat inflated at the very base, 

up to 14 µm diam, occasionally proliferating enteroblastically; in 

vitro conidiophores arising laterally from plagiotropous hyphae or 

terminally from ascending hyphae, the latter usually appearing 

more filiform than those arising laterally from plagiotropous hyphae, 

erect, straight to slightly flexuous, cylindrical-oblong, not geniculate, 

usually unbranched, rarely with a short lateral prolongation near 

the apex, 18–60(–100) × 3–6 µm, slightly attenuated towards 

the apex, septate, pale to medium brown or olivaceous-brown, 

smooth to asperulate, walls somewhat thickened. Conidiogenous 

cells integrated, terminal or intercalary, subcylindrical, 7–45 µm 


97


Fig. 2. Dichocladosporium chlorocephalum (HAL 1924 F), periconioid, stem rotting morph. Conidiophores and conidia. Scale bar = 10 µm. 

Table 1. Isolates subjected to DNA analysis and morphological examination. 

Species Accession number 1 Host Country Collector GenBank accession 

number 

Dichocladosporium chlorocephalum CBS 213.73; IMI 048108a Paeonia sp. United Kingdom F. Rilstone EU009455 

CBS 100405 Paeonia sp. New Zealand M. Braithwaite EU009456 

CBS 121522; CPC 11383 Paeonia delavayi Germany K. Schubert EU009457 

CBS 121523*; CPC 11969 Paeonia officinalis Germany K. Schubert EU009458 

1 

CBS: Centraalbureau voor Schimmelcultures, Utrecht, The Netherlands; CPC: Culture collection of Pedro Crous, housed at CBS; IMI: International 

Mycological Institute, CABI-Bioscience, Egham, Bakeham Lane, U.K. 


98


Fig. 3. Dichocladosporium chlorocephalum (CBS 121523 = CPC 11969). A–B. Symptoms of the periconioid, stem rotting morph. C–E, H. Macroconidiophores and conidia. F–G. 

Semi-macronematous conidiophores and conidia. I–J. Ramoconidia and conidia. K–M. Scanning electron microscopic photographs. K. Conidiophores. L. Conidial chain. M. 

Single conidium showing the surface ornamentation and scar structure. Scale bars: C–J, L = 10 µm; K = 100 µm; M = 2 µm. 


99


Fig. 4. Dichocladosporium chlorocephalum (HAL 2011 F), cladosporioid, leaf-spotting morph. Conidiophores and conidia. Scale bar = 10 µm. 

long, proliferation sympodial, with one to several conidiogenous 

loci, subdenticulate or denticulate, protuberant, terminally broadly 

truncate, 1.5–3 µm wide, unthickened or almost so, somewhat 

darkened-refractive. Conidia catenate, in simple or branched 

chains, polymorphous, small conidia globose, subglobose, broadly 

obovoid, 3–9 × 3–5 µm, aseptate, pale to medium brown, smooth, 

intercalary conidia limoniform, ellipsoid-fusiform, oblong, 5–23 

× 3.5–6.5 µm, 0–2-septate, medium brown, smooth to minutely 

verruculose or irregularly rough-walled, large conidia ellipsoid, 

oblong-cylindrical, ampulliform, 22–45(–52) × (4.5–)5–8 µm, 

0–5-septate, medium brown, smooth to minutely verruculose or 

irregularly rough-walled, walls somewhat thickened, hila truncate, 

1–3 µm wide, unthickened or almost so, somewhat darkenedrefractive; 

occasionally with microcyclic conidiogenesis; in vitro 

numerous, polymorphous, catenate, in loosely branched chains, 

small conidia globose, subglobose, or obovoid, 3–8 × 3–4 µm, 

aseptate, intercalary conidia limoniform to ellipsoid-fusiform, 9–18 

× 3.5–4.5 µm, 0–1-septate, large conidia ellipsoid to cylindricaloblong, 

14–30(–38) × 3–6(–7) µm, 0–3-septate, pale to medium 

brown, asperulate, minutely verruculose to irregularly roughwalled, 

walls thickened, hila usually short denticle-like, protuberant, 

truncate, in smaller conidia 0.5–1.8 µm wide, in larger conidia 

(1.5–)2–3 µm wide, unthickened or almost so but usually darkenedrefractive; 

with occasional microcyclic conidiogenesis. 

Characters of the periconioid morph: Macronematous conidiophores 

formed on faded or dead stems in late autumn, winter or after 

overwintering; colonies at first visible as reddish brown streaks, later 

turning olivaceous-brown to black, sometimes linear, sometimes 

encircling the stems, often occupying large stem segments, effuse, 

densely caespitose, velvety. Mycelium immersed, subcuticular 

100


to intraepidermal; hyphae at first sparsely branched, 3–7 µm 

wide, septate, not constricted at the septa, becoming swollen 

and wider, up to 11 µm wide, often branched, pale to medium 

olivaceous-brown, walls thickened, forming loose to dense hyphal 

aggregations; in vitro mycelium immersed to superficial, loosely 

branched, 2–6(–7) µm wide, pluriseptate, usually without swellings 

and constrictions, subhyaline to medium brown or olivaceousbrown, 

almost smooth to asperulate or irregularly rough-walled, in 

older colonies on PDA up to 10 µm wide, sometimes single hyphal 

cells distinctly swollen, up to 16(–20) µm wide, mainly at the base 

of conidiophores, sometimes covered by a slime coat or enveloped 

in a polysaccharide-like layer. Stromata well-developed, large and 

expanded, up to about 50–320 µm in length, 15–30 µm deep, 

composed of a single to several layers of swollen pale to medium 

brown stromatic cells, 5–18 µm diam, thick-walled. Conidiophores 

solitary or in loose groups, arising from swollen hyphal cells or 

stromata, erumpent through the cuticle, erect, straight, rigid to 

slightly flexuous, 150–680 µm long, composed of a subcylindrical 

stipe, 13–24 µm wide at the base, slightly attenuated towards 

the apex, 5–15 µm just below the branched head, pluriseptate, 

not constricted at the septa, young conidiophores pale medium 

olivaceous-brown, later medium to usually dark brown, sometimes 

slightly paler at the distal end, smooth or almost so, often appearing 

somewhat granular, roughened, walls distinctly thickened, 1.5–3(– 

4) µm wide; apex with a roughly subglobose to ovoid head, about 

35–70 µm diam, composed of dense branchlets and ramoconidia, 

primary branchlets close to the apex and below the first and 

sometimes second and third septa, solitary, in pairs or small verticils, 

appressed against the stipe or somewhat divergent, subcylindrical 

to ellipsoid-oval, aseptate, rarely 1-septate, pale olivaceous to dark 

brown, 10–20 × 5–8.5 µm; in vitro conidiophores initially microand 

semimacronematous, then progressively macronematous 

as colonies age, arising laterally from plagiotropous hyphae or 

terminally from ascending hyphae, sometimes also from swollen 

hyphal cells; micronematous conidiophores filiform, narrowly 

cylindrical-oblong, unbranched, up to 150 µm long, 2–3.5 µm wide, 

septate, septa often appear to be darkened, pale to pale medium 

olivaceous-brown, asperulate, walls slightly thickened; semimacronematous 

conidiophores often resembling those formed 

by the leaf-blotching (cladosporioid) morph on the natural host, 

subcylindrical to cylindrical-oblong, straight to slightly flexuous, 

unbranched, rarely branched, (10–)15–120 × 3–5(–6) µm, slightly 

attenuated towards the apex, septate, medium brown, minutely 

verruculose to irregularly rough-walled, walls more or less thickened; 

macronematous conidiophores formed in older cultures on SNA, 

PDA and also MEA (according to McKemy & Morgan-Jones 1991), 

but more prominent on PDA and MEA, resembling those formed 

by the stem-rotting morph (i.e., the periconioid morph, in planta), 

consisting of a long unbranched stipe and a subglobose head, but 

in culture the heads are often more loosely branched than on the 

natural substratum, not always forming a compact head, up to 580 

µm long, 5–13 µm wide, attenuated towards the apex, 4–8 µm 

just below the branched upper part, somewhat swollen at the base, 

septate, medium to very dark brown, minutely verruculose, walls 

distinctly thickened, two distinct wall layers visible, 1–2 µm thick. 

Conidiogenous cells holoblastic, integrated, terminal, intercalary or 

even discrete, ellipsoid to cylindrical or doliiform, subdenticulate, 

proliferation sympodial, multilocal, conidiogenous loci truncate, 

flat, unthickened, 1–3 µm wide, somewhat darkened-refractive; in 

culture conidiogenous loci appearing to be somewhat thickened 

and distinctly darkened-refractive, 1–2.5(–3) µm wide. Conidia 

catenate, in long, branched chains, straight, subglobose, aseptate, 

3.5–7 µm diam, or ellipsoid-ovoid, 6–15 × 4–9 µm, 0(–1)-septate, 

pale olivaceous to olivaceous-brown, smooth to verruculose (under 

the light microscope), hila flat, truncate, unthickened, (0.5–)1– 

2(–2.5) µm wide, not darkened, but somewhat refractive; in vitro 

conidia numerous, catenate, formed in long, branched chains, small 

conidia globose to subglobose, (2–)3–7 × (2–)3–4 µm, aseptate, 

intercalary ones ellipsoid-ovoid, 6–16 × 3.5–5 µm, 0(–1)-septate, 

secondary ramoconidia ellipsoid to cylindrical-oblong, (13–) 

15–34(–47) × (3–)4–6(–7) µm, 0–2-septate, sometimes slightly 

constricted at the septa, medium olivaceous-brown, verruculose or 

irregularly rough-walled, walls slightly to distinctly thickened, hila 

more or less protuberant, subdenticulate to denticulate, in small and 

intercalary conidia 0.5–1(–1.5) µm, in secondary ramoconidia 1–2.5 

(–3) µm, unthickened or somewhat thickened, darkened-refractive; 

occasional microcyclic conidiogenesis. 

Cultural characteristics: Colonies on PDA at first whitish or smoke 

grey, reverse smoke-grey to olivaceous-grey, with age smoke-grey 

to olivaceous or olivaceous-grey, sometimes even dark mouse-grey, 

reverse iron-grey to dark mouse-grey or black, felty; margin white 

to smoke-grey, narrow to more or less broad, regular to slightly 

undulate, glabrous to somewhat feathery; aerial mycelium at first 

mainly in the colony centre, with age abundantly formed, covering 

almost the whole colony, whitish, smoke-grey to olivaceous, felty; 

growth low convex to raised; numerous small exudates formed, 

sometimes becoming prominent; fertile. 

Specimens examined: Czechoslovakia, Bohemia, Turnau, on leaves of Paeonia 

arborea, 15 Sep. 1905, J.E. Kabát, Kabát & Bubák, Fungi Imperf. Exs. 396, B 70- 

6669. France, on dead stems of Paeonia sp., 1901, ex Herbario Musei Parisiensis, 

ex herb. Magnus, exs. Desmazières, Pl. Crypt. N. France, Ed. 2, Ser. 1, 1621, HBG, 

as “Periconia atra”; Chailly-en-Biere, Seine-et-Marne, Feuilleaubois, on stems of 

P. officinalis, 27 Mar. 1881, Roumeguère, Fungi Sel. Gall. Exs. 1803, HBG, as 

“Periconia atra”. Germany, Baden-Würtemberg, Kreis Tübingen, Drusslingen, on 

leaves of P. officinalis, Jun. 1935, Raabe, B 70-6670; Bayern, Freising, on leaves 

of P. officinalis, Sep. 1918, Prof. Dr. J.E. Weiß, Herbarium pathologicum, B 70- 

6663; Brandenburg, Schloßpark zu Tamsel, on leaves of P. officinalis, 15 Aug. 

1924, P. Vogel, Sydow, Mycoth. Germ. 2447, M-57751, PH; Triglitz, on leaves of 

P. officinalis, 3 Oct. 1909, Jaap, B 70-6668; Hessen, Frankfurt am Main, botanical 

garden, on leaves of P. potaninii, 7 Oct. 2004, R. Kirschner, HAL, RoKi 2222; Kreis 

Kassel, Hofgeismar, Garten von Prof. Grupe, on leaves of P. officinalis, 3 Sep. 

1947, Schulz, B 70-6658; Mecklenburg-Vorpommern, Rostock, neuer botanischer 

Garten, on leaves of P. corallina (= P. mascula), 27 Aug. 1950, Becker, B 70-6662; 

Nordrhein-Westfalen, Duisburg, Dinslake, private garden, on leaves of P. anomala, 

9 Aug. 2005, N. Ale-Agha, HAL 2014 F; Hamborn, botanical garden, on leaves of P. 

obovata, 10 Aug. 2005, N. Ale-Agha, HAL 2017 F; Essen, botanical garden of the 

university of Essen, on leaves of P. mlokosewitschii, 10 Aug. 2005, N. Ale-Agha, 

HAL 2013 F; on leaves of P. officinalis and P. suffruticosa, 11 Aug. 2005, N. Ale- 

Agha, HAL 2016, 2017 F; Sachsen, Königstein, in Gärten, verbreitet, on leaves of 

P. officinalis, Aug. 1896, W. Krieger, Krieger, Fungi Saxon. Exs. 1545, M-57749; 

Aug., Sep. 1896, 1915, W. Krieger, Krieger, Schädliche Pilze, B 70-6666, 70-6667; 

Sachsen-Anhalt, Halle (Saale), Botanical Garden, on leaves of P. delavayi, 22 

Jun. 2004, K. Schubert, HAL 2011 F, culture deposited at the CBS, CBS 121522 = 

CPC 11383; on leaves of P. officinalis, 22 Jun. 2004, K. Schubert, HAL 2012 F; on 

stems of P. officinalis, 16 Mar. 2005, K. Schubert, neotype of Dichocladosporium 

chlorocephalum designated here HAL 1924 F, isoneotype CBS-H 19869, culture 

ex-neotype CBS 121523 = CPC 11969; on dead stems of Paeonia sp., Jan. 1873, 

G. Winter, Rabenhorst, Fungi Eur. Exs. 1661, HBG, as “Periconia chlorocephala”; 

Thüringen, Fürstlicher Park zu Sondershausen, on leaves of P. arborea, 20 Aug. 

1903, G. Oertel, Sydow, Mycoth. Germ. 196, PH. Italy, Thümen, Herb. Mycol. 

Oecon. 416, on living leaves of Paeonia lactiflora [= P. edulis] (M-57753), lectotype 

of “Cladosporium paeoniae” designated here; isolectotypes: Thümen, Herb. 

Mycol. Oecon. 416; Padova, on leaves of P. officinalis, Aug. 1902, P.A. Saccardo, 

Saccardo, Mycoth. Ital. 1186, B 70-6660, SIENA; Parma, on leaves of P. officinalis, 

Jul. 1876, Prof. Passerini, Thümen, Mycoth. Univ. 670, B 70-6654, 70-6655, M- 

57752; Pavia, botanical garden, on leaves of P. officinalis, summer 1889, Briosi & 

Cavara, Fung. Paras. Piante Colt. Utili Ess. 78, M-57748; F. Cavara, Roumeguère, 

Fungi Sel. Gall. Exs. 5193, mixed infection with Cladosporium herbarum, B 70- 

6656; Siena, Hort. Bot., on leaves of Paeonia sp., Nov. 1899, SIENA. Latvia, prov. 

Vidzeme, Kreis Riga, Riga, in a garden, on leaves of P. foemina [= P. officinalis], 28 

Aug. 1936, J. Smarods, Fungi Lat. Exs. 799, M-57747. New Zealand, isolated from 


101


Fig. 5. Dichocladosporium chlorocephalum (CBS 121522 = CPC 11383). A–B. Symptoms on leaves of Paeonia officinalis and P. delavayi caused by the cladosporioid, leaf spotting 

morph. C–D. Conidiophores and conidia. E. Ramoconidia and conidia. F–G. Scanning electron microscopic photographs. F. Conidial chain still attached to a conidiophore. 

G. Conidia showing surface ornamentation and scar structure. Scale bars: C–E, G = 10 µm; F = 5 µm. 

red leaf and stem lesions on Paeonia sp., M. Braithwaite, CBS 100405. Romania, 

Râmnicu-Vâlcea, distr. Vâlcea, Oltenia, on leaves of P. officinalis, 17 Aug. 1930, 

Tr. Săvulescu & C. Sandu, Săvulescu, Herb. Mycol. Roman. 298, M-57742. U.K., 

England, Cornwall, Lambounce Hill, Perranzubuloe, isolated from dead stems of 

Paeonia sp., isol. F. Rilstone, CBS 213.73 = IMI 048108a. U.S.A., on leaves of 

Paeonia sp., Sep. 1878, Ellis, N. Amer. Fungi 543, B 70-6659, M-57744, PH; Illinois, 

Cobden, on leaves of Paeonia sp., 8 Aug. 1882, F.S. Earle, No. 91, B 70-6657; 

Kansas, Topeka, on leaves of P. officinalis, 7 Jul. 1922, C.F. Menninger, US Dept. 

Agric., Pathol. Mycol. Coll. 60085, B 70-6661, F; Montana, Columbia, on leaves of 

P. officinalis, Aug. 1886, B.T. Galloway, Ellis & Everh., N. Amer. Fungi Ser. II, 1991, 

PH; on leaves of Paeonia sp., 18 Oct. 1931, W.E. Maneval, F. 

Host range and distribution: On Paeonia anomala, P. arborea, P. 

delavayi, P. hybrida, P. lactiflora, P. mascula, P. mlokosewitschii, 

P. moutan, P. obovata, P. obovata var. willmotiae, P. officinalis, 

P. potaninii, P. suffruticosa, Paeonia spp. (Paeoniaceae), Asia 

(Armenia, China, Georgia, Kazakhstan, Russia), Europe (Belgium, 

102


Czechoslovakia, Denmark, France, Germany, Italy, Latvia, Moldova, 

Poland, Romania, Switzerland, U.K., Ukraine), North America 

(Canada, U.S.A.), New Zealand. 

Notes: Type material of Periconia chlorocephala is not preserved 

in the herbarium of G. Fresenius at FR (Forschungsinstitut 

Senkenberg, Frankfurt a. M., Germany). Hence, a new specimen 

collected in the Botanical Garden of the Martin-Luther-University 

Halle (Saale), Germany, is proposed to serve as neotype. A culture 

derived from this collection is deposited at the CBS, Utrecht, the 

Netherlands as ex-neotype culture. A leaf-blotch sample, also 

collected in the Botanical Garden at Halle (Saale), from which we 

also derived a living culture, is designated as representative of the 

synanamorph, Cladosporium paeoniae. Both cultures have been 

used to generate DNA sequence data. 

The two stages (morphs) of this fungus are usually ecologically 

and seasonally separated, but sometimes conidiophores of the 

leaf-blotching (cladosporioid) morph also occur on dead stems of 

peony intermixed with the macronematous conidiophores of the 

periconioid morph. In culture conidiophore and conidial width tends 

to be narrower than on the natural substratum, and the conidia are 

not as frequently septate. 

DISCUSSION 

Cultural studies by ourselves and McKemy & Morgan-Jones 

(1991), and molecular sequence analyses documented herein 

clearly demonstrate that Cladosporium chlorocephalum, occurring 

on necrotic stems, and C. paeoniae, causing leaf-blotch symptoms 

on living leaves of Paeonia spp., are two synanamorphs of a single 

species, which has to be excluded from Cladosporium s. str. since 

the conidiogenous loci are quite distinct from the characteristically 

coronate scars in the latter genus and because ITS sequences 

indicate clear separation from Cladosporium s. str. 

Analysis of the morphology and conidiogenesis showed that 

the macronematous stage of this fungus (C. chlorocephalum, the 

periconioid morph) closely resembles Metulocladosporiella, recently 

introduced for the Cladosporium speckle disease of banana. There 

are, however, some differences. In Metulocladosporiella musae 

(E.W. Mason) Crous et al., the type species, micronematous 

conidiophores occur in vitro and in vivo, and macronematous 

conidiophores occur on leaf-spots, whereas in C. chlorocephalum 

the semi-macronematous conidiophores usually accompany 

leaf-blotch symptoms on living leaves and the macronematous 

conidiophores occur in saprobic growth on old necrotic stems. 

Rhizoid hyphae arising from the swollen basal cells of the 

macronematous conidiophores are characteristic for M. musae, but 

lacking in C. chlorocephalum, and the conidia in the latter species 

are 0–5-septate, but only 0(–1)-septate in M. musae. The semimacronematous, 

leaf-blotching stage (the cladosporioid morph) 

is barely distinguishable from the present concept of Fusicladium, 

which includes species with catenate conidia (Schubert et al. 2003). 

However, the peony fungus does not cluster within the Venturiaceae. 

Since C. chlorocephalum clusters apart of the Chaetothyriales, 

the clade to which Metulocladosporiella belongs, the differences 

observed here seem to be sufficient to place this fungus in a new 

genus (also see Crous et al. 2007 – this volume). Crous et al. 

(2006a) discussed differences between Metulocladosporiella and 

allied dematiaceous hyphomycete genera and provided a key to 

the latter genus and morphologically similar genera. Using this key, 

attempts to determine the macronematous morph of Cladosporium 


chlorocephalum lead to Metulocladosporiella. Differences between 

morphologically similar genera have been discussed in the paper 

by Crous et al. (2006a) and are also valid for the new genus 

Dichocladosporium. Parapericoniella U. Braun, Heuchert & K. 

Schub., a fungicolous genus recently introduced to accommodate 

Cladosporium asterinae Deighton, is also morphologically similar 

in having apically, densely branched conidiophores and truncate, 

unthickened conidiogenous loci and hila, but is quite distinct in not 

having micronematous conidiophores (Heuchert et al. 2005). 


We are much obliged to the curators of B, F, HBG, M, PH and SIENA for the 

loans of the collections studied. R. Kirschner and N. Ale-Agha are thanked for 

sending collections and cultures on Paeonia spp. We are very grateful to the 

Institute of Zoology of the Martin-Luther-University, above all to G. Tschuch, for 

providing access to SEM facilities. This work was supported in part by a grant of 

the “Graduiertenförderung des Landes Sachsen-Anhalt” and a grant of Synthesys 

(No. 2559) awarded to K.S. We thank M. Vermaas for preparing the photographic 

plates. 

REFERENCES 



nov., the teleomorph of Cladosporium s.str. Mycological Progress 2: 3–18. 



Crous PW, Braun U, Schubert K, Groenewald JZ (2007). Delimiting Cladosporium 






A, Burgess T, Barber P, Groenewald JZ (2006b). Phylogenetic lineages in the 




Dhingra OD, Sinclair JB (1985). Basic plant pathology methods. CRC Press, Boca 

Raton, Florida. 

Fresenius JBGW (1850). Beiträge zur Mykologie 1. Heinrich Ludwig Brömmer 

Verlag, Frankfurt. 

Gams W, Verkley GJM, Crous PW (2007). CBS Course of Mycology. 5 th ed. CBS, 

Utrecht. 





Kiffer E, Morelet M (1999). The Deuteromycetes. Mitosporic Fungi, Classification 

and Generic Key. Science Publishers, Enfield, NJ. 

Lee SB, Taylor JW (1990). Isolation of DNA from fungal mycelia and single spores. 

In: PCR Protocols: a guide to methods and applications (Innis MA, Gelfand DH, 

Sninsky JJ, White TJ, eds). Academic Press, San Diego, California: 282–287. 

Mason EW, Ellis MB (1953). British species of Periconia. Mycological Papers 56: 

1–127. 

McKemy JM, Morgan-Jones G (1991). Studies in the genus Cladosporium sensu lato 

III. Concerning Cladosporium chlorocephalum and its synonym Cladosporium 

paeoniae, the causal organism of leaf-blotch of peony. Mycotaxon 41: 135– 

146. 

Meuli LJ (1937). Cladosporium leaf blotch of peony. Phytopathology 27: 172–182. 




Passerini G (1876). Cladosporium paeoniae. Just’s Botanische Jahresberichte 4: 

235. 


Kew. 



98: 625–634. 

103


Saccardo PA (1882). Fungorum extra-europaeorum Pugillus. Michelia 2(6): 136– 

149 ‘1880’. 

Saccardo PA (1886). Sylloge Fungorum vol. 4. Padova. 

Schubert K (2005). Morphotaxonomic revision of foliicolous Cladosporium species 

(hyphomycetes). Ph.D. dissertation. Martin-Luther-University Halle-Wittenberg. 

http://sundoc.bibliothek.uni-halle.de/diss-online/05/05H208/index.htm 






Vries GA de (1952). Contribution to the knowledge of the genus Cladosporium Link 

ex Fr. CBS, Baarn. 





104


doi:10.3114/sim.2007.58.05 


Biodiversity in the Cladosporium herbarum complex (Davidiellaceae, Capnodiales), 

with standardisation of methods for Cladosporium taxonomy and diagnostics 

K. Schubert 1* , J. Z. Groenewald 2 , U. Braun 3 , J. Dijksterhuis 2 , M. Starink 2 , C.F. Hill 4 , P. Zalar 5 , G.S. de Hoog 2 and P.W. Crous 2 

1 

Botanische Staatssammlung München, Menzinger Strasse 67, D-80638 München, Germany; 2 CBS Fungal Biodiversity Centre, P.O. Box 85167, 3508 AD Utrecht, The Netherlands; 

3 

Martin-Luther-Universität, Institut für Biologie, Geobotanik und Botanischer Garten, Herbarium, Neuwerk 21, D-06099 Halle (Saale), Germany; 4 Plant & Environment Laboratory 

Biosecurity NZ, Ministry of Agriculture & Forestry, P.O. Box 2095, Auckland 1140, New Zealand; 5 Biotechnical Faculty, Department of Biology, Večna pot 111, SI-1000 Ljubljana, 

Slovenia 

*Correspondence: Konstanze Schubert, konstanze.schubert@gmx.de 

Abstract: The Cladosporium herbarum complex comprises five species for which Davidiella teleomorphs are known. Cladosporium herbarum s. str. (D. tassiana), C. 

macrocarpum (D. macrocarpa) and C. bruhnei (D. allicina) are distinguishable by having conidia of different width, and by teleomorph characters. Davidiella variabile is 

introduced as teleomorph of C. variabile, a homothallic species occurring on Spinacia, and D. macrospora is known to be the teleomorph of C. iridis on Iris spp. The C. herbarum 

complex combines low molecular distance with a high degree of clonal or inbreeding diversity. Entities differ from each other by multilocus sequence data and by phenetic 

differences, and thus can be interpreted to represent individual taxa. Isolates of the C. herbarum complex that were formerly associated with opportunistic human infections, 

cluster with C. bruhnei. Several species are newly described from hypersaline water, namely C. ramotenellum, C. tenellum, C. subinflatum, and C. herbaroides. Cladosporium 

pseudiridis collected from Iris sp. in New Zealand, is also a member of this species complex and shown to be distinct from C. iridis that occurs on this host elsewhere in the 

world. A further new species from New Zealand is C. sinuosum on Fuchsia excorticata. Cladosporium antarcticum is newly described from a lichen, Caloplaca regalis, collected 

in Antarctica, and C. subtilissimum from grape berries in the U.S.A., while the new combination C. ossifragi, the oldest valid name of the Cladosporium known from Narthecium 

in Europe, is proposed. Standard protocols and media are herewith proposed to facilitate future morphological examination of Cladosporium spp. in culture, and neotypes or 

epitypes are proposed for all species treated. 

Taxonomic novelties: Cladosporium antarcticum K. Schub., Crous & U. Braun, sp. nov., C. herbaroides K. Schub., Zalar, Crous & U. Braun, sp. nov., C. ossifragi (Rostr.) U. 

Braun & K. Schub., comb. nov., C. pseudiridis K. Schub., C.F. Hill, Crous & U. Braun, sp. nov., C. ramotenellum K. Schub., Zalar, Crous & U. Braun, sp. nov., C. sinuosum K. 

Schub., C.F. Hill, Crous & U. Braun, sp. nov., C. subinflatum K. Schub., Zalar, Crous & U. Braun, sp. nov., C. subtilissimum K. Schub., Dugan, Crous & U. Braun, sp. nov., C. 

tenellum K. Schub., Zalar, Crous & U. Braun sp. nov., Davidiella macrocarpa Crous, K. Schub. & U. Braun, sp. nov., D. variabile Crous, K. Schub. & U. Braun, sp. nov. 

Key words: Clonality, Davidiella, homothallism, new species, phylogeny, recombination, taxonomy. 

Introduction 

Cladosporium herbarum (Pers. : Fr.) Link, type species of the genus 

Cladosporium Link, is one of the most common environmental fungi 

to be isolated worldwide. It abundantly occurs on fading or dead 

leaves of herbaceous and woody plants, as secondary invader 

on necrotic leaf spots, and has frequently been isolated from 

air (Samson et al. 2000), soil (Domsch et al. 1980), foodstuffs, 

paints, textiles, humans (de Hoog et al. 2000) and numerous 

other substrates. It is also known to occur on old carpophores 

of mushrooms and other fungi (Heuchert et al. 2005) and to be 

a common endophyte (Riesen & Sieber 1985, Brown et al. 1998, 

El-Morsy 2000), especially in temperate regions. Under favourable 

climatic conditions C. herbarum also germinates and grows as an 

epiphyte on the surface of green, healthy leaves (Schubert 2005). 

Persoon (1794) introduced C. herbarum as Dematium 

herbarum Pers., which was later reclassified by Link (1809) as 

Acladium herbarum (Pers.) Link. In 1816, Link included C. herbarum 

together with three additional species in his newly described genus 

Cladosporium. Clements & Shear (1931) proposed C. herbarum 

as lectotype species of the latter genus, a decision followed by de 

Vries (1952) and Hughes (1958). Several authors provided detailed 

treatments of C. herbarum (de Vries 1952, Ellis 1971, Domsch et al. 

1980, Prasil & de Hoog 1988), and there are literally thousands of 

records of this species in the literature. McKemy & Morgan-Jones 

(1991) and Ho et al. (1999) examined C. herbarum in culture and 

published detailed descriptions of its features in vitro. 

Cladosporium macrocarpum Preuss, a second component 

within the herbarum complex, has hitherto been known and treated 

as an allied, but morphologically distinct species on the basis of its 

wider and somewhat larger, frequently 2–3-septate, more regularly 

verrucose conidia, shorter conidial chains and more pronounced 

prolongations of the conidiophores. Dugan & Roberts (1994) carried 

out examinations of morphological and reproductive aspects of both 

species, and in so doing demonstrated a morphological continuum 

between C. macrocarpum and C. herbarum, concluding that the 

name herbarum should have preference. Therefore, Ho et al. (1999) 

introduced the new combination C. herbarum var. macrocarpum 

(Preuss) M.H.M. Ho & Dugan. Although transitional forms have 

been discussed to occur between the two species, several authors 

still prefer to retain C. macrocarpum as a separate species. 

In an attempt to elucidate the species within the C. herbarum 

complex, therefore, a multilocus DNA sequence typing approach 

was used, employing five genes, namely the internal transcribed 

spacers of the rDNA genes (ITS), actin, calmodulin, translation 

elongation factor 1-α, and histone H3. These data were 

supplemented with morphological examinations under standardised 

conditions, using light and scanning electron microscopy, as well as 

cultural characteristics and growth studies. 

Material and methods 

Isolates 

Isolates included in this study were obtained from the culture 

collection of the Centraalbureau voor Schimmelcultures (CBS), 

Utrecht, Netherlands, or were freshly isolated from a range of 

different substrates. Single-conidial and ascospore isolates were 

obtained using the techniques as explained in Crous (1998) for 

species of Mycosphaerella Johanson and its anamorphs. Isolates 

105


were inoculated onto 2 % potato-dextrose agar (PDA), synthetic 

nutrient-poor agar (SNA), 2 % malt extract agar (MEA) and oatmeal 

agar (OA) (Gams et al. 2007), and incubated under continuous 

near-ultraviolet light at 25 °C to promote sporulation. All cultures 

obtained in this study are maintained in the culture collection of 

the CBS (Table 1). Nomenclatural novelties and descriptions were 

deposited in MycoBank (www.MycoBank.org). 

DNA isolation, amplification and sequence analysis 


DNA was isolated as described in Gams et al. (2007). Partial gene 

sequences were determined as described by Crous et al. (2006) 

for actin (ACT), calmodulin (CAL), translation elongation factor 1- 

alpha (EF), histone H3 (HIS) and part (ITS) of the nuclear rDNA 

operon spanning the 3’ end of the 18S rRNA gene (SSU), the 

first internal transcribed spacer, the 5.8S rRNA gene, the second 

internal transcribed spacer and the 5’ end of the 28S rRNA gene 

(LSU). The nucleotide sequences were generated using both PCR 

primers to ensure good quality sequences over the entire length of 

the amplicon. Subsequent sequence alignment and phylogenetic 

analysis followed the methods of Crous et al. (2006). Gaps longer 

than 10 bases were coded as single events for the phylogenetic 

analyses; the remaining gaps were treated as new character 

states. Sequence data were deposited in GenBank (Table 1) and 

the alignment and tree in TreeBASE (www.treebase.org). 

Data analysis 

The number of entities in the dataset of 79 strains was inferred 

with Structure v. 2.2 software (Pritchard et al. 2000, Falush et al. 

2003) using an UPGMA tree of data of the ACT gene compared 

with CAL, EF and HIS with the exclusion of the nearly invariant ITS 

region. For this analysis group indications were derived from a tree 

produced with MrAic (Nylander 2004). The length of the burn-in 

period was set to 1 000 000, number of MCMC repeats after burn-in 

10 000, with admixture ancestry and allele frequencies correlated 

models, assuming that all groups diverged from a recent ancestral 

population and that allele frequencies are due to drift. Uniform prior 

for ALPHA was set to 1.0 (default) and allele frequencies with λ set to 

1.0 (default). The numbers of MCMC repetitions after burn-in were 

set as 10 000 and 100 000. The number of clusters (K) in Structure 

was assumed from 5 to 7. Population differentiation F ST 

(index: θ) 

was calculated with 1–6 runs using the same software. The null 

hypothesis for this analysis is no population differentiation. When 

observed theta (θ) is significantly different from those of random 

data sets (p < 0.05), population differentiation is considered. 

Association of multilocus genotypes was screened with the 

multilocus option in BioNumerics v. 4.5. To test for reproductive 

mode in each population, the standardised index of association 

(I S ; Haubold et al. 1998) was calculated with start2 software 

A 

(Jolley et al. 2001). The null hypothesis for this analysis is complete 

panmixia. The values of I S were compared between observed and 

A 

randomised datasets. The hypothesis would be rejected when p < 

0.05. Mean genetic diversity (H) and diversities of individual loci 

were calculated with lian v. 3.5 (Haubold & Hudson 2000). Degrees 

of recombination or horizontal gene transfer were also visualised 

using SplitsTree v. 4.8 software (Huson & Bryant 2006). Split 

decomposition was carried out with default settings, i.e., character 

transformation using uncorrected (observed, “P”) distances, splits 

transformation using “equal angle”, and optimise boxes iteration set 

to 2. 

Morphology 

As the present study represents the first in a series dealing 

with Cladosporium spp. and their Davidiella Crous & U. Braun 

teleomorphs in culture, a specific, standardised protocol was 

established by which all species complexes will be treated in 

future. 

Morphology of the anamorph: Microscopic observations were made 

from colonies cultivated for 7 d under continuous near-ultraviolet 

light at 25 °C on SNA. Preparations were mounted in Shear’s 

solution (Gams et al. 2007). To study conidial development and 

branching patterns, squares of transparent adhesive tape (Titan 

Ultra Clear Tape, Conglom Inc., Toronto, Canada) were placed on 

conidiophores growing in the zone between the colony margin and 

2 cm inwards, and mounted between two drops of Shear’s solution 

under a glass coverslip. Different types of conidia are formed by 

Cladosporium species for which different terms need to be adopted. 

Ramoconidia are conidia with usually more than one (mostly 2 

or 3) conidial hilum, which typically accumulate at the tip of these 

conidia. Conidiogenous cells with more than one conidiogenous 

locus are first formed as apical parts of conidiophores. Such apical 

Fig. 1. Cladosporium conidiophore with ramoconidia, secondary ramoconidia, 

intercalary conidia, and small, terminal conidia. Scale bar = 10 µm. K. Schubert 

del. 

106

Cladosporium herbarum species complex 

Aculeate 

Spinulose 

Digitate 

Muricate 

Granulate 

Colliculate 

Pustulate 

Pedicellate 

Fig. 2. Terms used to describe conidium wall ornamentation under the cryo-electron 

microscope. Adapted from David (1997). 

than the small terminal conidia. In older literature true ramoconidia 

were often cited as “ramoconidia s. str.”, whereas secondary 

ramoconidia have been referred to as “ramoconidia s. lat.” 

Morphology of the teleomorph: Teleomorphs were induced by 

inoculating plates of 2 % tap water agar onto which autoclaved 

stem pieces of Urtica dioica (European stinging nettle) were placed. 

Inoculated plates were incubated on the laboratory bench for 7 d, 

after that period they were further incubated at 10 °C in the dark for 

1–2 mo to stimulate teleomorph development. Wherever possible, 

30 measurements (× 1 000 magnification) were made of conidia 

and ascospores, with the extremes of spore measurements given 

in parentheses. Cultural characteristics: Colonies were cultivated 

on PDA, MEA and OA plates for 14 d at 25 °C in the dark, after 

which the surface and reverse colours were rated using the charts 

of Rayner (1970). Linear growth was determined on MEA, PDA and 

OA plates by inoculating three plates per isolate for each medium, 

and incubating them for 14 d at 25 ºC, after that period colony 

diameters were determined. 

Low-temperature scanning electron microscopy 

Isolates of Cladosporium spp. were grown on SNA with 30 g agar/L 

for 3–4 d at room temperature under black light. Relevant parts of 

the small colonies with conidiophores and conidia were selected 

under a binocular, excised with a surgical blade as small agar (3 

× 3 mm) blocks, and transferred to a copper cup for snap-freezing 

in nitrogen slush. Agar blocks were glued to the copper surface 

with frozen tissue medium (KP-Cryoblock, Klinipath, Duiven, 

Netherlands) mixed with 1 part colloidal graphite (Agar Scientific, 

Stansted, U.K.). Samples were examined in a JEOL 5600LV 

scanning electron microscope (JEOL, Tokyo, Japan) equipped 

with an Oxford CT1500 Cryostation for cryo-electron microscopy 

(cryoSEM). Electron micrographs were acquired from uncoated 

frozen samples, or after sputter-coating by means of a gold/ 

palladium target for 3 times during 30 s (Fig. 2). Micrographs of 

uncoated samples were taken at an acceleration voltage of 3 kV, 

and consisted out of 30 averaged fast scans (SCAN 2 mode), and 

at 5 kV in case of the coated samples (PHOTO mode). 

parts of conidiophores are called ramoconidia if they secede at 

a septum from the conidiophore (Kirk et al. 2001). The septum at 

which the ramoconidium secedes often appears to be somewhat 

refractive or darkened. Ramoconidia are characterised by having 

a truncate, undifferentiated base (thus they lack a differentiated, 

coronate basal hilum formed in the context of conidiogenesis) 

and they can be very long, aseptate to sometimes multi-septate. 

Although they were formed initially as part of the conidiophore, 

they function as propagules. Only few of the species known 

until now have the ability to form true ramoconidia. Secondary 

ramoconidia also have more than one distal conidial hilum but 

they always derive from a conidiogenous locus of an earlier formed 

cell, which can be either a conidiogenous cell or a ramoconidium. 

Secondary ramoconidia are often shorter but somewhat wider than 

ramoconidia; they are often septate, and typically have a narrowed 

base with a coronate hilum (Fig. 1). Conidia in Cladosporium are 

cells with a coronate basal hilum, which is formed in the context of 

conidiogenesis and with either a single (when formed as intercalary 

units in unbranched parts of chains) or without any distal conidial 

hilum (when formed at the tip of conidial chains). For the first, the 

term “intercalary conidium” and for the latter, “small terminal 

conidium” is used. Intercalary conidia typically are larger and more 

pigmented and have a more differentiated surface ornamentation 

RESULTS 

Phylogeny and differentiation 

The manually adjusted alignment contained 80 sequences (including 

the outgroup sequence) and the five loci were represented by a 

total of 1 516 characters including alignment gaps which were 

used in the analysis. Of the 1 516 characters, 369 were parsimonyinformative, 

259 were variable and parsimony-uninformative, and 

888 were constant. 

Forty equally most parsimonious trees (TL = 1 933 steps; CI 

= 0.569; RI = 0.786; RC = 0.447), one of which is shown in Fig. 

3, were obtained from the parsimony analysis of the combined 

genes. Neighbour-joining analysis using three substitution models 

(uncorrected “p”, Kimura 2-parameter and HKY85) on the sequence 

data yielded trees with identical topologies. These differed from 

the tree presented in Fig. 3 with regard to the placement of C. 

macrocarpum strain CPC 12054 which was placed as a sister 

branch to the C. bruhnei Linder clade in the distance analyses 

(results not shown) because it shares an identical CAL sequence. 

All cryptic species consisting of multiple strains are clustering in 

well-supported clades with bootstrap support values ranging from 

71 % (C. herbarum) to 100 % [e.g. C. ramotenellum K. Schub., 


107


Table 1. Isolates subjected to DNA sequence analyses and morphological examinations. 


(ITS, EF, ACT, CAL, HIS) 

Cladosporium antarcticum — CBS 690.92* (ex-type) Caloplaca regalis Antarctica C. Möller EF679334, EF679405, EF679484, EF679560, EF679636 

Cladosporium bruhnei Davidiella allicina CBS 134.31 = ATCC 11283 — Germany — EF679335, EF679406, EF679485, EF679561, EF679637 

CBS 157.82 Quercus robur Belgium — EF679336, EF679407, EF679486, EF679562, EF679638 

CBS 159.54 = ATCC 36948 Man, skin The Netherlands — EF679337, EF679408, EF679487, EF679563, EF679639 

CBS 161.55 Man, sputum The Netherlands — EF679338, EF679409, EF679488, EF679564, EF679640 

CBS 177.71 Thuja tincture The Netherlands — EF679339, EF679410, EF679489, EF679565, EF679641 

CBS 188.54 = ATCC 11290 = IMI 049638 = CPC — — — AY251077, EF679411, EF679490, EF679566, EF679642 

3686 

CBS 366.80 Man, skin The Netherlands — EF679340, EF679412, EF679491, EF679567, EF679643 

CBS 399.80 Man, skin The Netherlands — AJ244227, EF679413, EF679492, EF679568, EF679644 

CBS 521.68 Air The Netherlands — EF679341, EF679414, EF679493, EF679569, EF679645 

CBS 572.78 Polyporus radiatus Russia V.K. Melnik DQ289799, EF679415, DQ289866, DQ289831, EF679646 

CBS 813.71 Polygonatum odoratum Czech Republic — EF679342, EF679416, EF679494, EF679570, EF679647 

CBS 110024 Industrial water Germany, Nordrhein-Westfalen — EF679343, EF679417, EF679495, EF679571, EF679648 

CBS 115683 = ATCC 66670 = CPC 5101 CCA-treated Douglas-fire pole U.S.A., New York C.J. Wang AY361959, EF679418, AY752193, AY752224, AY752255 

CBS 121624* = CPC 12211 (neotype) Hordeum vulgare Belgium J.Z. Groenewald EF679350, EF679425, EF679502, EF679578, EF679655 

CPC 11386 Tilia cordata Germany, Sachsen-Anhalt K. Schubert EF679344, EF679419, EF679496, EF679572, EF679649 

CPC 11840 Ourisia macrophylla New Zealand A. Blouin EF679345, EF679420, EF679497, EF679573, EF679650 

CPC 12042 = EXF-389 Hypersaline water from salterns Slovenia P. Zalar EF679346, EF679421, EF679498, EF679574, EF679651 

CPC 12045 = EXF-594 Hypersaline water from salterns Spain P. Zalar EF679347, EF679422, EF679499, EF679575, EF679652 

CPC 12046 = EXF-680 Air conditioning system Slovenia P. Zalar EF679348, EF679423, EF679500, EF679576, EF679653 

CPC 12139 Hordeum vulgare The Netherlands — EF679349, EF679424, EF679501, EF679577, EF679654 

CPC 12212 Hordeum vulgare Belgium J.Z. Groenewald EF679351, EF679426, EF679503, EF679579, EF679656 

CPC 12921 Eucalyptus sp. Australia — EF679352, EF679427, EF679504, EF679580, EF679657 

Cladosporium cladosporioides — CBS 673.69 Air The Netherlands — EF679353, EF679428, EF679505, EF679581, EF679658 

complex 

Davidiella sp. CBS 109082 Silene maritima United Kingdom A. Aptroot EF679354, EF679429, EF679506, EF679582, EF679659 

— CPC 11606 Musa sp. India M. Arzanlou EF679355, EF679430, EF679507, EF679583, EF679660 

— CPC 11609 Musa sp. India M. Arzanlou EF679356, EF679431, EF679508, EF679584, EF679661 

Cladosporium herbaroides — CBS 121626* = CPC 12052 = EXF-1733 (ex-type) Hypersaline water from salterns Israel P. Zalar EF679357, EF679432, EF679509, EF679585, EF679662 

Cladosporium herbarum Davidiella tassiana CBS 111.82 Arctostaphylos uva-ursi Switzerland E. Müller AJ238469, EF679433, EF679510, EF679586, EF679663 

CBS 300.49 Biscutella laevigata Switzerland J.A. von Arx EF679358, EF679434, EF679511, EF679587, EF679664 

CBS 121621* = CPC 12177 (epitype) Hordeum vulgare The Netherlands — EF679363, EF679440, EF679516, EF679592, EF679670 

CPC 11600 Delphinium barbeyi U.S.A., Colorado A. Ramalay DQ289800, EF679435, DQ289867, DQ289832, EF679665 

CPC 11601 Delphinium barbeyi U.S.A., Colorado A. Ramalay EF679359, EF679436, EF679512, EF679588, EF679666 



108


Cladosporium iridis Davidiella 

macrospora 

Cladosporium macrocarpum Davidiella 

macrocarpa 







CBS 107.20 Iris sp. — — EF679369, EF679446, EF679522, EF679598, EF679676 

CBS 138.40* (epitype) Iris sp. The Netherlands — EF679370, EF679447, EF679523, EF679599, EF679677 

CBS 175.82 Water Romania — EF679371, EF679448, EF679524, EF679600, EF679678 

CBS 223.31 = ATCC 11287 Mycosphaerella tulasnei — — AF222830, EF679449, EF679525, EF679601, EF679679 

CBS 299.67 Triticum aestivum Turkey — EF679372, EF679450, EF679526, EF679602, EF679680 

CBS 121811* = CPC 12755 (neotype) Spinacia oleracea U.S.A. — EF679376, EF679454, EF679530, EF679606, EF679684 

CPC 11817 Corylus sp. U.S.A. — EF679373, EF679451, EF679527, EF679603, EF679681 


CBS H-19855 = CPC 12752 = CBS 121623 Spinacia oleracea U.S.A. — EF679375, EF679453, EF679529, EF679605, EF679683 

CPC 12756 Spinacia oleracea U.S.A. — EF679377, EF679455, EF679531, EF679607, EF679685 




Cladosporium ossifragi — CBS 842.91* (epitype) Narthecium ossifragum Norway M. di Menna EF679381, EF679459, EF679535, EF679611, EF679689 

CBS 843.91 Narthecium ossifragum Norway M. di Menna EF679382, EF679460, EF679536, EF679612, EF679690 

Cladosporium pseudiridis — CBS 116463* = LYN 1065 = ICMP 15579 (ex-type) Iris sp. New Zealand C.F. Hill EF679383, EF679461, EF679537, EF679613, EF679691 

Cladosporium ramotenellum — CBS 121628* = CPC 12043 = EXF-454 (ex-type) Hypersaline water from salterns Slovenia P. Zalar EF679384, EF679462, EF679538, EF679614, EF679692 

CPC 12047 = EXF-967 Air conditioning system Slovenia P. Zalar EF679385, EF679463, EF679539, EF679615, EF679693 

Fuchsia excorticata New Zealand A. Blouin EF679386, EF679464, EF679540, EF679616, EF679694 

Cladosporium sinuosum — CBS 121629* = CPC 11839 = ICMP 15819 (extype) 

Cladosporium spinulosum — CBS 102044 Hypersaline water from salterns Slovenia S. Soujak EF679387, EF679465, EF679541, EF679617, EF679695 

CBS 119907* = CPC 12040 = EXF-334 (ex-type) Hypersaline water from salterns Slovenia P. Zalar EF679388, EF679466, EF679542, EF679618, EF679696 

Cladosporium subinflatum — CBS 121630* = CPC 12041 = EXF-343 (ex-type) Hypersaline water from salterns Slovenia P. Zalar EF679389, EF679467, EF679543, EF679619, EF679697 

Cladosporium sp. — CBS 172.52 = ATCC 11320 Carya illinoensis U.S.A. — EF679390, EF679468, EF679544, EF679620, EF679698 

CBS 113741 Grape berry U.S.A. — EF679391, EF679469, EF679545, EF679621, EF679699 

CBS 113742 Grape berry U.S.A. — EF679392, EF679470, EF679546, EF679622, EF679700 

CBS 113744 Grape bud U.S.A. — EF679393, EF679471, EF679547, EF679623, EF679701 

CPC 12484 Pinus ponderosa Argentina A. Greslebin EF679394, EF679472, EF679548, EF679624, EF679702 

CPC 12485 Pinus ponderosa Argentina A. Greslebin EF679395, EF679473, EF679549, EF679625, EF679703 

Cladosporium subtilissimum — CBS 113753 Bing cherry fruits U.S.A. — EF679396, EF679474, EF679550, EF679626, EF679704 

CBS 113754* Grape berry U.S.A. — EF679397, EF679475, EF679551, EF679627, EF679705 


Cladosporium tenellum — CBS 121634* = CPC 12053 = EXF-1735 (ex-type) Hypersaline water from salterns Israel P. Zalar EF679401, EF679479, EF679555, EF679631, EF679709 


109




(ITS, EF, ACT, CAL, HIS) 

CPC 11813 Phyllactinia sp. on Corylus sp. U.S.A. D. Glawe EF679399, EF679477, EF679553, EF679629, EF679707 

CPC 12051 = EXF-1083 Hypersaline water from salterns Israel P. Zalar EF679400, EF679478, EF679554, EF679630, EF679708 

Cladosporium variabile Davidiella variabile CBS 121636* = CPC 12751 (epitype) Spinacia oleracea U.S.A. — EF679402, EF679480, EF679556, EF679632, EF679710 


— Davidiella sp. CBS 289.49 Allium schoenoprasum Switzerland E. Müller AY152552, EF679482, EF679558, EF679634, EF679712 

CBS 290.49 Trisetum distichophyllum Switzerland E. Müller EF679404, EF679483, EF679559, EF679635, EF679713 

1 ATCC: American Type Culture Collection, Virginia, U.S.A.; CBS: Centraalbureau voor Schimmelcultures, Utrecht, The Netherlands; CPC: Culture collection of Pedro Crous, housed at CBS; EXF: Extremophilic Fungi Culture Collection of the Department 

of Biology, Biotechnical Faculty, University of Ljubljana, Slovenia; ICMP: International Collection of Micro-organisms from Plants, Landcare Research, Private Bag 92170, Auckland, New Zealand; IMI: International Mycological Institute, CABI-Bioscience, 

Egham, Bakeham Lane, U.K. 

2 ACT: partial actin gene, CAL: partial calmodulin gene, EF: partial elongation factor 1-alpha gene, HIS: partial histone H3 gene, ITS: internal transcribed spacer region. 


Zalar, Crous & U. Braun and C. ossifragi (Rostr.) U. Braun & K. 

Schub.]. The intraspecific variation in the C. bruhnei clade is due 

to genetic variation present in the sequence data of all loci except 

for ITS, those in the C. macrocarpum clade in all loci except for ITS 

and ACT, and those in the C. herbarum clade in all loci except for 

ITS and CAL (data not shown). However, none of the variation for 

these species could be linked to host specificity or morphological 

differences. In general, ITS data did not provide any resolution 

within the C. herbarum complex, whereas EF data provided species 

clades with very little intraspecific variation and ACT, CAL and HIS 

revealed increasing intraspecific variation (ACT the least and HIS 

the most). 

The mean genetic diversity (H) of the entire data set excluding 

the nearly invariant ITS region was 0.9307, with little difference 

between genes (ACT = 0.9257, CAL = 0.9289, EF = 0.9322, HIS 

= 0.9361). The loci showed different numbers of alleles (ACT: 22, 

CAL: 16, EF: 21, HIS: 20, ITS: 6). Differentiation of entities when 

calculated with Structure software using the admixture/correlated 

model showed highest value with K = 6. At this value F ST 

varied 

between 0.1362 and 0.3381. Linkage disequilibrium calculated 

using the standardised index of association (I S ) for the entire 

A 

dataset (observed variance V o 

= 0.5602, expected variance V e 

= 

0.2576) was 0.3914 (P = 0.0001), consistent with a small amount 

of recombination that did not destroy the linkage between alleles. 

Only few groups appeared to be separated for all alleles; degrees 

of gene flow are indicated in Fig. 4. SplitsTree software produced 

unresolved star-shaped structures for all genes, without any sign of 

reticulation (Fig. 5). 

110


100 

74 

100 

10 changes 

Cercospora beticola CPC11557 

CPC 11609 



Cladosporium cladosporioides complex 

98 

CPC 11606 

Davidiella sp. CBS 290.49 

100 CPC 12043 

CPC 12047 Cladosporium ramotenellum 

100 CBS 842.91 

80 

Cladosporium ossifragi 


Davidiella sp. CBS 289.49 

CPC 12053 

91 

100 

CPC 11813 Cladosporium tenellum 

64 

CPC 12051 

Cladosporium sp. CBS 113744 

63 

54 

94 

100 

98 

72 62 

81 

66 

67 

54 

67 

91 

51 



CPC 12044 

CPC 12139 






63 






CPC 12921 

CPC 12046 


CPC 12045 

CPC 11840 


CPC 12212 

CPC 12211 

CPC 12042 

CPC 11386 


Cladosporium sp. CPC 12485 

74 

83 

58 

85 


CPC 12040 

Cladosporium sp. CPC 12484 

Cladosporium sp. CBS 172.52 


Cladosporium antarcticum CBS 690.92 


Cladosporium subinflatum CPC 12041 

Cladosporium pseudiridis CBS 116463 

Cladosporium sinuosum CPC 11839 

93 

100 

51 

66 

81 

94 

65 

CBS 107.20 Cladosporium iridis 


Cladosporium herbaroides CPC 12052 

100 

64 

95 

71 

61 

99 

CPC 12751 

CPC 12753 

CPC 11817 

CPC 12758 

CPC 12759 

CPC 12054 



51 CPC 12757 


CPC 12756 

67 CPC 12755 

CPC 12752 

CPC 11603 


CPC 11600 

CPC 11604 

CPC 11602 

CPC 11601 


CPC 12177 

CPC 12183 

100 

CPC 12181 

CPC 12180 

CPC 12179 

CPC 12178 

Cladosporium subtilissimum 

Cladosporium bruhnei 

Cladosporium spinulosum 

Cladosporium variabile 

Cladosporium macrocarpum 

Cladosporium herbarum 

Fig. 3. One of 40 equally most parsimonious trees obtained from a heuristic search with 100 random taxon additions of the combined sequence alignment (ITS, ACT, CAL, EF, 

HIS). The scale bar shows ten changes, and bootstrap support values from 1 000 replicates are shown at the nodes. Thickened lines indicate the strict consensus branches 

and strain numbers in bold represent ex-type sequences. The tree was rooted to sequences of Cercospora beticola strain CPC 11557 (GenBank accession numbers AY840527, 

AY840458, AY840425, AY840494, AY840392, respectively). 


111


Davidiella sp. 

Global (Gapcost:0%) 

ACT 

80 

85 

D. macrosporum D. tassiana 

D. macrocarpa 

D. macrocarpa 

90 

D. macrospora 


D. variabile 

D. Variabile 

D. tassiana 

D. allicina 

D. alliacea 

95 

100 

• . CBS 673.69 

• . CPC 11606 

• . CBS 109082 

• . CPC 11609 

 

. CBS 110024 

 

. CBS 157.82 

 


 


 

. CPC 11386 

 

. CPC 12042 

 

. CPC 12046 

 

. CPC 12211 

 

. CPC 12212 

 

. CPC 11840 

 


 


 

. CPC 12921 

 


 


 


 


 

. CPC 12139 

 


 

. CPC12041 

 

. CPC 11839 

 


 

 


 

. CPC 12045 

 


 


 

. CPC 12044 

 

. CPC 12485 

 


 


 


 


 


 


 

. CPC 12040 

 


 


 


 


 

. CPC 12484 

 

. CPC 12751 

 

. CPC 12753 

• 


• 


 

. CPC 11600 

 

. CPC 11601 

 

. CPC 11602 

 

. CPC 11604 

 


 


 

. CPC 11603 

• 


• 


• 


• 

. CPC 11817 

• 

. CPC 12054 

 

. CPC 12752 

 

. CPC 12755 

 

. CPC 12756 

• 

. CPC 12758 

• 

. CPC 12759 

• 

. CPC 12757 

 

. CPC 12177 

 

. CPC 12178 

 

. CPC 12179 

 

. CPC 12180 

 

. CPC 12181 

 

. CPC 12183 

• 

. CPC 12052 

 

. CPC 11813 

 

. CPC 12051 

 

. CPC 12053 

 

. CPC 12043 

 

. CPC 12047 

 

























Cladosporium subinflatum 

Cladosporium sinuosum 




Cladosporium subtilissimum s.str. 



Cladosporium subtilissimum s.lat. 

Cladosporium antarcticum 




Cladosporium pseudiridis 










Cladosporium iridis 

Cladosporium iridis 

























Cladosporium herbaroides 

Cladosporium tenellum 



Cladosporium ramotenellum 



Water 

Germany 

Wood 

Belgium 

Human 

Netherlands 

Fungus 

Russia 

Tree 

Germany 

Hypersaline Slovenia 

Airco 

Slovenia 

Grass 

Belgium 

Grass 

Belgium 

Plant 

Australia 

? ? 

Human 


Tree 

Australia 

Human 


Chemical Netherlands 

Human 


Air 


Grass 


Treated pole USA 


Plant 

New Zealand 

? Germany 

Plant 

Czechia 

Hypersaline Spain 

Cherry 

USA . 

Grape 

USA 


. 

Wood 

Argentina 

Lichen 

Antarctica 

Grape 

USA 

Grape 

USA 

Plant 

USA 

Plant 

New Zealand 



Plant 

Switzerland 

Plant 

Norway 

Plant 

Norway 

Grape 

USA 

Wood 

Argentina 

Spinach 

USA 

Spinach 

USA 

Plant 


Plant 


Plant 

USA 

Plant 

USA 

Plant 

USA 

Plant 

USA 

Plant 


Plant 


Plant 

USA 

Water 

Romania 

? ? 

Straw 

Turkey 

Branch 

USA 


Spinach 

USA 

Spinach 

USA 

Spinach 

USA 

Spinach 

USA 

Spinach 

USA 

Spinach 

USA 

Grass 


Grass 


Grass 


Grass 


Grass 


Grass 


Saltern 

Israel 

Fungus 

USA 




Airco 

Slovenia 

Plant 


Fig. 4. Distance tree of the Cladosporium herbarum complex based on ACT sequence data generated with UPGMA, showing Structure analysis at K = 6 under admixture model 

with correlated allele frequencies. Group indications (18) are taken from a tree based on EF sequences with AIC under the HKYG model. 

112


B 

A 

C 

D 

Fig. 5. Split decomposition of the Cladosporium herbarum complex using SplitsTree of 16–22 unique alleles obtained from 79 Cladosporium isolates for four loci. The star-like 

structures suggest clonal development. A = ACT, B = CAL, C = HIS, D = EF. Scale bars = 0.01 nucleotide substitutions per site. 

Taxonomy 

Key to the Cladosporium species treated 

Morphological features used in the key to distinguish the species treated in this study were determined after 7 d growth at 25 ºC on SNA using 

light microscopy, and cultural characteristics after 14 d incubation on PDA. 

1. Conidia usually smooth, rarely minutely verruculose ..................................................................... C. cladosporioides (species complex) 

1. Conidia with different surface ornamentation, minutely to distinctly verruculose, verrucose to echinulate or spiny 

.................................................................................................................................................................................................................... 2 

2. Conidiophores uniform, macronematous; conidia solitary, sometimes formed in short unbranched chains .............................................. 3 

2. Conidiophores both macronematous and micronematous; conidia always catenate, usually formed in branched chains 

.................................................................................................................................................................................................................... 5 


113


3. Conidiophores due to geniculations often growing zigzag-like, (4–)5–7 µm wide; conidia 9–21 × (5–)6–8 µm, 0–1-septate; 

conidiogenous loci and conidial hila 1.2–2(–2.2) µm diam .................................................................................................... C. sinuosum 

3. Conidiophores not growing zigzag-like, wider, 6–11 µm; conidia very large and wide, 15–75(–87) × (7–)10–19(–21) µm, often with more 

septa; conidiogenous loci and hila wider, (2–)2.5–4 µm diam ................................................................................................................... 4 

4. Conidia (18–)30–75(–87) × (7–)10–16(–18) µm, (0–)2–6(–7)-septate, walls thickened, especially in older conidia, up to 1 µm thick 

......................................................................................................................................................................................................... C. iridis 

4. Conidia shorter and wider, 15–55 × (9–)11–19(–21) µm, 0–3-septate, walls distinctly thickened, up to 2 µm, usually appearing zonate 

.............................................................................................................................................................................................. C. pseudiridis 

5(2) Macronematous conidiophores nodulose or nodose with conidiogenous loci usually confined to swellings ........................................... 6 

5. Macronematous conidiophores non-nodulose or only occasionally subnodulose due to geniculate proliferation, but conidiogenous loci not 

confined to swellings ................................................................................................................................................................................ 11 

6. Macronematous conidiophores 3–6 µm wide, swellings 5–11 µm wide .................................................................................................... 7 

6. Macronematous conidiophores somewhat narrower, (1.5–)2.5–5 µm wide, swellings 3–8 µm wide ........................................................ 8 

7. Aerial mycelium twisted; conidial septa often distinctly darkened, becoming sinuous with age, apex and base of the conidia often appear 

to be distinctly darkened; slower growing in culture (29 mm after 14 d on PDA) ...................................................................... C. variabile 

7. Aerial mycelium not twisted; conidial septa as well as apex and base not distinctly darkened, septa not sinuous with age; faster growing 

in culture (on average 38 mm after 14 d on PDA) .......................................................................................................... C. macrocarpum 

8. Macronematous conidiophores (1.5–)2.5–4.5(–5.5) µm wide, swellings 3–6.5 µm wide; conidia 4–17(–22) µm long, ornamentation 

variable, but usually densely echinulate, spines up to 0.8 µm long .................................................................................... C. subinflatum 

8. Macronematous conidiophores slightly wider, 3–5 µm, swellings (4–)5–8(–9) µm wide; conidia longer, up to 25(–35) µm, ornamentation 

minutely verruculose to verrucose, but not echinulate or spiny ................................................................................................................. 9 

9. Conidia formed by macronematous conidiophores 3–33 × (2–)3–6(–7) µm, with age becoming wider, (3.5–)5–9(–11) µm, darker and 

more thick-walled ................................................................................................................................................................ C. herbaroides 

9. Conidia formed by macronematous conidiophores not becoming wider and darker with age, usually up to 7 µm wide 

.................................................................................................................................................................................................................. 10 

10. Conidiophores usually with small head-like swellings, sometimes also with a second intercalary nodule; small terminal conidia 

4–9 × 2.5–3.5 µm, secondary ramoconidia and occasionally formed ramoconidia 10–24(–31) × 3–5(–7) µm ........................ C. bruhnei 

10. Conidiophores with a single or often numerous swellings in short succession giving the stalk a knotty/gnarled appearance; 

conidia wider, small terminal conidia 4–10 × 3–5(–6) µm, intercalary conidia 6–16 × 4–6 µm, secondary ramoconidia 12–25(–35) × 

(3–)5–7(–9) µm ....................................................................................................................................................................... C. herbarum 

11(5) Small terminal and intercalary conidia 4–15 × 3–5 µm, secondary ramoconidia 16–36(–40) × (4–)5–8 µm, 0–3(–4)-septate, 

ramoconidia absent ................................................................................................................................................................. C. ossifragi 

11. Small terminal conidia, ramoconidia and secondary ramoconidia distinctly narrower, 2–5(–6) µm wide, 0–2(–3)-septate 

.................................................................................................................................................................................................................. 12 

12. Mycelium dimorphic, narrow hyphae 1–3 µm wide, hyaline to subhyaline, thin-walled, hyphae of the second type wider, 3.5–8(–9) 

µm, pale to dark greyish olivaceous or olivaceous-brown, thick-walled, sometimes even two-layered, 1(–1.5) µm thick, hyphae 

appearing consistently enveloped in polysaccharide-like material or covered by a slime coat; conidiophores usually several times slightly 

to distinctly geniculate towards the apex, with numerous conidiogenous loci crowded towards the apex, up to 14 per conidiogenous cell 

............................................................................................................................................................................................. C. antarcticum 

12. Mycelium not dimorphic, neither enveloped in polysaccharide-like material nor covered by a slime coat; conidiophores usually not 

geniculate, occasionally only slightly so ................................................................................................................................................... 13 

13. Conidial ornamentation distinctly echinulate, spiny (baculate, digitate or capitate under SEM), spines 0.5–1.3 µm long, loose to moderately 

dense, conidial hila usually situated on small peg-like prolongations or denticles .............................................................. C. spinulosum 

13. Conidial ornamentation different, minutely verruculose to verruculose, conidial hila not situated on peg-like prolongations 

.................................................................................................................................................................................................................. 14 

14. Small terminal conidia narrowly obovoid, limoniform or fusiform, but neither globose nor subglobose; conidiogenous loci and conidial hila 

0.5–2(–2.5) µm diam ........................................................................................................................ C. subtilissimum (species complex) 

14. Numerous small globose or subglobose terminal conidia formed, also ovoid or limoniform; conidiogenous loci and conidial hila somewhat 

smaller, 0.5–1.5(–2) µm diam .................................................................................................................................................................. 15 

114


15. Conidiophores usually with numerous conidiogenous loci forming sympodial clusters of pronounced scars at the apex, sometimes up to 

10 or even more denticulate loci; conidia 3–20(–28) × 2.5–5(–6) µm, 0–1(–2)-septate, often with several apically crowded hila, up to 7(–9) 

.................................................................................................................................................................................................. C. tenellum 

15. Conidiophores usually only with few conidiogenous loci, mostly 1–3; conidia longer and narrower, 2.5–35 × 2–4(–5) µm, 0–3-septate, 

usually with up to three distal conidial hila ....................................................................................................................... C. ramotenellum 

Key to the Davidiella species treated 

1. Ascospores frequently wider than 7 µm when mounted in Shear’s solution or lactic acid, apical cell obtusely rounded 

.................................................................................................................................................................................................................... 2 

1. Ascospores not wider than 7 µm when mounted in Shear’s solution or lactic acid, apical cell acutely rounded, ascospores (20–)25–27 

(–30) × (5.5–)6–7 µm ................................................................................................................................................................. D. allicina 

2. Pseudoparaphyses prominent; asci frequently >95 µm; ascospores (22–)23–26(–28) × (6–)6.5–7(–8) µm...................... D. macrocarpa 

2. Pseudoparaphyses mostly absent in older ascomata; asci


Fig. 6. Cladosporium antarcticum (CBS 690.92). Macro- and micronematous conidiophores and conidia. Scale bar = 10 µm. K. Schubert del. 

geniculate with several conidiogenous loci at the apex, 2–22 × 

2–3 µm, pale greyish olivaceous, loci denticulate. Ramoconidia 

occasionally occurring, cylindrical, up to 30 µm long, 4–5 µm 

wide, 0–1-septate, concolorous with the tips of conidiophores, with 

a broadly truncate, unthickened and not darkened base, without 

dome and rim, 2.5 µm wide. Conidia catenate, in branched chains, 

straight, small terminal conidia obovoid, limoniform or narrowly 

ellipsoid, 4–14 × 2.5–4 µm [av. ± SD, 8.5 (± 3.3) × 3.5 (± 0.6)], 

0(–1)-septate, secondary ramoconidia ellipsoid to cylindrical, often 

with several or numerous conidial hila crowded at the distal end, up 

to 12, 13–30 × 4–5 µm [av. ± SD, 20.1 (± 5.8) × 4.3 (± 0.5) µm], 

0–3-septate, sometimes slightly constricted at the median septum, 

pale olivaceous-brown or greyish brown, minutely verruculose to 

verrucose (granulate under SEM), walls more or less thickened, 

rounded or slightly attenuated towards apex and base, hila 

protuberant, denticulate, 0.8–1.5(–2) µm diam, thickened and 

darkened-refractive; microcyclic conidiogenesis occurring. 

Cultural characteristics: Colonies on PDA attaining 9 mm diam after 

14 d at 25 ºC, greenish olivaceous to grey-olivaceous, at the margin 

becoming dull green, reverse with a pale olivaceous-grey centre 

and a broad olivaceous-black margin, margin narrow, regular, 

entire edge, white, feathery, aerial mycelium sparse but colonies 

appearing felty, growth flat with somewhat elevated colony centre, 

prominent exudates not formed, sporulation dense, covering almost 

the whole colony. Colonies on MEA attaining 12 mm diam after 14 

d at 25 ºC, olivaceous-grey to iron-grey, iron-grey reverse, velvety 

to powdery, aerial mycelium sparse, sporulation profuse. Colonies 

on OA attaining 4 mm after 14 d at 25 ºC, olivaceous-grey, aerial 

mycelium sparse, diffuse, growth flat, without prominent exudates, 

sporulating. 

Specimen examined: Antarctica, King George, Arctowski, isolated from the lichen 

Caloplaca regalis (Teloschistaceae), C. Möller, No. 32/12, 1991, CBS-H 19857, 

holotype, isotype HAL 2024 F, culture ex-type CBS 690.92. 

Substrate and distribution: On the lichen Caloplaca regalis; 

Antarctica. 

Notes: This is the second genuine lichenicolous species of the 

genus Cladosporium. Cladosporium licheniphilum Heuchert & U. 

Braun, occurring on apothecia of Pertusaria alpina in Russia, is 

quite distinct from C. antarcticum by having subcylindrical or only 

slightly geniculate-sinuous, wider conidiophores, 5–8 µm, with 

numerous characteristic terminal branches and much shorter, 0– 

1-septate, smooth conidia, 3.5–13 × 3–7 µm (Heuchert & Braun 

2006). Cladosporium lichenicola Linds. was invalidly published and 

C. arthoniae M.S. Christ. & D. Hawksw. as well as C. lichenum 

Keissl. are to be excluded from the genus Cladosporium since 

they do not possess the typical cladosporioid scar structure but 

inconspicuous, unthickened conidiogenous loci and conidial hila 

(Hawksworth 1979, Heuchert et al. 2005). The fungicolous species 

C. uredinicola Speg. and the foliicolous species C. alneum Pass. 

ex K. Schub. and C. psoraleae M.B. Ellis are morphologically 

superficially similar. However, C. uredinicola, a widespread fungus 

on rust fungi, downy mildews and powdery mildew fungi, differs 

in having somewhat longer and wider, smooth conidia, 3–39 × 2– 

6.5(–8) µm, and wider conidiogenous loci and conidial hila, 0.5–3 

116


Fig. 7. Cladosporium antarcticum (CBS 690.92). A. 

Overview of the growth pattern on SNA. Note the very 

large bulbous cells formed at the base of different 

conidiophores. Other conidiophores sprout from the 

agar surface. B. Overview of conidiophores and conidia. 

Note the large distance of the scars on the conidiophore 

and the different stages of conidial formation on the 

tips of other conidia. The long secondary ramoconidia 

are also visible, and sparse aerial hyphae. C. Detail 

of B with details of the ornamentation and scars. The 

absence of ornamentation at the apical (spore-forming) 

end of the secondary ramoconidium is clearly visible. D– 

E. Tubular structures on coniophore (D) and secondary 

ramoconidium (E). Scale bars: A–B = 10 μm, C–D = 5 

µm, E = 2 µm. 

Fig. 8. Cladosporium antarcticum (CBS 690.92). A–B. Macronematous conidiophores. C, G. Mycelium enveloped by a polysaccharide-like layer. D, F. Conidia. E. Micronematous 

conidiophore. H. Ramoconidium with numerous distal scars. Scale bars = 10 µm. 

µm (Heuchert et al. 2005); C. alneum, which causes leaf spots on 

Alnus glutinosa, possesses longer and wider conidiophores, 25– 

260 × (2–)3–7(–8.5) µm, and somewhat shorter, smooth conidia 

(Schubert 2005, Schubert et al. 2006); and C. psoraleae, known 

from Myanmar on Psoralea corylifolia, can easily be distinguished 

from C. antarcticum by its smooth and wider conidia, 3.5–7 µm, 

and wider conidiogenous loci and conidial hila, 1–3 µm diam (Ellis 

1972, Schubert 2005). 


117


Fig. 9. Cladosporium bruhnei (CPC 12211). Macro- and micronematous conidiophores and conidia. Scale bar = 10 µm. K. Schubert del. 

Cladosporium bruhnei Linder, Bull. Natl. Mus. Canada 97: 259. 

1947. Figs 9–12. 

≡ Hormodendrum hordei Bruhne, in W. Zopf, Beitr. Physiol. Morph. nied. 

Org. 4: 1. 1894, non C. hordei Pass., 1887. 

≡ Cladosporium herbarum (Pers.: Fr.) Link var. (δ) cerealium Sacc. f. 

hordei (Bruhne) Ferraris, Flora Ital. Crypt., Pars I, Fungi, Fasc.13: 882. 

1914. 

≡ Cladosporium hordei (Bruhne) Pidopl., Gribnaja Flora Grubych Kormov: 

268. 1953, nom. illeg., homonym, non C. hordei Pass., 1887. 

Teleomorph: Davidiella allicina (Fr. : Fr.) Crous & Aptroot, in 

Aptroot, Mycosphaerella and its anamorphs: 2. Conspectus of 

Mycosphaerella. CBS Biodiversity Ser. 5: 30. 2006. 

Basionym: Sphaeria allicina Fr., Kongl. Vetensk. Acad. Handl. 38: 

247. 1817, sactioned by Fr., Syst. Mycol. 2: 437. 1823. 

≡ Sphaerella allicina (Fr. : Fr.) Auersw., in Gonn. & Rabenh., Mycol. 

Europaea 5–6: 19. 1869. 

Ascomata pseudothecial, black, superficial, situated on a small 

stroma, globose, up to 250 µm diam; ostioles periphysate, with 

apical periphysoids present; wall consisting of 3–6 layers of 

reddish brown textura angularis. Asci fasciculate, bitunicate, 

subsessile, obovoid to broadly ellipsoid, straight to slightly curved, 

8-spored, 65–90 × 16–25 µm; with pseudoparenchymatal cells 

of the hamathecium persistent. Ascospores tri- to multiseriate, 

overlapping, hyaline, with irregular lumina, thick-walled, straight 

to slightly curved, fusoid-ellipsoidal with obtuse basal end, and 

acutely rounded apical end, widest near the middle of the apical 

cell, medianly 1-septate, not to slightly constricted at the septum, 

(20–)25–27(–30) × (5.5–)6–7 µm. 

Mycelium superficial, hyphae branched, 1.5–8 µm wide, 

pluriseptate, broader hyphae usually slightly constricted at the 

septa and somewhat swollen, hyaline to subhyaline, almost smooth 

to somewhat verruculose or irregularly rough-walled, sometimes 

appearing to have a slime coat, walls unthickened. Conidiophores 

macronematous, sometimes also micronematous, arising as lateral 

or terminal branches from plagiotropous or ascending hyphae, erect, 

straight to more or less flexuous, sometimes geniculate, nodulose, 

usually with small head-like swellings, sometimes also with 

intercalary nodules, sometimes swellings protruding and elongated 

to one side, unbranched, occasionally branched, (7–)20–330 µm, 

sometimes even longer, (2–)3–5 µm wide, swellings (4–)5–8 µm 

wide, pluriseptate, not constricted at the septa, septa sometimes 

not very conspicuous, subhyaline to pale brown or pale olivaceous, 

smooth or somewhat verruculose, walls unthickened or almost so, 

more thickened with age. Conidiogenous cells integrated, usually 

terminal, cylindrical with a terminal head-like swelling, sometimes 

with a second swelling, 15–40 µm long, proliferation sympodial, 

with few conidiogenous loci confined to swellings, up to five per 

swelling, loci protuberant, conspicuous, 1–2 µm diam, thickened 

and darkened-refractive. Conidia catenate, formed in branched 

chains, straight to slightly curved, small terminal conidia subglobose, 

ovoid to obovoid or somewhat limoniform, 4–9 × 2.5–3.5 µm [av. ± 

SD, 6.5 (± 1.5) × 3.1 (± 0.5) µm], aseptate; secondary ramoconidia 

and occasionally formed ramoconidia ellipsoid to subcylindrical 

or cylindrical, 10–24(–31) × 3–5(–7) µm [av. ± SD, 16.1 (± 4.1) 

× 4.1 (± 0.8) µm], rarely up to 40 µm long, 0–1(–3)-septate, very 

rarely 5-septate, subhyaline to pale brown or pale olivaceous, 

minutely verruculose to verrucose (mostly granulate with some 

muricate projections under SEM), walls unthickened or almost so, 

apex rounded or slightly attenuated towards apex and base, hila 

protuberant, conspicuous, 1–2 µm wide, up to 1 µm high, thickened 

and darkened-refractive; microcyclic conidiogenesis occurring. 

118


Belgium, isolated from Quercus robur (Fagaceae), CBS 157.82; Kampenhout, 

isolated from Hordeum vulgare (Poaceae), 26 June 2005, J.Z. Groenewald, CBS-H 

19856, neotype designated here of C. bruhnei, isoneotype HAL 2023 F, cultures 

ex-type CBS 121624 = CPC 12211, CPC 12212. Czech Republic, Lisen, isolated 

from Polygonatum odoratum (Liliaceae), CBS 813.71, albino mutant of CBS 812.71. 

Germany, CBS 134.31 = ATCC 11283 = IMI 049632; Nordrhein-Westfalen, Mühlheim 

an der Ruhr, isolated from industrial water, IWW 727, CBS 110024; Sachsen-Anhalt, 

Halle (Saale), Robert-Franz-Ring, isolated from leaves of Tilia cordata (Tiliaceae), 

2004, K. Schubert, CPC 11386. Netherlands, isolated from air, CBS 521.68; isolated 

from Hordeum vulgare, 1 Jan. 2005, P.W. Crous, CPC 12139; isolated from man, 

skin, CBS 159.54 = ATCC 36948; Amsterdam, isolated from Thuja tincture, CBS 

177.71; Geleen, St. Barbara Ziekenhuis, isolated from man, skin, CBS 366.80, CBS 

399.80; isolated from man, sputum, Aug. 1955, CBS 161.55. New Zealand, Otago, 

Lake Harris, isolated from Ourisia macrophylla (Scrophulariaceae), 30 Jan. 2005, 

A. Blouin, Hill 1135, CPC 11840. Russia, Moscow region, isolated from Polyporus 

radiatus (Polyporaceae), Oct. 1978, CBS 572.78 = VKM F-405. Slovenia, Ljubljana, 

isolated from an air conditioning system, 2004, M. Butala, EXF-680 = CPC 12046; 

Sečovlje, isolated from hypersaline water from salterns (reserve pond), 2005, P. 

Zalar, EXF-389 = CPC 12042. Spain, Ebro Delta, isolated from hypersaline water 

from salterns (crystallisation pond), 2004, P. Zalar, EXF-594 = CPC 12045. Sweden, 

Skåne, on tip blight of living leaves of Allium sp. (Alliaceae), Fr. no. F-09810, UPS- 

FRIES, holotype of Davidiella allicina. U.S.A., New York, Geneva, isolated from 

CCA-treated Douglas-fir pole, CBS 115683 = ATCC 66670 = CPC 5101. 

Substrate and distribution: Living and decaying plant material, man, 

air, hypersaline and industrial water; widespread. 

Literature: Saccardo (1899: 1076), Linder (1947: 289). 

Fig. 10. Cladosporium bruhnei (CPC 12211). A. Conidiophore with characteristic 

long secondary ramoconidium and complex conidiophore. B. Detail of hila on 

secondary ramoconidia. C. Details of prominent ornamentation on conidia. Scale 

bars: A = 10 µm, B = 2 µm, C = 5 µm. 

Cultural characteristics: Colonies on PDA reaching 22–32 mm diam 

after 14 d at 25 ºC, olivaceous-grey to iron-grey, sometimes whitish, 

smoke-grey to pale olivaceous due to abundant aerial mycelium 

covering almost the whole colony, with age collapsing becoming 

olivaceous-grey, occasionally zonate, velvety to floccose, margin 

narrow, entire edge, white, glabrous to somewhat feathery, aerial 

mycelium sparse to abundant, white, fluffy, growth regular, flat to 

low convex, sometimes forming few exudates in the colony centre, 

sporulating. Colonies on MEA reaching 21–32 mm diam after 14 

d at 25 ºC, grey-olivaceous, olivaceous-grey to dull green or irongrey, 

sometimes whitish to pale smoke-grey due to abundant aerial 

mycelium, olivaceous-grey to iron-grey reverse, velvety, margin 

narrow, entire edge to slightly undulate, white, radially furrowed, 

glabrous to slightly feathery, aerial mycelium sparse to abundant, 

mainly in the centre, white, fluffy, growth convex to raised, radially 

furrowed, distinctly wrinkled in the colony centre, without prominent 

exudates, sporulating. Colonies on OA reaching 20–32 mm diam 

after 14 d at 25 ºC, smoke-grey, grey-olivaceous to olivaceous-grey, 

greenish black or iron-grey reverse, margin narrow, entire edge, 

colourless to white, glabrous, aerial mycelium sparse to abundant, 

dark smoke-grey, diffuse, high, later collapsed, felty, growth flat, 

without prominent exudates, sporulation profuse. 

Specimens examined: Sine loco et dato, CBS 188.54 = ATCC 11290 = IMI 

049638. Australia, N.S.W., Barrington Tops National Park, isolated from leaves 

of Eucalyptus stellulata (Myrtaceae), 3 Jan. 2006, B. Summerell, CPC 12921. 

Fig. 11. Davidiella allicina (F-09810, UPS-FRIES, holotype). Ascus and ascospores. 

Scale bar = 10 µm. P.W. Crous del. 


119


Fig. 12. Cladosporium bruhnei (CPC 12211) and its teleomorph Davidiella allicina. A–B. Macronematous conidiophores. C. Conidial chains. D. Micronematous conidiophore. E. 

Ascomata of the teleomorph formed on the host. F–G. Asci. Scale bars: A–B, D, F = 10 µm, E = 200 µm. 

Notes: Cladosporium bruhnei proved to be an additional component 

of the herbarum complex. The species resembles C. herbarum s. 

str. as already stated by Linder (1947), but possesses consistently 

narrower conidia, usually 2.5–5 µm wide, and the conidiophores 

often form only a single apical swelling. The species was described 

by Bruhne (l.c.) as Hormodendrum hordei from Germany but type 

material could not be located. Linder (1947) examined No. 1481a-5 

(Canada, N. Quebec, Sugluk, on Elymus arenarius var. villosus, 

31 Jul. 1936, E. Meyer), presumably in the National Museum, and 

stated that this specimen agreed well with the description and 

illustration given by Bruhne (l.c.). Although the species occurs 

on numerous substrates and is widely distributed, it has not yet 

been recognised as a distinct species since it has probably been 

interpreted as a narrow variant of C. herbarum. 

Based on morphology and DNA sequence data, the CBS strain 

CBS 177.71 chosen by Prasil & de Hoog (1988) as representative 

living strain of C. herbarum, rather clusters together with isolates 

of C. bruhnei. The strain CBS 813.71 is an albino mutant of the 

latter species as it does not appear to contain colour pigment. 

Furthermore, all isolates from humans treated until now as C. 

herbarum proved to be conspecific with the narrow-spored C. 

bruhnei. 

Although Davidiella tassiana (ascospores 17–25 × 6–8.5 µm, 

RO) was treated as synonymous to D. allicina (ascospores 20–27 

× 6–7 µm, UPS) in Aptroot (2006), they differ in apical ascospore 

taper, with ascospores of D. allicina being acutely rounded, while 

those of D. tassiana are obtusely rounded. The same ascospore 

taper was also observed in the teleomorph of C. bruhnei, and thus 

the name D. allicina is herewith linked to C. bruhnei, which is distinct 

from C. herbarum, having D. tassiana as teleomorph. 

Cladosporium herbaroides K. Schub., Zalar, Crous & U. Braun, 

sp. nov. MycoBank MB504574. Figs 13–15. 

Etymology: Refers to its morphological similarity to Cladosporium 

herbarum. 

Differt a Cladosporio herbaro conidiis polymorphis, 3–33 × (2–)3–6(–7) µm, 

postremo latioribus, (3.5–)5–9(–11) µm, fuscis et crassitunicatis; et a Cladosporio 

macrocarpo conidiophoris leniter angustioribus, 3–5 µm latis, nodulis angustioribus, 

5–8 µm latis. 

Mycelium branched, (1–)2–8 µm wide, septate, often with small 

swellings and constrictions, subhyaline to pale brown or pale 

olivaceous-brown, smooth or almost so to somewhat verruculose, 

walls unthickened or almost so. Conidiophores macronematous 

and micronematous, arising lateral from plagiotropous hyphae or 

terminally from ascending hyphae. Macronematous conidiophores 

erect, straight to slightly flexuous, often geniculate, nodulose, with 

unilateral or multilateral swellings, often numerous swellings in 

short succession giving them a gnarled appearance, often forming 

somewhat protruding or prolonged lateral swellings or a branch- 

120


Fig. 13. Cladosporium herbaroides (CPC 12052). Macro- and micronematous conidiophores and conidia. Scale bar = 10 µm. K. Schubert del. 

like prolongation below the terminal swelling (due to sympodial 

proliferation), unbranched or sometimes branched, 30–230 µm 

long or even longer, 3–5 µm wide, swellings 5–8 µm wide, septate, 

not constricted at septa, pale to medium olivaceous-brown, 

smooth or almost so, walls slightly thickened. Conidiogenous cells 

integrated, terminal or intercalary, cylindrical, usually nodulose to 

nodose forming distinct swellings, sometimes geniculate, 15–55 

µm long, with numerous conidiogenous loci usually confined to 

swellings or situated on small lateral shoulders, sometimes on the 

top of short peg-like prolongations or denticles, loci protuberant, 

1–2 µm diam, thickened and darkened-refractive. Micronematous 

conidiophores much shorter, narrower, paler, neither nodulose nor 

geniculate, arising laterally from plagiotropous hyphae, often only 

as short lateral denticles or branchlets of hyphae, erect, straight, 

conical to cylindrical, unbranched, 3–65 × 2–3 µm, mostly aseptate, 

sometimes up to five septa, subhyaline, smooth, walls unthickened. 


to conidiogenous cells, conidiogenous loci solitary or sometimes 

as sympodial clusters of pronounced denticles, protuberant, 1–1.5 

µm diam, thickened and somewhat darkened-refractive. Conidia 

polymorphous, two main morphological types recognisable, formed 


by the two different types of conidiophores, conidia formed by 

macronematous conidiophores catenate, in branched chains, 

straight to slightly curved, subglobose, obovoid, limoniform, ellipsoid 

to cylindrical, 3–33 × (2–)3–6(–7) µm [av. ± SD, 14.5 (± 7.9) × 

5.2 (± 1.2) µm], 0–2(–3)-septate, sometimes slightly constricted at 

septa, septa median or somewhat in the lower half, pale to medium 

olivaceous-brown, verruculose to verrucose (granulate under 

SEM), walls slightly thickened, with up to three rarely four distal 

scars, with age becoming medium or even dark brown (chocolate 

brown), wider and more thick-walled, 5.5–33 × (3.5–)5–9(–11) µm 

[av. ± SD, 14.4 (± 6.9) × 7.2 (± 1.9) µm], walls up to 1 µm thick, 

hila protuberant, 0.8–2(–2.5) µm diam, thickened and darkenedrefractive; 

microcyclic conidiogenesis occurring. Conidia formed by 

micronematous conidiophores paler and narrower, mostly formed 

in unbranched chains, sometimes in branched chains with up to 

three distal hila, straight to slightly curved, limoniform, narrowly 

fusiform, almost filiform to subcylindrical, 10–26(–35) × 2–3.5 

µm [av. ± SD, 15.6 (± 6.2) × 2.9 (± 0.5) µm], 0–1(–3)-septate, 

subhyaline to pale brown, almost smooth to minutely verruculose, 

walls unthickened, hila protuberant, 1–1.5 µm diam, thickened and 

somewhat darkened-refractive. 

121


Fig. 14. Cladosporium herbaroides (CPC 12052). A–B, D. Macronematous conidiophores. C. Conidial chain. E. Micronematous conidiophore. F. Microcyclic conidiogenesis. G. 

Conidia formed by micronematous conidiophores. Scale bars = 10 µm. 

Cultural characteristics: Colonies on PDA attaining 23 mm diam 

after 14 d at 25 ºC, grey-olivaceous to olivaceous, olivaceous-grey 

reverse, velvety, margin regular, entire edge, narrow, feathery, aerial 

mycelium abundantly formed, loose, with age covering large parts 

of the colony, woolly, growth flat with somewhat elevated colony 

centre, folded, regular, deep into the agar, with few prominent 

exudates, sporulation profuse. Colonies on MEA attaining 24 mm 

diam after 14 d at 25 ºC, grey- to greenish olivaceous, olivaceousgrey 

or iron-grey reverse, velvety to powdery, margin narrow, 

colourless, entire edge, somewhat feathery, aerial mycelium pale 

olivaceous-grey, sparse, growth convex, radially furrowed, folded 

in the colony centre, without prominent exudates, sporulating. 

Colonies on OA attaining 23 mm diam after 14 d at 25 ºC, greyolivaceous, 

margin more or less regular, entire edge, colourless, 

somewhat feathery, aerial mycelium whitish to smoke grey, at first 

sparse, later more abundantly formed, growth flat, without exudates, 

sporulation profuse. 

Specimen examined: Israel, from hypersaline water of Eilat salterns, 2004, coll. 

N. Gunde-Cimerman, isol. M. Ota, CBS-H 19858, holotype, isotype HAL 2025 F, 

culture ex-type CBS 121626 = EXF-1733 = CPC 12052. 

Substrate and distribution: Hypersaline water; Israel. 

Notes: Cladosporium herbaroides is morphologically similar to C. 

herbarum but differs in having somewhat longer conidia becoming 

wider, darker and even more thick-walled with age [at first conidia 

3–33 × (2–)3–6(–7) µm, with age (3.5–)5–9(–11) µm wide]. Besides 

that, the species often produces a second conidial type formed on 

micronematous conidiophores, giving rise to unbranched conidial 

chains which are almost filiform, limoniform, narrowly fusiform to 

subcylindrical, much narrower and paler than the ones formed by 

macronematous conidiophores, 10–26(–35) × 2–3.5 µm. In C. 

herbarum, conidia formed by micronematous conidiophores do not 

occur as frequently as in C. herbaroides, and differ in being often 

clavate and somewhat wider, up to 4(–5) µm wide. Cladosporium 

macrocarpum is easily distinguishable by having somewhat wider 

conidiophores (3–)4–6 µm, with distinctly wider swellings, 5–10 µm 

wide, and the conidia are usually (3–)5–9(–10) µm wide. 

Cladosporium herbarum (Pers. : Fr.) Link, Ges. Naturf. Freunde 

Berlin Mag. Neuesten Entdeck. Gesammten Naturk. 7: 37. 1816: 

Fr., Syst. mycol. 3(2): 370. 1832. Figs 16–19. 

Basionym: Dematium herbarum Pers., Ann. Bot. (Usteri) 11: 32. 

1794: Fr., Syst. mycol. 3(2): 370. 1832. 

= Dematium epiphyllum var. (β) chionanthi Pers., Mycol. eur. 1: 16. 1822, syn. 

nov. 

For additional synonyms see Dugan et al. (2004), Schubert 

(2005). 

Teleomorph: Davidiella tassiana (De Not.) Crous & U. Braun, 

Mycol. Progr. 2: 8. 2003. 

Basionym: Sphaerella tassiana De Not., Sferiacei Italici 1: 87. 

1863. 

≡ Mycosphaerella tassiana (De Not.) Johanson, Öfvers. Förh. Kongl. 

Svenska Vetensk.-Akad. 41: 167. 1884. 

122


Fig. 15. Cladosporium herbaroides (CPC 12052). A. Overview of the growth characteristics of this fungus. Broad hyphae run over the surface of the agar, and possibly give rise 

to conidiophore branches. The conidiophores of this fungus can be rather long, resembling aerial hyphae. Clusters of conidia are clearly visible in this micrograph. B. The very 

wide surface hyphae can anastomose. C. Conidiophore with secondary ramoconidia and conidia. Note the variation in scar size. D. A very elaborate, complex conidiophore with 

different scars of variable size, one being more than 2 μm wide! E. Details of secondary ramoconidia and hila. Note the rather strong ornamentation in which smaller “particles” 

are between larger ones. F. Three conidia in a row. Note the scar formation in the chain and the reduction of the size of the cells throughout the spore-chain. The inset shows 

the resemblance of the scars on a conidiophore and on a secondary ramoconidium. Scale bars: A = 50 µm, B–C, F (inset) = 10 µm, D–E = 5 µm, F = 2 µm. 

Ascomata pseudothecial, black, globose, erumpent to superficial, 

up to 200 µm diam, with 1(–3) short, periphysate ostiolar necks; 

wall consisting of 3–6 layers of medium red-brown textura 

angularis. Asci fasciculate, bitunicate, subsessile, obovoid to 

broadly ellipsoid, straight to slightly curved, 8-spored, 65–85 × 13– 

17 µm. Pseudoparaphyses absent in host material, but remnants 

observed when studied in culture, hyaline, septate, subcylindrical, 

anastomosing, 3–4 µm wide. Ascospores tri- to multiseriate, 

overlapping, hyaline, with irregular luminar inclusions, thickwalled, 

straight to slightly curved, fusoid-ellipsoidal with obtuse 

ends, widest near middle of apical cell, medianly 1-septate, not to 

slightly constricted at the septum, tapering towards both ends, but 

more prominently towards the lower end, (17–)20–23(–25) × (6–) 

7(–8) µm; becoming brown and verruculose in asci. Ascospores 

germinating after 24 h on MEA from both ends, with spore body 

becoming prominently constricted at the septum, but not distorting, 

up to 7 µm wide, hyaline to pale brown and appearing somewhat 

verruculose, enclosed in a mucoid sheath, with germ tubes being 

irregular, somewhat nodular. 


123


almost smooth to minutely verruculose or irregularly rough-walled, 

sometimes forming clavate conidia, up to 33 µm long, 0–2-septate. 


to conidiogenous cells, narrowly cylindrical or filiform, with a single 

or two loci. Conidia catenate, in unbranched or loosely branched 

chains with branching mostly occurring in the lower part of the chain, 

straight to slightly curved, small terminal conidia without distal hilum 

obovoid, 4–10 × 3–5(–6) µm [av. ± SD, 7.8 (± 1.9) × 4.7 (± 0.9) µm], 

aseptate, intercalary conidia with a single or sometimes up to three 

distal hila limoniform, ellipsoid to subcylindrical, 6–16 × 4–6 µm 

[av. ± SD, 12.4 (± 1.6) × 5.3 (± 0.6) µm], 0–1-septate, secondary 

ramoconidia with up to four distal hila, ellipsoid to cylindrical-oblong, 

12–25(–35) × (3–)5–7(–9) µm [av. ± SD, 18.8 (± 4.5) × 6.2 (± 0.9) 

µm], 0–1(–2)-septate, rarely with up to three septa, sometimes 

distinctly constricted at the septum, septum median or somewhat 

in the upper or lower half, pale greyish brown or brown to medium 

brown or greyish brown, minutely verruculose to verrucose, walls 

slightly to distinctly thickened, guttulate to somewhat granular, 

usually only slightly attenuated towards apex and base, apex 

obtuse or slightly truncate, towards the base sometimes distinctly 

attenuated with hila situated on short stalk-like prolongations, hila 

slightly to distinctly protuberant, truncate to slightly convex, (0.8–) 

1–2.5(–3) µm wide, 0.5–1 µm high, somewhat thickened and 

darkened-refractive; microcyclic conidiogenesis occurring, conidia 

forming micro- and macronematous secondary conidiophores. 

Fig. 16. Davidiella tassiana (RO, holotype). Ascus and ascospores. Scale bar = 10 

µm. P.W. Crous del. 

Mycelium superficial, loosely branched, (0.5–)1–5 µm wide, septate, 

sometimes constricted at septa, hyaline, subhyaline to pale brown, 

smooth or almost so to verruculose or irregularly rough-walled, 

sometimes appearing irregular in outline due to small swellings 

and constrictions, walls unthickened to somewhat thickened, cell 

lumen appearing to be granular. Conidiophores both macro- and 

micronematous, arising laterally from plagiotropous hyphae or 

terminally from ascending hyphae. Macronematous conidiophores 

erect, straight to flexuous, somewhat geniculate-sinuous, nodulose 

to nodose with unilateral or multilateral swellings, with a single to 

numerous swellings in short succession giving the stalk a knotty/ 

gnarled appearance, unbranched or occasionally branched, up to 

three times, sometimes with a lateral branch-like proliferation below 

or at the apex, 10–320 × 3.5–5 µm, swellings 5–8(–9) µm wide, 

pluriseptate, septa sometimes constricted when formed after a 

node, pale to medium brown, older ones almost dark brown, paler 

towards the apex, smooth or minutely verruculose, walls thickened, 

sometimes even two-layered. Conidiogenous cells integrated, 

terminal or intercalary, nodulose to nodose, with a single or up 

to five swellings per cell, 10–24 µm long, proliferation sympodial, 

with several conidiogenous loci confined to swellings, mostly 

situated on small lateral shoulders, more or less protuberant, 

broadly truncate to slightly convex, 1.5–2.5 µm diam, thickened 

and somewhat darkened-refractive. Micronematous conidiophores 

hardly distinguishable from hyphae, sometimes only as short lateral 

outgrowth with a single apical scar, short, conical to almost filiform 

or narrowly cylindrical, non-nodulose, not geniculate, unbranched, 

5–120 × 1.5–3(–4) µm, pluriseptate, not constricted at septa, 

cells usually very short, 5–15 µm long, subhyaline to pale brown, 


after 14 d at 25 ºC, grey-olivaceous to olivaceous-grey, whitish 

to smoke-grey or pale olivaceous-grey due to abundant aerial 

mycelium, velvety, reverse olivaceous-grey or iron-grey, margin 

almost colourless, regular, entire edge, glabrous to feathery, 

aerial mycelium abundant mainly in the colony centre, dense, 

felty, woolly, sometimes becoming somewhat reddish brown, 

fawn coloured, growth regular, flat to low convex with an elevated 

colony centre, sometimes forming few large prominent exudates, 

sporulation profuse. Colonies on MEA reaching 17–37 mm diam 

after 14 d at 25 ºC, smoke-grey to pale olivaceous-grey towards 

margin, olivaceous-grey to iron-grey reverse, velvety, margin white, 

entire edge to slightly undulate, aerial mycelium abundant, dense, 

fluffy to felty, growth low convex or raised, radially furrowed, folded 

and wrinkled in the colony centre, without prominent exudates but 

sporulating. Colonies on OA reaching 12–28 mm diam after 14 

d at 25 ºC, olivaceous-grey to iron-grey, due to abundant aerial 

mycelium pale olivaceous-grey, olivaceous-grey reverse, margin 

narrow, more or less undulate, white, aerial mycelium white, loose to 

dense, high, fluffy to felty, covering large parts of the colony, growth 

flat to low convex, without prominent exudates, sporulating. 

Specimens examined: Sine loco, sine dato, L 910.225-733, lectotype of C. 

herbarum, selected by Prasil & de Hoog, 1988. Sine loco, on leaves of Chionanthus 

sp. (Oleaceae), L 910.255-872 = L-0115833, holotype of Dematium epiphyllum 

var. (β) chionanthi. Netherlands, Wageningen, isolated from Hordeum vulgare 

(Poaceae), 2005, P.W. Crous, CBS-H 19853, epitype designated here of C. 

herbarum and D. tassiana, isoepitype HAL 2022 F, ex-type cultures, CPC 12177 = 

CBS 121621, CPC 12178–12179, 12181, 12183. Italy, on upper and lower surface 

of dead leaves of Carex nigra [“fusca”] (Cyperaceae), Tassi no. 862, RO, holotype 

of Davidiella tassiana. U.S.A., Colorado, San Juan Co., above Little Molas Lake, 

isolated from stems of Delphinium barbeyi (Ranunculaceae), 12 Sep. 2004, A. 

Ramaley, CBS-H 19868 (teleomorph), single ascospore isolates, CBS 121622 = 

CPC 11600, CPC 11601–11604. 

Substrate and distribution: On fading and decaying plant material, 

on living leaves (phylloplane fungus), as secondary invader, as an 

endophyte, isolated from air, soil, foodstuffs, paints, textiles and 

numerous other materials; cosmopolitan. 

124


Fig. 17. Cladosporium herbarum (CPC 11600). Macro- and micronematous conidiophores and conidia. Scale bar = 10 µm. K. Schubert del. 

Literature: de Vries (1952: 71), Hughes (1958: 750), Ellis (1971: 

313), Domsch et al. (1980: 204), Sivanesan (1984: 225), Ellis & 

Ellis (1985: 290, 468, 1988: 168), Prasil & de Hoog (1988), Wang 

& Zabel (1990: 202), McKemy & Morgan-Jones (1991), Dugan & 

Roberts (1994), David (1997: 59), Ho et al. (1999: 129), de Hoog et 

al. (2000: 587), Samson et al. (2000: 110), Samson et al. (2001). 

Notes: De Vries (1952) incorrectly selected a specimen of Link’s 

herbarium at herb. B as lectotype. Prasil & de Hoog (1988) discussed 

this typification and designated one of Persoon’s original specimens 

as lectotype in which C. herbarum could be recognised. The latter 

material, which is in poor condition, could be re-examined within 

the course of these investigations and showed conidia agreeing 

with the current species concept of C. herbarum being (6–)9.5– 

14.5(–21) × (5–)6–7(–8) µm. Since the identity of the strain CBS 

177.71 chosen by Prasil & de Hoog (1988) as representative living 

strain of C. herbarum could not be corroborated, an epitype with 

a living ex-epitype culture is designated. The holotype specimen 

of D. tassiana (RO) is morphologically similar to that observed on 

the epitype of C. herbarum, having ascospores which are (17–)21– 


23(–25) × (6–)7–8(–8.5) µm, turning brown and verruculose in asci 

with age. However, no hamathecial remnants were observed in 

ascomata in vivo. 

The connection to the teleomorph D. tassiana could be 

confirmed, which is in agreement with the findings of von Arx (1950) 

and Barr (1958). Ascospore isolates formed the typical C. herbarum 

anamorph in culture, and these anamorph cultures developed 

some immature fruiting bodies within the agar. When inoculated 

onto water agar plates with nettle stems, numerous ascomata with 

viable ascospores were formed in culture. 

Cladosporium iridis (Fautrey & Roum.) G.A. de Vries, Contr. 

Knowl. Genus Cladosporium: 49. 1952. Figs 20–21. 

Basionym: Scolicotrichum iridis Fautrey & Roum., Rev. Mycol. 

(Toulouse) 13: 82. 1891. 

≡ Heterosporium iridis (Fautrey & Roum.) J.E. Jacques, Contr. Inst. Bot. 

Univ. Montréal 39: 18. 1941. 

For additional synonyms see Dugan et al. (2004). 

Teleomorph: Davidiella macrospora (Kleb.) Crous & U. Braun, 

Mycol. Progr. 2: 10. 2003. 

125


Fig. 18. Cladosporium herbarum (CPC 11600) and its teleomorph Davidiella tassiana (from the host and CPC 12181). A–B. Macronematous conidiophores. C. Micronematous 

conidiophore. D. Microcyclic conidiogenesis. E. Conidial chain. F. Ascomata on the leaf. G. Ascomata formed in culture on nettle stems. H–I. Asci on the host. J–K. Ascospores 

in culture. L. Asci in culture. Scale bars: A, E, H, J–L = 10 µm, F–G, I = 200 µm. 

Basionym: Didymellina macrospora Kleb., Ber. Deutsch. Bot. Ges. 

42: 60, 1924. 1925. 

≡ Mycosphaerella macrospora (Kleb.) Jørst., Meld. Stat. Plantepatol. Inst. 

1: 20. 1945. 

Mycelium branched, 2–8 µm wide, septate, not constricted at the 

septa, hyaline to pale brown, smooth, walls slightly thickened, 

sometimes guttulate. Conidiophores very long, usually terminally 

arising from ascending hyphae, erect to subdecumbent, slightly 

to distinctly flexuous, geniculate-sinuous, usually several times, 

subnodulose due to geniculate, sympodial proliferation forming 

swollen lateral shoulders, unbranched, rarely branched, up to 

720 µm long, 6–11 µm wide, swellings 8–11(–14) µm wide, 

pluriseptate, often very regularly septate, not constricted at 

the septa, pale to medium olivaceous-brown, somewhat paler 

towards the apex, smooth to minutely verruculose, walls only 

slightly thickened. Conidiogenous cells integrated, terminal as 

well as intercalary, cylindrical-oblong, 15–55 µm long, proliferation 

percurrent to sympodial, usually with a single geniculation forming 

laterally swollen shoulders often below a septum, conidiogenous 

loci confined to swellings, usually one locus per swelling, rarely 

two, protuberant, (2–)2.5–4 µm diam, somewhat thickened 

and darkened-refractive. Conidia solitary, sometimes in short, 

unbranched chains, straight to curved, young conidia pyriform to 

subcylindrical, connection between conidiophore and conidium 

being rather broad, subhyaline to pale olivaceous-brown, walls 

slightly thickened, then enlarging and becoming more thick-walled, 

cylindrical-oblong, soleiform with age, both ends rounded, usually 

with a slightly to distinctly bulbous base, visible from a very early 

stage, but broadest part often towards the apex not at the base, (18–) 

30–75(–87) × (7–)10–16(–18) µm [av. ± SD, 53.3 (± 17.8) × 12.6 (± 

2.2) µm], (0–)2–6(–7)-septate, usually not constricted at the septa, 

rarely slightly constricted, septa often becoming sinuous with age, 

pale to medium olivaceous-brown, sometimes darker, verrucose to 

echinulate, walls thickened, especially in older conidia, up to 1 µm 

thick, hila protuberant, often stalk-like or conically prolonged, up 

to 2 µm long, (2–)2.5–3.5(–4) µm diam, with age becoming more 

sessile, sometimes just visible as a thickened plate just below the 

outer wall layer, especially in distal scars of branched conidia, 

periclinal rim often distinctly visible, hila somewhat thickened and 

darkened-refractive; microcyclic conidiogenesis not observed. 

126


Fig. 19. Cladosporium herbarum (CPC 11600). A. Overview of hyphal growth and conidiophore formation of a colony on SNA. Conidiophores are often formed on very wide 

(approx. 10 μm), septate hyphae that often grow near the agar surface. B. A more detailed view on colony organisation reveals the ornamented conidia. Note the septum near the 

conidiophore (arrow). C. Detail of spore ornamentation and hila on a secondary ramoconidium (arrow). Ornamentation is visible during early stages of spore formation (arrow). 

D. Structure of the conidiophore, illustrating the complex morphology of the spore-forming apparatus. In addition, secondary ramoconidia, conidia, and a hilum on the conidium 

are visible. E. Complex structure of the spore-forming apparatus. F. Details of secondary ramoconidia with complex scar-pattern on the right cell. G. Details of a secondary 

ramoconidium giving rise to conidia. Note the lack of ornamentation at the location of spore formation. Scale bars: A = 50 µm, B, F = 10 µm, C–E, G = 5 µm. 

Cultural characteristics: Colonies on PDA reaching 19–23 mm 

diam after 14 d at 25 ºC, pale greenish olivaceous, smoke-grey 

to olivaceous-grey due to abundant aerial mycelium, greenish 

olivaceous to olivaceous reverse, margin broad, regular, entire 

edge to slightly undulate, feathery, aerial mycelium abundantly 

formed, felty, fluffy, covering large parts of the colony, mainly in 

the central parts, high, growth low convex with a somewhat raised 

colony centre. Colonies on MEA reaching 9–23 mm diam after 14 d 

at 25 ºC, pale olivaceous-grey to olivaceous-grey, olivaceous-grey 

reverse, felty, margin slightly undulate, white, somewhat raised, 

aerial mycelium abundant, loose, diffuse, high, growth low convex, 

radially furrowed, slightly folded. Colonies on OA reaching 10–19 

mm diam after 14 d at 25 ºC, olivaceous, margin broad, undulate, 

white, aerial mycelium white, very high, loose, diffuse, hairy, growth 

flat, due to the mycelium low convex, without prominent exudates 

and sporulating on all media. 

Specimens examined: Isolated from Iris sp. (Iridaceae), CBS 107.20. France, 

Cote d’Or, Jardin de Noidan, on leaves of Iris germanica, Jul. 1880, F. Fautrey, 

Roumeguère, Fungi Sel. Gall. Exs. No. 5689, PC, lectotype of C. iridis, selected 

by David, 1997; K, isolectotype. Netherlands, Boterenbrood, isolated from leaves 

of Iris sp., Aug. 1940, CBS-H 19859, epitype designated here of C. iridis, culture 

ex-epitype CBS 138.40. 


127


Fig. 20. Cladosporium iridis (CBS 138.40). Conidiophores and conidia. Scale bar = 10 µm. K. Schubert del. 

Fig. 21. Cladosporium iridis (teleomorph Davidiella macrospora) (CBS 138.40). A–C. Conidiophores with conidia. D. Conidium. Scale bar = 10 µm. 

128


Fig. 22. Cladosporium macrocarpum (CBS 299.67). Macro- and micronematous conidiophores and conidia. Scale bar = 10 µm. K. Schubert del. 

Substrates and distribution: Leaf spot and blotch of Iris spp. 

including I. crocea, I. florentina, I. foetidissima, I. germanica, I. 

gueldenstaedtiana, I. kamaonensis, I. pallida, I. plicata (= I. swertii 

Hort.), I. pseudacorus, I. pumila, I. spuria ssp. halophila, and other 

species, also on Belacamanda chinensis (= Gemmingia chinensis), 

Hemerocallis fulva, Gladiolus gandavensis; Africa (Algeria, 

Morocco, South Africa, Zambia, Zimbabwe), Asia (Armenia, 

Azerbaijan, China, Georgia, India, Iran, Israel, Japan, Kazakhstan, 

Kirgizstan, Korea, Russia, Turkey, Turkmenistan, Uzbekistan), 

Australasia (Australia, New Zealand), Europe (Austria, Belgium, 

Belorussia, Cyprus, Czech Republic, Denmark, Estonia, France, 

Germany, Great Britain, Greece, Italy, Latvia, Lithuania, Malta, 

Moldavia, Montenegro, Netherlands, Norway, Poland, Romania, 

Russia, Serbia, Spain, Sweden, Ukraine), North America (Canada, 

U.S.A.), Central & South America (Argentina, Chile, Jamaica, 

Panama, Uruguay). 

Literature: Ellis (1971: 312), Ellis & Waller (1974), Sivanesan (1984: 

222), McKemy & Morgan-Jones (1990), David (1997: 43), Shin et 

al. (1999). 

Notes: The description of the morphological parameters in culture 

is based on the isolate sporulating on PDA, since sporulation on 

SNA was not observed. The conidiophores and conidia in vivo are 

usually wider than in culture [conidiophores (6–)9–15(–17) µm 

wide, conidia (11–)15–23(–28) µm]. 


Cladosporium macrocarpum Preuss, in Sturm, Deutsch. Fl. 

3(26): 27. 1848. Figs 22–25. 

≡ Cladosporium herbarum var. macrocarpum (Preuss) M.H.M. Ho & 

Dugan, in Ho et al., Mycotaxon 72: 131. 1999. 

= Dematium herbarum var. (β) brassicae Pers., Syn. meth. fung. 2: 699. 1801, 

syn. nov. 

= Dematium graminum Pers., Mycol. eur. 1: 16. 1822, syn. nov. 

= Dematium vulgare var. (δ) typharum Pers., Mycol. eur. 1: 14. 1822, syn. 

nov. 

= Dematium vulgare var. (β) foliorum Pers., Mycol. eur. 1: 14. 1822, syn. nov. 

For additional synonyms see Dugan et al. (2004), Schubert 

(2005). 

Teleomorph: Davidiella macrocarpa Crous, K. Schub. & U. Braun, 

sp. nov. MycoBank MB504582. 

Davidiellae tassianae similis, sed pseudoparaphysibus prominentibus et ascosporis 

maioribus, (22–)23–26(–28) × (6–)6.5–7(–8) µm. 

Ascomata superficial on a small stroma, black, up to 200 µm 

diam, globose, separate, but developing with 1–3 necks with age; 

ostioles consisting of pale brown to subhyaline cells, periphysate, 

with periphysoids growing into the cavity; wall consisting of 3–6 

layers of medium brown textura angularis. Pseudoparaphyses 

present, hyaline, subcylindrical, septate, anastomosing, 3–4 

µm diam; hamathecial cells persistent in cavity. Asci fasciculate, 

bitunicate, subsessile, broadly ellipsoid with a long tapered stalk, 

straight to curved, 8-spored, 70–110 × 15–20 µm. Ascospores 

tri- to multiseriate, overlapping, hyaline, guttulate, irregular lumina 

129


Fig. 23. Cladosporium macrocarpum (CBS 299.67) and its teleomorph Davidiella macrocarpa (CPC 12755). A–C. Macronematous conidiophores and conidia. D–G. 

Micronematous conidiophores. H. Microcyclic conidiogenesis. I. Ascomata formed on nettle stems in culture. J. Periphyses. K, M–N. Asci. L. Ostiole. Scale bars: A, D–H, J–N 

= 10 µm, I = 200 µm. 

rarely observed, thick-walled, straight to slightly curved, fusoidellipsoidal 

with obtuse ends, widest in the middle of the apical 

cell, medianly 1-septate, not to slightly constricted at the septum, 

tapering towards both ends, but more prominently towards lower 

end, (22–)23–26(–28) × (6–)6.5–7(–8) µm; mucoid sheath rarely 

observed, mostly absent. 

Mycelium unbranched or loosely branched, 1–4.5(–5) µm wide, 

septate, sometimes slightly constricted at septa, hyaline to pale 

brown, smooth to minutely verruculose, walls unthickened or slightly 

thickened. Conidiophores micronematous and macronematous, 

solitary, arising terminally from plagiotropous hyphae or terminally 

from ascending hyphae. Macronematous conidiophores erect, 

straight to somewhat flexuous, cylindrical-oblong, nodulose to 

nodose, with a single apical or usually several swellings either 

somewhat distinct from each other or often in short succession giving 

conidiophores a knotty appearance, swellings sometimes laterally 

elongated or formed at the top of a branch-like outgrowth below 

the apical swelling, sometimes distinctly geniculate, unbranched, 

sometimes branched, 12–260 × (3–)4–6 µm, swellings 5–10 µm 

wide, pluriseptate, sometimes slightly constricted at septa, pale to 

medium brown or olivaceous-brown, somewhat paler at apices, 

130


Fig. 24. Cladosporium macrocarpum (CBS 299.67). A. Survey of a conidiophore that forms several secondary ramoconidia and conidia. Several aerial hyphae are also visible 

in this picture. B. Conidiophore with broadly ellipsoid secondary ramoconidia and obovoid conidia. Note the different scars on the conidiophore at the lower left. C. Ellipsoid 

or obovoid conidia with notable areas of scar formation. The ornamentation is relatively widely distributed over the body of the cell and similar to C. variabile. D. Detail of 

a conidiophore (see B) with scars. Note the relatively shallow rings of the scars. E. Details of conidia and a secondary ramoconidium. F. Conidiophore with a secondary 

ramoconidium and conidia. Note the hila on several spores and the lack of ornamentation at the site where spores are formed. Scale bars: A–C, = 10 µm, D, F = 5 µm, E = 2 

µm. 

smooth to minutely verruculose or verruculose, walls somewhat 

thickened, sometimes even two-layered. Conidiogenous cells 

integrated, terminal or intercalary, cylindrical, nodulose with lateral 

shoulders or nodose with swellings round about the stalk, with 

conidiogenous loci confined to swellings, 12–37 µm long, with up to 

12 loci per cell, usually with up to six, loci conspicuous, protuberant, 

(1–)1.5–2 µm diam, somewhat thickened and darkened-refractive. 

Micronematous conidiophores almost indistinguishable from 

hyphae, straight, narrowly filiform, non-nodulose or with a single or 

few swellings, mostly with small head-like swollen apices, usually 


131


or branched chains, subglobose, obovoid to limoniform, ellipsoid 

or fusiform, 2.5–16 × 1.5–5 µm, 0(–1)-septate, few longer conidia 

subcylindrical to clavate, up to 37(–43) µm long, 0–2(–3)-septate, 

occasionally with up to four septa, sometimes slightly constricted 

at the septa, subhyaline to pale brown, almost smooth to minutely 

verruculose, walls unthickened, hila 0.8–1.2 µm diam, thickened 

and darkened-refractive. 


in diam after 14 d at 25 ºC, dark dull green to olivaceous-grey, 

olivaceous-grey, dark olivaceous- to iron-grey reverse, pulvinate, 

velvety, sometimes somewhat zonate, paler zones towards the 

margin, margin regular, entire edge, almost colourless to white, 

glabrous to feathery, aerial mycelium sparse to more abundant 

in the colony centre or covering large areas of the colony, hairy, 

fluffy or felty, whitish to smoke-grey, sometimes becoming reddish, 

livid red to vinaceous, growth flat, regular, sometimes forming 

few prominent exudates, exudates sometimes slightly reddish, 

sporulation profuse with two kinds of conidiophores, low and high. 

Colonies on MEA reaching 31–50 mm in diam after 14 d at 25 

ºC, grey-olivaceous to olivaceous-grey or iron-grey, sometimes 

pale olivaceous-grey to whitish due to abundant aerial mycelium, 

olivaceous-grey or iron-grey reverse, velvety or powdery, margin 

narrow, entire edge, colourless to white, glabrous, aerial mycelium 

sparse to abundant, hairy or felty, growth regular, flat to low convex, 

radially furrowed, without prominent exudates, sporulation profuse. 

Colonies on OA reaching 29–40 mm in diam after 14 d at 25 ºC, 

grey-olivaceous, olivaceous-grey to dark smoke-grey, olivaceousblack 

or iron grey reverse, margin entire edge, narrow, colourless 

or white, glabrous, aerial mycelium sparse, mainly in the colony 

centre, felty, white to smoke-grey or grey-olivaceous, felty, growth 

flat, regular, without exudates, sporulating. 

Fig. 25. Davidiella macrocarpa (CPC 12755). Ascus and ascospores. Scale bar = 

10 µm. P.W. Crous del. 

only few micrometer long, 1.5–3 µm wide, aseptate or with only 

few septa, subhyaline, smooth or almost so, walls unthickened, 

with a single or only few conidiogenous loci, narrow, 0.8–1.2 µm 

diam, thickened and somewhat darkened-refractive. Conidia 

catenate, in branched chains, small terminal conidia subglobose, 

obovoid, oval, limoniform, 4–11 × (3–)4–6 µm [av. ± SD, 7.6 (± 

1.9) × 5.0 (± 0.8) µm], aseptate, intercalary conidia broadly ovoidellipsoid, 

10–17 × (4.5–)5–9 µm [av. ± SD, 12.7 (± 2.1) × 6.8 (± 

0.8) µm], 0–1-septate; secondary ramoconidia broadly ellipsoid 

to subcylindrical, 14–25(–30) × (5–)6–9(–10) µm [av. ± SD, 19.4 

(± 3.5) × 7.6 (± 1.0) µm], 0–2(–3)-septate, sometimes slightly 

constricted at the septa, septa somewhat sinuous with age, pale 

brown to medium olivaceous-brown or brown, sometimes even 

dark brown, verruculose to echinulate (muricate under SEM), 

walls thickened, up to 1 µm thick, mostly broadly rounded at 

apex and base, sometimes attenuated, sometimes guttulate 

by oil drops, with up to three apical hila, mostly 1–2, hila sessile 

(apparently somewhat immersed) to somewhat protuberant, 1– 

2(–2.5) µm diam, thickened and darkened-refractive; microcyclic 

conidiogenesis occurring with conidia forming secondary microand 

macronematous conidiophores, conidia often germinating with 

long hyphae. Conidia formed by micronematous conidiophores 

usually smaller, narrower and paler, catenate, in short unbranched 

Specimens examined: Sine loco et dato, L 910.255-723 = L-0115836, lectotype 

designated here of Dematium graminum. Sine loco, on dead stems of Brassica sp. 

(Brassicaceae), No. 601, L 910.255-716 = L-0115849, holotype of D. herbarum var. 

(β) brassicae. Sine loco, on leaves of Iris (Iridaceae), Quercus (Fagaceae), Brassica 

etc., L 910.255-736 = L-0115871, holotype of D. vulgare var. (β) foliorum, isotype L 

910.255-718 = L-0115872. Sine loco et dato, L 910.255-698 = L-0115852, lectotype 

designated here for D. vulgare var. (δ) typharum. Isolated from “Mycosphaerella 

tulasnei”, CBS 223.32 = ATCC 11287 = IMI 049635. Romania, isolated from water, 

CBS 175.82. Slovenia, Sečovlje, isolated from hypersaline water from salterns 

(precrystalisation pond), 2004, P. Zalar, EXF-2287 = CPC 12054. Turkey, Ankara, 

Tekeli, isolated from Triticum aestivum (Poaceae), isol. S. Tahsin, ident. A.C. Stolk, 

CBS 299.67. U.S.A., Seattle, University of Washington Campus, 47.6263530, - 

122.3331440, isolated from cleistothecia of Phyllactinia guttata (Erysiphaceae) 

on leaves of Corylus sp. (Corylaceae), 16 Sep. 2004, D. Glawe, CPC 11817; 

Washington, isolated from Spinacia oleracea (Chenopodiaceae), 1 Jan. 2003, 

L. DuToit, CBS-H 19855, neotype designated here for C. macrocarpum, and 

holotype of D. macrocarpa, isoneotype HAL 2020 F, isotype HAL 2021 F, culture 

ex-type CPC 12752, 12756–12759, CPC 12755 = CBS 121623. 

Substrate and distribution: Decaying plant material, human, 

hypersaline water, water; widespread. 

Literature: de Vries (1952: 76), Ellis (1971: 315), Domsch et al. 

(1980: 208), Ellis & Ellis (1985: 290, 468), Matsushima (1985: 5), 

McKemy & Morgan-Jones (1991), Dugan & Roberts (1994), David 

(1997: 71), Samson et al. (2000: 112). 

Notes: In the absence of Preuss’s type material (not preserved) 

de Vries (1952) “lectotypified” C. macrocarpum by a specimen in 

Saccardo’s herbarum (Herb. Myc. P.A. Saccardo no. 419, PAD). 

This material, subsequently distributed in Mycotheca Italica no. 

1396, should correctly be regarded as neotype (David 1997). A 

single collection of Saccardo’s Mycotheca Italica no. 1396 from 

herb. HBG, which can be considered as isoneotype material, 

132


Fig. 26. Cladosporium ossifragi (CBS 842.91). Conidiophores and conidia. Scale bar = 10 µm. K. Schubert del. 

was re-examined and proved to rather agree with the species 

concept of C. herbarum s. str. The conidia were formed in simple, 

rarely branched chains, 6–26 × (4–)5.5–8(–9) µm, 0–3-septate, 

almost smooth or minutely to densely verruculose or verrucose 

(Schubert 2005). However, since de Vries’ “lectotypification” was 

incorrect according to the code (ICBN, Art. 9.2, 9.17), a neotype is 

designated. 

The delimitation of C. macrocarpum as a morphologically 

distinct species from C. herbarum has been controversially 

discussed by several authors (McKemy & Morgan-Jones 1991, 

Dugan & Robert 1994, Ho et al. 1999). Based on molecular as well 

morphological studies, it can be shown that C. macrocarpum is a 

well-defined species distinguishable from C. herbarum s. str. by 

forming conidiophores with wider nodes, 5–10 µm, wider and more 

frequently septate conidia [small terminal conidia 4–11 × (3–)4–6 

µm versus 4–10 × 3–5(–6) µm in C. herbarum, intercalary conidia 

10–17 × (4.5–)5–9 µm versus 6–16 × 4–6 µm in C. herbarum, 

secondary ramoconidia 14–25(–30) × (5–)6–9(–10) µm versus 12– 

25(–35) × (3–)5–7(–9) µm in C. herbarum] and by being connected 

to Davidiella macrocarpa. On natural substrates the conidiophores 

are usually somewhat wider than in culture, 4–8(–10) µm wide, and 

also the conidia can be somewhat wider, sometimes up to 13(–15) µm. 

Cladosporium graminum, described by Persoon (1822), as 

well as C. brunneum and C. gracile, introduced by Corda (1837), 

are older synonyms of C. macrocarpum and, according to the 

code, would have priority. However, since C. macrocarpum is a 

well established, currently used name with numerous records in 

literature, a proposal to conserve the name against these older 

names is in preparation for formal publication in Taxon. 

A characteristic difference between ascomata of C. 

macrocarpum in comparison to those of C. herbarum, are the 

smaller, globose pseudothecia, asci with longer stalks, prominence 

of pseudoparaphyses, and rather inconspicuous luminar ascospore 

inclusions. 

Cladosporium ossifragi (Rostr.) U. Braun & K. Schub., comb. 


Basionym: Napicladium ossifragi Rostr., Bot. Fǽröes 1: 316. 

1901. 

≡ Heterosporium ossifragi (Rostr.) Lind, Dan. fung.: 531. 1913. 

= Heterosporium magnusianum Jaap, Schriften Naturwiss. Vereins Schleswig- 

Holstein 12: 346. 1902. 

≡ Cladosporium magnusianum (Jaap) M.B. Ellis in Ellis, More 



133


Fig. 27. Cladosporium ossifragi (CBS 842.91). A. Macronematous conidiophore. B. Micronematous conidiophore. C–D. Conidia. E. Conidia and microcyclic conidiogenesis. 


Fig. 28. Cladosporium ossifragi (CBS 842.91). A. Survey on different secondary ramoconidia and conidia. B. Details of conidia and hila. Note the very pronounced ornamentation 

and the absence of ornamentation near the site of spore formation. C. Detail of the end of a secondary ramoconidium with pronounced hila. D. Formation of a new conidium. 

Note the broad scar behind it (> 1 μm). E. Formation of a new conidium from a smooth-walled stalk. F. Hila on a secondary ramoconidium. This micrograph is from the sample 

before coating with gold-palladium and shows similar features as the sample after sputter coating. Scale bars: A = 10 µm, B–D, F = 2 µm, E = 5 µm. 

134


Mycelium abundantly formed, twisted, often somewhat aggregated, 

forming ropes, branched, 1–5 µm wide, septate, often irregularly 

swollen and constricted, hyaline or subhyaline to pale brown, 

smooth, walls unthickened or only slightly thickened. Conidiophores 

macronematous and micronematous, arising from plagiotropous 

hyphae, terminally or laterally, erect to subdecumbent, more 

or less straight to flexuous, cylindrical, sometimes geniculate, 

subnodulose with loci often situated on small lateral shoulders, 

unbranched, sometimes branched, often very long, up to 350 µm 

long, 3–4.5(–5) µm wide, pluriseptate, shorter ones aseptate, not 

constricted at septa, pale to pale medium brown, paler towards 

apices, sometimes subhyaline, smooth to minutely verruculose, 

especially towards apices, walls somewhat thickened, up to 0.5 µm, 

sometimes appearing two-layered. Conidiogenous cells integrated, 

terminal as well as intercalary, cylindrical, sometimes geniculate, 

subnodulose, 5–31 µm long, proliferation sympodial, with few 

loci (1–3) per cell, loci usually confined to small lateral shoulders, 

protuberant, conspicuous, short cylindrical, 1–2 µm wide, up to 

1 µm high, somewhat thickened, darkened-refractive. Conidia 

catenate, in short, unbranched or branched chains, straight, small 

terminal and intercalary conidia subglobose, obovoid to ellipsoid, 

4–15 × 3–5 µm [av. ± SD, 9.3 (± 3.7) × 4.0 (± 0.7) µm], 0–1- 

septate, not constricted at the septa, pale brown, hila 0.8–1 µm 

diam, secondary ramoconidia cylindrical, sometimes ellipsoid or 

subfusiform, 16–36(–40) × (4–)5–8 µm [av. ± SD, 26.6 (± 7.4) × 

6.0 (± 1.2) µm], (0–)1–3(–4)-septate [in vivo wider, (6–)7–9(–11) 

µm, and with up to five, rarely seven septa], not constricted at 

the septa, septa sometimes slightly sinuous, pale brown to pale 

medium brown, densely verruculose, verrucose to echinulate 

(densely muricate under SEM), walls unthickened to somewhat 

thickened, rounded or somewhat attenuated at apex and base, 

hila protuberant, conspicuous, sometimes situated on short, small 

prolongations, 1–2.5 µm diam, somewhat thickened and darkenedrefractive; 

microcyclic conidiogenesis occasionally occurring. 

Cultural characteristics: Colonies on PDA reaching 53 mm diam 

after 14 d at 25 ºC, greenish olivaceous, grey-olivaceous to 

olivaceous-grey or iron-grey, appearing somewhat zonate, dull 

green to olivaceous-black reverse, margin colourless, regular, 

entire edge, aerial mycelium abundantly formed, covering at first 

the colony centre later most of the surface, dense, high, growth flat 

with elevated colony centre, somewhat folded. Colonies on MEA 

reaching 54 mm diam after 14 d at 25 ºC, pale olivaceous-grey 

to olivaceous-grey in the centre, iron-grey reverse, velvety, margin 

colourless to white, entire edge, radially furrowed, aerial mycelium 

abundantly formed, fluffy to felty, growth flat with somewhat raised, 

folded colony centre. Colonies on OA attaining 52 mm diam after 

14 d at 25 ºC, olivaceous-grey to iron-grey, iron-grey to greenish 

black reverse, margin white, entire edge, aerial mycelium diffuse, 

loose, growth flat, prominent exudates absent, sporulation profuse 

on all media. 

Specimens examined: Denmark, Undallslund, on leaves of Narthecium ossifragum 

(Melanthiaceae), 13 Sep. 1885, E. Rostrup, CP, neotype designated here of C. 

ossifragi; Tǿnder, Rǿmǿ near Twismark, 19 Aug. 1911, H. Sydow, Sydow, Mycoth. 

Germ. 1047, M. Germany, Hamburg, Eppendorfer Moor, on leaves of Narthecium 

ossifragum, 12 Sep. 1897, O. Jaap, HBG, lectotype selected here of C. 

magnusianum; 4 Sep. 1903, O. Jaap, Jaap, Fungi Sel. Exs. 49, M; Wernerwald near 

Cuxhaven, Aug. 1927, A. Ludwig, Petrak, Mycoth. Gen. 146, M. Norway, Bjerkreim 

County, isolated from leaves of Narthecium ossifragum, M. di Menna, CBS-H 19860, 

epitype designated here of C. ossifragi, culture ex-epitype CBS 842.91 = ATCC 

200946; Møre og Romsdal County, isolated from leaves of Narthecium ossifragum, 

M. di Menna, CBS 843.91. 

Substrate and distribution: Causing leaf spots on Narthecium 

ossifragum; Europe (Austria, Denmark, Germany, Great Britain, 

Ireland, Norway). 

Literature: Ellis & Ellis (1985: 390), David (1995a; 1997: 85–86, 

88), Ho et al. (1999: 132). 

Notes: Type material of Napicladium ossifragi is not preserved in 

Rostrup’s herbarium (on Narthecium ossifragum, Faeroe Islands, 

Viderö, Viderejde and Österö, Svinaa, sine dato, leg. Ostenfeld & 

Harz). However, other authentic collections seen and examined 

by Rostrup are deposited at CP. Lind (1913) re-examined these 

samples, synonymised N. ossifragi with H. magnusianum and 

correctly introduced the combination H. ossifragi. Nevertheless, 

the correct oldest name for this fungus has been ignored by 

most authors. David (1997), who clearly stated that N. ossifragi is 

the earliest name for this species, preferred to use the name C. 

magnusianum because the typification of Rostrup’s name was still 

uncertain. Despite the lacking type material, there is no doubt about 

the correct identity of N. ossifragi since authentic material of this 

species, examined by and deposited in Rostrup’s herbarium (CP), 

is preserved. Therefore, there is no reason to reject the oldest valid 

name for this species. The original collection of C. magnusianum 

cited by Jaap (1902) (on leaves of Narthecium ossifragum, Denmark, 

Tǿnder, Rǿmǿ, peatbog by Twismark, Jul.–Aug. 1901, Jaap), but 

not designated as type, is not preserved (David 1997). It is neither 

deposited at B, HBG nor S. However, in the protologue Jaap (1902) 

also referred to material of this species found near Hamburg, which 

is, hence, syntype material available for lectotypification. 

Cladosporium pseudiridis K. Schub., C.F. Hill, Crous & U. Braun, 


Etymology: Epithet derived from its similar morphology to 

Cladosporium iridis. 

Differt a Cladosporio iridis conidiis 0–3-septatis, brevioribus et latioribus, 15–55 × 

(9–)11–19(–21) µm. 

Mycelium sparingly branched, 2–7 µm wide, septate, not constricted 

at the septa, subhyaline to pale brown, smooth or almost so, walls 

somewhat thickened, guttulate or protoplasm appearing granular, 

sometimes enveloped by a slime coat. Conidiophores arising 

mostly terminally from ascending hyphae, sometimes also laterally 

from plagiotropous hyphae, erect, more or less straight, broadly 

cylindrical-oblong, once or several times slightly to distinctly 

geniculate-sinuous, forming more or less pronounced lateral 

shoulders, nodulose, unbranched, 100–320(–500) × 7–11 µm, 

swellings 10–14 µm wide, becoming narrower and paler towards the 

apex, septate, not constricted at the septa, septa mainly basal, apical 

cell often very long, pale to medium olivaceous-brown, subhyaline 

at the apex, smooth or almost so, sometimes minutely verruculose, 

walls usually distinctly thickened, sometimes even two-layered, up 

to 1(–2) µm thick, protoplasm granular, often clearly contrasting 

from the outer wall. Conidiogenous cells integrated, terminal and 

intercalary, cylindrical-oblong, slightly to distinctly geniculatesinuous, 

nodulose with conidiogenous loci confined to swellings 

or lateral shoulders, 30–110 µm long, proliferation percurrent to 

sympodial, with a single or three, sometimes up to five geniculations 

per cell, usually only a single locus per swelling, protuberant, very 

prominent, short cylindrical, peg-like, clearly composed of a dome 

and surrounding rim, dome often higher than the periclinal rim, 

broad, somewhat paler than rim, conically narrowed, (2–)2.5–4 

µm wide, up to 2 µm high, thickened and darkened-refractive. 


135


Fig. 29. Cladosporium pseudiridis (CBS 116463). Conidiophores and conidia. Scale bar = 10 µm. K. Schubert del. 

Conidia solitary, sometimes in short unbranched chains of two or 

three, straight to slightly curved, young conidia small, 0–1-septate, 

broadly ovoid to pyriform, 15–26 × (9–)11–16(–18) µm [av. ± SD, 

19.2 (± 4.3) × 14.2 (± 3) µm], first septum somewhat in the upper 

half, the upper cell is much smaller but gradually extending as the 

conidium matures, mature conidia 1–3-septate, broadly pyriform, 

cylindrical-oblong or soleiform, usually with a distinctly bulbous 

base, 30–55 × 12–19(–21) µm [av. ± SD, 41.5 (± 6.8) × 17.1 (± 2.1) 

µm], broadest part of conidia usually at the bulbous base, mostly 

attenuated towards the basal septum, septa becoming sinuous with 

age, pale to medium olivaceous-brown or brown, usually echinulate, 

sometimes coarsely verrucose, walls distinctly thickened, up to 2 

µm thick, often appearing layered with a large lumen in the centre 

of the cell, broadly rounded to flattened at apex and base, hila often 

very prominent, often peg-like elongated, up to 3 µm long, with age 

becoming less prominent, visible as a thickened flat plate just below 

the outer echinulate wall layer, slightly raised towards the middle, 

2–3.5 µm diam, thickened and darkened-refractive; microcyclic 

conidiogenesis not observed. 


after 14 d at 25 ºC, whitish, smoke-grey to pale olivaceous-grey 

due to abundant aerial mycelium, olivaceous-black reverse, margin 

narrow, white, more or less crenate, aerial mycelium zonate, fluffy, 

covering most of the colony, mainly in the colony centre, growth 

136


Fig. 30. Cladosporium pseudiridis (CBS 116463). A–C. Conidiophores and conidia. D. Part of a conidiogenous cell showing a protuberant cladosporioid conidiogenous locus. 

E–F. Conidia. Scale bars = 10 µm. 

convex to raised, deep into the agar, with age few large prominent 

exudates formed, sparingly sporulating. Colonies on MEA attaining 

7 mm diam after 14 d at 25 ºC, olivaceous-grey, pale olivaceousgrey 

to pale rosy-buff due to abundant aerial mycelium covering 

almost the whole colony, iron-grey reverse, margin colourless or 

white, broad, regular, more or less glabrous, aerial mycelium fluffy, 

dense, high, growth convex to umbonate, sometimes with elevated 

colony centre, prominent exudates lacking, sporulation sparse. 

Colonies on OA attaining 8 mm diam after 14 d at 25 ºC, white, pale 

buff to pale olivaceous-grey in the centre, margin grey-olivaceous, 

olivaceous- to iron-grey reverse, margin entire edge or somewhat 

undulate, somewhat feathery, growth raised with a somewhat 

depressed centre forming an elevated outer rim, without prominent 

exudates, sporulation more abundant. 

Specimen examined: New Zealand, Auckland, Mt. Albert, Carrington Road, Unitec 

Campus, isolated from large leaf lesions on Iris sp. (Iridaceae), 15 Aug. 2004, C.F. 

Hill, CBS-H 19861, holotype, culture ex-type CBS 116463 = LYN 1065 = ICMP 

15579. 

Substrate and distribution: On living leaves of Iris sp.; New 

Zealand. 

Notes: Cladosporium pseudiridis closely resembles C. iridis, a 

common and widespread species causing leaf spots on numerous 

Iris spp. and a few additional hosts of the host family Iridaceae, 

but the latter species is easily distinguishable by having longer 

and narrower, more frequently septate conidia, (18–)30–75(–87) × 

(7–)10–16(–18) µm, (0–)2–6(–7)-septate. 


It is unlikely that C. pseudiridis is of New Zealand origin since 

the genus Iris is not indigenous to New Zealand. All Iris species 

that are found in this country have been introduced, mainly for 

horticultural purposes. The species is, therefore, probably more 

common than indicated above. However, within the course of the 

recent monographic studies in the genus Cladosporium numerous 

herbarium specimens, mainly of European origin, have been 

examined and proved to be correctly identified agreeing with the 

species concept of C. iridis. Additional collections and cultures are 

necessary to determine its distribution. 

Cladosporium ramotenellum K. Schub., Zalar, Crous & U. Braun, 


Etymology: Refers to the morphological similarity with Cladosporium 

tenellum. 

Differt a Cladosporio cladosporioide conidiophoris et conidiis leniter angustioribus, 

2–4(–5) µm latis, conidiis 0–2(–3)-septatis, semper verruculosis; et a Cladosporio 

tenello locis conidiogenis non numerosis et non aggregatos ad apicem, conidiis 

longioribus et angustioribus, 2.5–35 × 2–4(–5) µm, 0–3-septatis. 

Mycelium unbranched or only sparingly branched, 1.5–4 µm wide, 

septate, without swellings and constrictions, hyaline or subhyaline, 

smooth, sometimes irregularly rough-walled, walls unthickened. 

Conidiophores solitary, macronematous and micronematous, 

arising as lateral branches of plagiotropous hyphae or terminally 

from ascending hyphae, erect, straight or slightly flexuous, 

cylindrical, neither geniculate nor nodulose, without head-like 

swollen apices or intercalary swellings, unbranched, sometimes 

137


Fig. 31. Cladosporium ramotenellum (CPC 12043). Conidiophores and conidia. Scale bar = 10 µm. K. Schubert del. 

branched, branches often only as short lateral prolongations, 

mainly formed below a septum, 14–110 × 2–4 µm, septate, not 

constricted at the septa, subhyaline to pale olivaceous or brown, 

smooth to minutely verruculose, walls unthickened, sometimes 

guttulate. Conidiogenous cells integrated, terminal, sometimes also 

intercalary, cylindrical, not geniculate, non-nodulose, 10–28(–50) 

µm long, proliferation sympodial, with few conidiogenous loci, 

mostly 1–3, loci sometimes situated on small lateral prolongations, 

protuberant, 0.5–1.5(–2) µm diam, thickened and somewhat 

darkened-refractive. Ramoconidia formed, cylindrical-oblong, up 

to 47 µm long, 2–4 µm wide, 0–1-septate, rarely up to 4-septate, 

subhyaline to very pale olivaceous, smooth or almost so, with a 

broadly truncate base, without any dome and raised rim, 2–3 µm 

wide, not thickened but somewhat refractive. Conidia numerous, 

polymorphous, catenate, in branched chains, straight, sometimes 

slightly curved, small terminal conidia numerous, globose, 

subglobose or ovoid, obovoid or limoniform, 2.5–7 × 2–4(–4.5) µm 

[av. ± SD, 5.1 (± 1.3) × 3.1 (± 0.6) µm], aseptate, without distal 

hilum or with a single apical scar, intercalary conidia ellipsoid to 

subcylindrical, 8–15 × 3–4(–4.5) µm [av. ± SD, 11.5 (± 2.4) × 3.6 

(± 0.5) µm], 0–1-septate; secondary ramoconidia subcylindrical 

to cylindrical-oblong, 17–35 × 3–4(–5) µm [av. ± SD, 22.5 (± 

5.6) × 3.7 (± 0.5) µm], 0–3-septate, not constricted at the septa, 

subhyaline to very pale olivaceous, minutely verruculose (granulate 

under SEM), walls unthickened or almost so, apex broadly rounded 

or slightly attenuated towards apex and base, sometimes guttulate, 

hila protuberant, conspicuous, 0.8–1.5(–2) µm diam, somewhat 

thickened and darkened-refractive; microcyclic conidiogenesis 

occurring. 


after 14 d at 25 ºC, olivaceous to grey-olivaceous due to abundant 

sporulation, appearing zonate in forming concentric zones, margin 

entire edge to slightly undulate, white, glabrous, aerial mycelium 

absent or sparse, growth flat with a somewhat folded and wrinkled 

colony centre, without prominent exudates, sporulation profuse. 

Colonies on MEA reaching 48–49 mm diam after 14 d at 25 ºC, 

grey-olivaceous to olivaceous-grey, velvety, olivaceous-grey to 

138


Fig. 32. Cladosporium ramotenellum (CPC 12043). A, C. Macronematous conidiophore. B. Conidial chain. D. Micronematous conidiophore. E. Ramoconidia and conidia. Scale 

bars = 10 µm. 

Fig. 33. Cladosporium ramotenellum (CPC 12043). A. Survey of colony development showing a large bulbous “foot cell” that gives rise to conidiophores, which can be branched. 

B. Details of conidiophores showing secondary ramoconidia and conidia. The inset shows scar formation on a conidiophore. C. Conidiophore and several conidia. D. Details 

of ornamentation on conidia. Note the wide, but relatively low ornamentation units. E. A micrograph illustrating the organisation within a conidiophore. Scale bars A–D = 5 µm, 

E = 10 µm. 


139


Fig. 34. Cladosporium sinuosum (CPC 11839). Conidiophores and conidia. Scale bar = 10 µm. K. Schubert del. 

iron-grey reverse, margin entire edge to undulate, radially furrowed, 

colourless, glabrous to feathery, aerial mycelium sparse, diffuse, 

growth flat with slightly elevated colony centre, distinctly wrinkled, 

prominent exudates not formed, abundantly sporulating. Colonies 

on OA attaining 40 mm diam after 14 d at 25 ºC, grey-olivaceous, 

margin entire edge, colourless or white, glabrous, aerial mycelium 

absent or sparse, growth flat, without exudates, sporulation 

profuse. 

Specimens examined: Slovenia, Ljubljana, isolated from an air conditioning system 

(bathroom), 2004, M. Butala, CBS 121627 = CPC 12047 = EXF-967; Sečovlje, 

isolated from hypersaline water from reverse ponds, salterns, 2005, P. Zalar, CBS-H 

19862, holotype, isotype HAL 2026 F, culture ex-type CBS 121628 = CPC 12043 

= EXF-454. 

Substrate and distribution: Hypersaline water, air; Slovenia. 

Notes: Cladosporium ramotenellum, which appears to be a 

saprobe in air and hypersaline water, morphologically resembles 

C. cladosporioides and C. tenellum K. Schub., Zalar, Crous & 

U. Braun, but is quite distinct from C. cladosporioides by having 

somewhat narrower conidiophores and conidia, 2–4(–5) µm 

wide, and 0–3-septate, always minutely verruculose conidia. 

Cladosporium tenellum, a newly introduced species (see below) 

isolated from hypersaline water and plant material, possesses 

conidiophores with numerous conidiogenous loci, usually crowded 

towards the apex forming sympodial clusters of pronounced scars, 

and shorter and somewhat wider, 0–1(–2)-septate conidia, 3–20(– 

28) × (2.5–)3–5(–6) µm. Besides these morphological differences, 

C. ramotenellum is faster growing in culture than C. tenellum. 

Cladosporium arthrinioides Thüm. & Beltr. and C. hypophyllum 

Fuckel are also close to C. ramotenellum, but C. arthrinioides, 

known from Italy on leaves of Bougainvillea spectabilis, deviates in 

having shorter and wider, 0–1(–2)-septate, mostly smooth conidia 

(2–18 × 2–6.5 µm) which become larger and more frequently 

septate with age (up to 32 µm long and with up to four septa); 

and C. hypophyllum occurring in Europe on leaves of Ulmus 

minor differs in having often mildly to distinctly geniculate-sinuous, 

sometimes subnodulose conidiophores and shorter and somewhat 

wider, 0–1(–3)-septate conidia, 4–17(–19) × 2–5 µm, becoming 

distinctly swollen, darker, longer and wider with age, 5–7 µm, with 

the septa often being constricted (Schubert 2005). 

140


Fig. 35. Cladosporium sinuosum (CPC 11839). A–D. Conidiophores. E–F. Conidia. Scale bars = 10 µm. 

Cladosporium sinuosum K. Schub., C.F. Hill, Crous & U. Braun, 


Etymology: Refers to the usually distinctly sinuous conidiophores. 

Differt a Cladosporio herbaro conidiophoris distincte sinuosis, conidiis solitariis vel 

breve catenatis, catenis non ramosis, echinulatis. 

Mycelium sparingly branched, 1–7 µm wide, septate, not 

constricted at the septa, subhyaline to pale brown, smooth to 

minutely verruculose, walls unthickened or slightly thickened, 

sometimes with small swellings. Conidiophores arising laterally 

from plagiotropous hyphae or terminally from ascending hyphae, 

erect, more or less straight to flexuous, often once or several times 

slightly to distinctly geniculate-sinuous, sometimes even zigzag-like, 

nodulose with small to large lateral shoulders, shoulders somewhat 

distant from each other or in close succession giving them a knotty/ 

gnarled appearance, unbranched or once branched, 25–260 × 5–7 

µm, shoulders up to 10 µm wide, pluriseptate, septa sometimes 

in short succession, not constricted at the septa, pale brown to 

medium brown, smooth to minutely verruculose, walls thickened, 

often distinctly two-layered, up to 1 µm thick. Conidiogenous 

cells integrated, terminal or intercalary, often slightly to distinctly 

geniculate-sinuous, nodulose with small to large laterally swollen 

shoulders, 8–30 µm long, proliferation sympodial, with a single 

or up to three conidiogenous loci, usually confined to lateral 


shoulders, protuberant, often denticle-like or on the top of short 

cylindrical stalk-like prolongations, 1.2–2(–2.2) µm diam, mainly 

2 µm, somewhat thickened and darkened-refractive, dome often 

slightly higher than the surrounding rim. Conidia solitary or in short 

unbranched chains with up to three conidia, straight, obovoid, oval, 

broadly ellipsoid to subcylindrical or sometimes clavate (broader at 

the apex), 9–21 × (5–)6–8 µm [av. ± SD, 14.5 (± 2.5) × 6.6 (± 0.7) 

µm], 0–1-septate, not constricted at the septa, septum more or less 

median, pale greyish brown, densely echinulate, spines up to 1 µm 

long, walls thickened, apex mostly broadly rounded or sometimes 

attenuated, towards the base mostly distinctly attenuated forming a 

peg-like prolongation, up to 2 µm long, hila protuberant, 1.2–2 µm 

diam, mainly 2 µm, somewhat thickened and darkened-refractive; 

microcyclic conidiogenesis not observed. 

Cultural characteristics: Colonies on PDA attaining 20 mm 

diam after 14 d at 25 ºC, pale olivaceous-grey due to abundant 

aerial mycelium, olivaceous-grey towards margins, iron-grey 

to olivaceous-black reverse, margin regular, entire edge, aerial 

mycelium abundant, cottony, dense, high, growth regular, low 

convex, radially furrowed in the centre, growing deep into the agar, 

with age numerous small to large prominent exudates, sporulation 

sparse. Colonies on MEA attaining 16 mm diam after 14 d at 25 ºC, 

white to pale smoke-grey, fawn reverse, velvety, margin undulate, 

glabrous, aerial mycelium abundant, dense, high, fluffy, growth 

raised with elevated colony centre, laterally furrowed, without 

141


Fig. 36. Cladosporium spinulosum (CPC 12040). A. Overview on agar surface with conidiophores arising from the surface. The spore clusters on the conidiophore are very 

compact. Note several simple, tubular conidiophore ends. The inset shows details of a conidium showing two pronounced hila and a unique, very distinct ornamentation on the 

cell wall. B. Conidiophore with globose or subsphaerical secondary ramoconidia and conidia. Note the newly forming cells and hila. C. Two conidiophores. D. Details of spores 

and spore formation. E. The end of a conidiophore and two scars. Scale bars: A = 20 µm, A (inset) = 1 µm, B, D–E = 5 µm, C = 10 µm. 

prominent exudates. Colonies on OA attaining 18 mm diam after 14 

d at 25 ºC, olivaceous, white to pale olivaceous-grey in the centre 

due to abundant aerial mycelium, olivaceous-grey reverse, margin 

white, entire edge, glabrous, aerial mycelium loose to dense, high, 

fluffy to felty, growth flat to low convex, regular, without prominent 

exudates, sporulating. 

Specimen examined: New Zealand, Te Anau, isolated from leaves of Fuchsia 

excorticata (Onagraceae), 31 Jan. 2005, A. Blouin, Hill 1134A, CBS-H 19863, 

holotype, culture ex-type CBS 121629 = CPC 11839 = ICMP 15819. 

Substrate and distribution: On living leaves of Fuchsia excorticata; 

New Zealand. 

Notes: This new species is well characterised by its slightly to 

distinctly geniculate-sinuous, often zigzag-like conidiophores and 

its conidia formed solitary or rarely in short unbranched chains and 

is therefore morphologically not comparable with any of the species 

described until now. Most Cladosporium species with conidia usually 

formed solitary or in short unbranched chains have previously 

been treated as species of the genus Heterosporium Klotzsch ex 

Cooke, now considered to be synonymous with Cladosporium. All 

of them, including the newly introduced C. arthropodii K. Schub. 

& C.F. Hill from New Zealand, which also belongs to this species 

complex (Braun et al. 2006), possess very large and wide, often 

pluriseptate conidia quite distinct from those of C. sinuosum (David 

1997). Cladosporium alopecuri (Ellis & Everh.) U. Braun, known 

from the U.S.A. on Alopecurus geniculatus is also quite different by 

having larger and wider conidia, 20–40 × 7–13(–15) µm, and wider 

conidiogenous loci and conidial hila, 3.5–5 µm diam (Braun 2000). 

Cladosporium herbarum is superficially similar but the 

conidiophores of the latter species are sometimes only slightly 

geniculate-sinuous but never zigzag-like and the verruculose to 

verrucose conidia are frequently formed in unbranched or branched 

chains. 

Cladosporium spinulosum Zalar, de Hoog & Gunde-Cimerman, 

Studies in Mycology 58: 180. 2007 – this volume. Fig. 36. 

Note: This new species is described and illustrated in Zalar et al. 

(2007 – this volume). 

142


Fig. 37. Cladosporium subinflatum (CPC 12041). Macro- and micronematous conidiophores and conidia. Scale bar = 10 µm. K. Schubert del. 

Cladosporium subinflatum K. Schub., Zalar, Crous & U. Braun, 


Etymology: Refers to its nodulose conidiophores. 

Differt a Cladosporio bruhnei conidiophoris cum nodulis angustioribus, 3–6.5 µm 

latis, conidiis brevioribus, 4–17(–22) µm longis, spinulosis, cum spinulis ad 0.8 µm 

longis; et a Cladosporio spinuloso conidiophoris nodulosis, conidiis spinulosis, cum 

spinulis brevioribus, ad 0.8 longis, locis conidiogenis et hilis latioribus, (0.5–)1–2 

µm latis. 

Mycelium unbranched or occasionally branched, 1.5–3 µm wide, 

later more frequently branched and wider, up to 7 µm wide, 

septate, not constricted at the septa, hyaline or subhyaline, almost 

smooth to somewhat verruculose or irregularly rough-walled, walls 

unthickened. Conidiophores mainly macronematous, sometimes 

also micronematous, arising terminally from ascending hyphae 

or laterally from plagiotropous hyphae, erect or subdecumbent, 

straight or flexuous, sometimes bent, cylindrical, nodulose, usually 

with small head-like swellings, sometimes swellings also on a 

lower level or intercalary, occasionally geniculate, unbranched, 

occasionally branched, (5–)10–270 × (1.5–)2.5–4.5(–5.5) µm, 


swellings 3–6.5 µm wide, aseptate or with few septa, not constricted 

at the septa, pale brown, pale olivaceous-brown or somewhat 

reddish brown, smooth, usually verruculose or irregularly roughwalled 

and paler, subhyaline towards the base, walls thickened, 

sometimes appearing even two-layered, up to 1 µm thick. 

Conidiogenous cells integrated, usually terminal or conidiophores 

reduced to conidiogenous cells, cylindrical, nodulose, usually with 

small head-like swellings with loci confined to swellings, sometimes 

geniculate, 5–42 µm long, proliferation sympodial, with several loci, 

up to four situated at nodules or on lateral swellings, protuberant, 

conspicuous, denticulate, (0.8–)1–2 µm diam, thickened and 

darkened-refractive. Conidia catenate, in branched chains, more 

or less straight, numerous globose and subglobose conidia, ovoid, 

obovoid, broadly ellipsoid to cylindrical, 4–17(–22) × (2.5–)3.5– 

5.5(–7) µm [av. ± SD, 11.7 (± 4.6) × 4.5 (± 0.8) µm], 0–1(–2)- 

septate, not constricted at septa, pale brown or pale olivaceousbrown, 

ornamentation variable, mainly densely verruculose to 

echinulate (loosely muricate under SEM), spines up to 0.8 µm 

high, sometimes irregularly verrucose with few scattered tubercles 

143


Fig. 38. Cladosporium subinflatum (CPC 12041). A–C. Macronematous conidiophores. D–E. Conidia. Scale bar = 10 µm. 

Fig. 39. Cladosporium subinflatum 

(CPC 12041). A–G. Images of an 11- 

d-old culture on SNA. A. Overview of 

colony with clusters of conidia and 

aerial hyphae. Many of the hyphae 

have a collapsed appearance. B. 

Detail of colony with conidiophores, 

conidia and aerial hyphae that 

are partly collapsed. C. Detail of a 

conidiophore end and a secondary 

ramoconidium. Note the scars at the 

end of the conidiophore. D. Details 

of conidia and ornamentation. 

The ornamentation consists out of 

markedly defined units, which have 

a relatively large distance from each 

other. Note the hilum on the right 

conidium. E. Conidiophore with 

large scars and conidia. F. Different 

blastoconidia with very early stages 

of new spore formation in the middle 

of the picture. G. Pattern of spore 

development. Scale bars: A = 20 

µm, B, E–G = 5 µm, C = 10 µm, D 

= 2 µm. 

144


or irregularly echinulate, walls unthickened or slightly thickened, 

apex rounded or slightly attenuated towards apex and base, hila 

conspicuous, protuberant, denticulate, 0.5–2 µm diam, thickened 

and darkened-refractive; microcyclic conidiogenesis observed. 


after 14 d at 25 ºC, olivaceous-black to olivaceous-grey towards 

margin, margin regular, entire edge, narrow, colourless to white, 

glabrous to feathery, aerial mycelium formed, fluffy, mainly near 

margins, growth flat, somewhat folded in the colony centre, deep 

into the agar, few prominent exudates formed with age, sporulation 

profuse. Colonies on MEA attaining 25 mm diam after 14 d at 25 

ºC, olivaceous-grey to olivaceous due to abundant sporulation 

in the colony centre, pale greenish grey towards margin, irongrey 

reverse, velvety to powdery, margin crenate, narrow, white, 

glabrous, radially furrowed, aerial mycelium diffuse, growth convex 

with papillate surface, wrinkled colony centre, without prominent 

exudates, sporulation profuse. Colonies on OA attaining 26 mm 

diam after 14 d at 25 ºC, olivaceous, iron-grey to greenish black 

reverse, growth flat, deep into the agar, with a single exudate, 

abundantly sporulating. 

Specimen examined: Slovenia, Sečovlje, isolated from hypersaline water from 

crystallization ponds, salterns, 2005, S. Sonjak, CBS-H 19864, holotype, isotype 

HAL 2027 F, culture ex-type CBS 121630 = CPC 12041 = EXF-343. 

Substrate and distribution: Hypersaline water; Slovenia. 

Notes: Cladosporium subinflatum, an additional saprobic species 

isolated from hypersaline water, was at first identified as C. 

spinulosum, but proved to be both morphologically as well as 

phylogenetically distinct from the latter species in having somewhat 

wider [(1.5–)2.5–4.5(–5.5) µm], nodulose macronematous 

conidiophores with conidiogenous loci confined to swellings, wider 

conidiogenous loci and hila, (0.8–)1–2 µm, and spiny conidia 

with shorter spines than in C. spinulosum (up to 0.8 µm versus 

0.5–1.3 µm long) (Zalar et al. 2007). With its narrow, nodulose 

macronematous conidiophores and catenate conidia, C. bruhnei 

is morphologically also similar but differs by having conidiophores 

with wider swellings, (4–)5–8 µm, and longer conidia 4–24(–31) 

µm, rarely up to 40 µm long which are minutely verruculose to 

verrucose but not spiny. 

Fig. 40. Cladosporium subtilissimum (CBS 113754). Macro- and micronematous conidiophores and conidia. Scale bar = 10 µm. K. Schubert del. 


145


Fig. 41. Cladosporium subtilissimum (CBS 113754). A–C. Macronematous conidiophores. D. Conidial chain. E. Micronematous conidiophore. F–G. Conidia. Scale bars = 10 

µm. 

Cladosporium subtilissimum K. Schub., Dugan, Crous & U. 

Braun, sp. nov. MycoBank MB504580. Figs 40–42. 

Etymology: Refers to its narrow conidiophores and conidia. 

Differt a Cladosporio cladosporioide conidiophoris et conidiis semper asperulatis ad 

verruculosis, conidiis 0–1(–2)-septatis. 

Mycelium unbranched or sparingly branched, 1–5 µm wide, septate, 

without swellings and constrictions, hyaline to subhyaline or pale 

brown, smooth to minutely verruculose, walls unthickened or almost 

so, protoplasm somewhat guttulate or granular. Conidiophores 

macronematous and micronematous, arising laterally from 

plagiotropous hyphae or terminally from ascending hyphae, erect, 

straight to slightly flexuous, filiform to cylindrical-oblong, nonnodulose, 

sometimes geniculate towards the apex, unbranched or 

once branched, branches short to somewhat longer, usually formed 

below a septum, sometimes only short, denticle-like or conical, 

25–140 × 2–4 µm, 0–4-septate, not constricted at the septa, 

subhyaline to pale brown, almost smooth, minutely verruculose 

to verruculose, sometimes irregularly rough-walled in the lower 

part, walls unthickened or slightly thickened, protoplasm guttulate 

or somewhat granular. Conidiogenous cells integrated, terminal 

or pleurogenous, sometimes also intercalary, filiform to narrowly 

cylindrical, non-nodulose, sometimes geniculate, 14–57 µm long, 

with usually sympodial clusters of pronounced conidiogenous loci 

at the apex or on a lower level, denticle-like or situated on short 

lateral prolongations, up to five loci, intercalary conidiogenous 

cells usually with a short denticle-like lateral outgrowth below a 

septum, protuberant, denticulate, somewhat truncate, 1.2–2 µm 

diam, thickened and darkened-refractive. Ramoconidia sometimes 

occurring, conidiogenous cells seceding at one of the upper septa 

of the conidiophore and behaving like conidia, filiform or cylindrical, 

20–40(–55) µm long, 1.5–4 µm wide, 0–1-septate, concolorous 

with conidiophores, not attenuated towards apex and base, base 

broadly truncate, non-cladosporioid, without any dome and raised 

rim, 2–3.5 µm wide, neither thickened nor darkened, sometimes 

slightly refractive. Conidia catenate, in branched chains, up to 12 

or even more in a chain, straight, small terminal conidia numerous, 

subglobose, narrowly obovoid, limoniform or fusiform, 4–9 × 2–3.5 

µm [av. ± SD, 6.4 (± 1.5) × 2.8 (± 0.4) µm], with up to three distal 

scars, aseptate, hila (0.5–)0.8–1 µm diam, intercalary conidia 

narrowly ellipsoid, fusiform to subcylindrical, 9–18 × 3–4(–6) µm 

[av. ± SD, 13.0 (± 2.5) × 3.8 (± 0.3) µm], 0(–1)-septate, hila 1–1.2(– 

1.8) µm diam, with up to four distal scars, secondary ramoconidia 

ellipsoid, fusiform or subcylindrical, (13–)17–32(–37) × 3–5(–6) µm 

[av. ± SD, 21.4 (± 4.4) × 4.1 (± 0.5) µm], 0–1(–2)-septate, septum 

median or somewhat in the lower half, usually not constricted at 

the septa, with up to six distal hila crowded at the apex, hila (1.2–) 

1.5–2(–2.5) µm diam, apex often somewhat laterally enlarged 

or prolonged with hila crowded there, very pale or pale brown or 

olivaceous-brown, minutely verruculose to verruculose (granulate 

under SEM), walls unthickened or only slightly thickened, often 

slightly attenuated towards apex and base, protoplasm often 

guttulate or granular, hila protuberant, denticulate, (0.5–)0.8–2(– 

2.2) µm diam, thickened and darkened-refractive; microcyclic 

conidiogenesis occasionally observed. 


after 14 d at 25 °C, grey-olivaceous to olivaceous, olivaceousgrey, 

iron-grey or olivaceous-black reverse, velvety, margin 

regular, entire edge, white or pale greenish olivaceous, glabrous 

to feathery, aerial mycelium sparse, only few areas with abundant 

146


Fig. 42. Cladosporium subtilissimum (CBS 113754). A. Overview on the organisation of spore formation. The micrograph shows a large basal secondary ramoconidium which 

has chains of secondary ramoconidia, intercalary and small terminal conidia. The conidia are formed in rows of often three cells. Note the size difference in the different cells. 

B. Conidiophore showing very pronounced scars that almost appear as branches. C. Detail of (A), illustrating the scar formation between the cells. D. Conidia during different 

stages of formation. E. Details of pronounced hila, and prominent ornamentation on secondary ramoconidia with the central dome-formed area. F. Different conidia and hila. 

Scale bars: A = 10 µm, B–D, F = 5 µm, E = 2 µm. 

mycelium, diffuse, growth regular, flat or with a raised and wrinkled 

colony centre, radially furrowed, effuse, usually without prominent 

exudates, with age several exudates formed, sporulation profuse, 

colonies consisting of two kinds of conidiophores, short and a few 

longer ones. Colonies on MEA reaching 25 mm diam after 14 d 

at 25 °C, greenish olivaceous to grey-olivaceous in the centre, 

olivaceous-grey to iron-grey reverse, velvety, margin entire edge, 

crenate or umbonate, narrow, pale greenish olivaceous, sometimes 

radially furrowed, aerial mycelium absent or sparse, growth low 

convex with distinctly wrinkled colony centre, without prominent 

exudates, abundantly sporulating. Colonies on OA attaining 25 mm 

diam after 14 d at 25 °C, dark grey-olivaceous to olivaceous due to 

profuse sporulation, iron-grey reverse, sometimes releasing some 

olivaceous-buff pigments into the agar, velvety, margin regular, 

entire edge or crenate, narrow, colourless or white, glabrous or 

feathery, aerial mycelium sparse, growth flat with slightly raised 

colony centre, prominent exudates lacking, sporulation profuse. 


Specimens examined: Slovenia, Sečovlje, isolated from hypersaline water from 

salterns (reserve pond), 2005, P. Zalar, CPC 12044 = EXF-462. U.S.A., isolated 

from bing cherry fruits, F. Dugan, CBS 113753; isolated from a grape berry, F. 

Dugan, wf 99-2-9 sci 1, CBS-H 19865, holotype, isotype HAL 2028 F, culture extype 

CBS 113754. 

Excluded strains within the subtilissimum complex: Argentina, 

isolated from Pinus ponderosa (Pinaceae), 2005, A. Greslebin, 

CPC 12484, CPC 12485. U.S.A., isolated from grape berry, F. 

Dugan, CBS 113741, CBS 113742; isolated from grape bud, F. 

Dugan, CBS 113744. 

Substrate and distribution: Plant material and hypersaline water; 

Slovenia, U.S.A. 

Notes: Cladosporium cladosporioides is morphologically comparable 

with the new species but deviates in having usually smooth 

conidiophores and conidia, with the conidia being mainly aseptate. 

C. subtilissimum is represented by three isolates of different origins 

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Fig. 43. Cladosporium tenellum (CPC 12053). Macro- and micronematous conidiophores and conidia. Scale bar = 10 µm. K. Schubert del. 

Fig. 44. Cladosporium tenellum (CPC 12053). A–C, E. Macronematous conidiophore. D. Micronematous conidiophore. F. Ramoconidium and conidia. Scale bars = 10 µm. 

148


Differt a Cladosporio cladosporioide conidiophoris et conidiis semper asperulatis, 

locis conidiogenis apicalibus, numerosis, hilis quoque numerosis, conidiophoris 

angustioribus, (1–)1.5–3.5(–4) µm latis; et a Cladosporio subtilissimo loci 

conidiogenis et hilis apicalibus, numerosis, angustioribus, saepe 1–1.5 µm latis, 

conidiis minutis numerosis, saepe globosis. 

Fig. 45. Cladosporium tenellum (CPC 12053). A. A bird’s eye view of a colony of 

C. tenellum with its very characteristic bundles of aerial hyphae. Numerous conidia 

are visible, formed on simple conidiophores. B. Hyphae that run on the agar surface 

give rise to conidiophores and numerous conidia, that are relatively rounded. C. 

Conidiophore ends are rather simple and have large scars. D. Hila on a secondary 

ramoconidium with non-ornamented area. E. Detail of the prominent ornamentation 

on a secondary ramoconidium. Scale bars: A = 20 µm, B = 10 µm, C, E = 2 µm, 

D = 5 µm. 

and substrates. Besides these strains, several additional isolates 

listed under excluded strains are morphologically indistinguishable 

from C. subtilissimum in culture, but genetically different, clustering 

in various subclades. They are indicated as Cladosporium sp. in 

the tree (Fig. 3). 

Cladosporium tenellum K. Schub., Zalar, Crous & U. Braun, sp. 


Etymology: Refers to its narrow conidiophores and conidia. 


Mycelium sparingly branched, 1–3 µm wide, septate, septa often 

not very conspicuous, not constricted at the septa, sometimes 

slightly swollen, subhyaline, smooth, walls unthickened. 

Conidiophores macronematous and micronematous, solitary, 

arising terminally or laterally from plagiotropous or ascending 

hyphae, erect or subdecumbent, almost straight to more or less 

flexuous, cylindrical, sometimes geniculate towards the apex, but 

not nodulose, sometimes with short lateral prolongations at the 

apex, unbranched to once or twice branched (angle usually 30–45° 

degree, sometimes up to 90°), branches usually below a septum, 

6–200 × (1–)2–4(–5) µm, septate, septa not very conspicuous, 

not constricted at the septa, subhyaline to pale brown, almost 

smooth to usually asperulate, walls unthickened or almost so. 

Conidiogenous cells integrated, terminal or intercalary, sometimes 

conidiophores reduced to conidiogenous cells, cylindrical, 

sometimes geniculate, non-nodulose, 6–40 µm long, proliferation 

sympodial, with several conidiogenous loci often crowded at the 

apex and sometimes also at a lower level, situated on small lateral 

shoulders, unilateral swellings or prolongations, with up to 6(–10) 

denticulate loci, forming sympodial clusters of pronounced scars, 

intercalar conidiogenous cells with short or somewhat long lateral 

outgrowths, short denticle-like or long branches with several scars 

at the apex, usually below a septum, loci protuberant, 1–1.5(–2) µm 

diam, thickened and darkened-refractive. Ramoconidia sometimes 

occurring, cylindrical, up to 32 µm long, 2.5–4 µm wide, with a 

broadly truncate, unthickened base, about 2 µm wide. Conidia 

catenate, formed in branched chains, straight, small terminal 

conidia globose, subglobose, ovoid, oval, 3–6 × 2.5–3.5 µm [av. 

± SD, 4.5 (± 1.3) × 2.8 (± 0.4) µm], aseptate, asperulate, with 0–2 

distal hila, intercalary conidia and secondary ramoconidia ellipsoidovoid, 

ellipsoid to subcylindrical, 3.5–20(–28) × (2.5–)3–5(–6) µm 

[av. ± SD, 12.4 (± 5.4) × 4.1 (± 0.7) µm], 0–1-septate, rarely with 

up to three septa, sometimes slightly constricted at the septa, 

subhyaline, pale brown to medium olivaceous-brown, asperulate 

or verruculose (muricate, granulate or colliculate under SEM), 

walls unthickened or slightly thickened, apex rounded or slightly 

to distinctly attenuated towards apex and base, often forming 

several apical hila, up to 7(–9), crowded, situated on small lateral 

outgrowths giving them a somewhat irregular appearance, hila 

protuberant, 0.5–1.5 µm diam, thickened and darkened-refractive; 

microcyclic conidiogenesis sometimes occurring. 


after 14 d at 25 ºC, smoke-grey, grey-olivaceous to olivaceous-grey, 

olivaceous-grey to iron-grey reverse, velvety to powdery, margin 

regular, entire edge, narrow, colourless to white, aerial mycelium 

absent or sparingly formed, felty, whitish, growth regular, flat, 

radially furrowed, with folded and elevated colony centre, deep into 

the agar, with age forming few to numerous prominent exudates, 

sporulation profuse, few high conidiophores formed. Colonies on 

MEA reaching 25–44 mm diam after 14 d at 25 ºC, olivaceous-grey 

to olivaceous- or iron-grey due to abundant sporulation in the colony 

centre, velvety, margin regular, entire edge, narrow, colourless, 

white to pale olivaceous-grey, aerial mycelium loose, diffuse, 

growth convex with papillate surface, radially furrowed, wrinkled, 

without prominent exudates, sporulating. Colonies on OA reaching 

23–32 mm diam after 14 d at 25 ºC, grey-olivaceous, olivaceous- 

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Fig. 46. Cladosporium variabile (CPC 12751). Macro- and micronematous conidiophores and conidia. Scale bar = 10 µm. K. Schubert del. 

grey to olivaceous due to abundant sporulation in the colony centre, 

olivaceous- or iron-grey reverse, velvety, margin regular, entire 

edge, narrow, colourless or white, aerial mycelium sparse, diffuse, 

floccose, growth flat to low convex, radially furrowed, wrinkled, 

without prominent exudates, sporulation profuse. 

Specimens examined: Israel, Eilat, isolated from hypersaline water from salterns, 

2004, N. Gunde-Cimerman, CBS 121633 = CPC 12051 = EXF-1083; Ein Bokek, 

isolated from hypersaline water of the Dead Sea, 2004, M. Ota, CBS-H 19866, 

holotype, isotype HAL 2029 F, culture ex-type CBS 121634 = CPC 12053 = EXF- 

1735. U.S.A., Seattle, University of Washington campus, isolated from Phyllactinia 

sp. (Erysiphaceae) on leaves of Corylus sp. (Corylaceae), 16 Sep. 2004, D. Glawe, 

CPC 11813. 

Substrates and distribution: Hypersaline water and plant material; 

Israel, U.S.A. 

Notes: Cladosporium subtilissimum and C. cladosporioides are 

morphologically comparable with the new species C. tenellum, but 

C. cladosporioides deviates in having usually smooth conidiophores 

and conidia with only few conidiogenous loci and conidial hila crowded 

at the apex and somewhat wider conidiophores, 3–5(–6) µm; and 

in C. subtilissimum the small terminal conidia are not globose but 

rather narrowly obovoid to limoniform, the conidiogenous loci and 

conidial hila are somewhat wider, (0.5–)0.8–2(–2.2) µm, and at the 

apices of conidiophores and conidia only few scars are formed. 

Cladosporium ramotenellum, which morphologically also 

resembles C. tenellum, possesses longer and narrower, 0–3-septate 

conidia, 2.5–35 × 2–4(–5) µm, but forms only few conidiogenous 

loci and conidial hila at the apices of conidiophores and conidia. 

Cladosporium variabile (Cooke) G.A. de Vries, Contr. Knowl. 

Genus Cladosporium: 85. 1952. Figs 46–48. 

Basionym: Heterosporium variabile Cooke, Grevillea 5(35): 123. 

1877. 

≡ Helminthosporium variabile Cooke, Fungi Brit. Exs. Ser. 1, No. 360. 

1870, nom. inval. 

= Cladosporium subnodosum Cooke, Grevillea 17(83): 67. 1889. 

150


Fig. 47. Cladosporium variabile and its teleomorph Davidiella variabile (CPC 12751). A–C. Macronematous conidiophores. D, F. Micronematous conidiophores. E, G–H. 

Conidia. I. Twisted aerial mycelium. J. Ascomata formed on nettle stem in culture. K. Surface view of ascomal wall of textura epidermoidea. L–M. Asci. N–P. Ascospores. Q. 

Ascus with a sheath. Scale bars A, D, G–J, K–N = 10 µm, J = 250 µm. 


151


Fig. 48. Cladosporium variabile (CPC 12753). A. Survey of hyphae that grow on the agar surface. Some of the fungal cells have a swollen appearance and could develop into 

a “foot cell” that gives rise to a conidiophore. B. A number of aerial hyphae obstruct the swollen, large structures on the agar surface, which give rise to conidiophores. Some 

of them appear ornamented. C. A series of conidia formed on a conidiophore (bottom of the micrograph). D. Detail of the ornamented conidia. The ornamentations are isolated 

and dispersed. Note also the ornamentation-free scar zone and the hilum of the left cell. E. Two conidia behind an aerial hypha. F. Two conidiophores forming secondary 

ramoconidia. Note the bulbous shape of the spore-forming apparatus. This micrograph is from an uncoated sample. Scale bars: A–C, F = 10 µm, D = 2 µm, E = 5 µm. 

Teleomorph: Davidiella variabile Crous, K. Schub. & U. Braun, sp. 


Davidiellae tassianae similis, sed ascosporis maioribus, (22–)26–30(–35) × (7–)7.5– 

8(–9) µm, et ascis latioribus, plus quam 18 µm. 

Ascomata pseudothecial, black, superficial, situated on a small 

stroma, globose, up to 250 µm diam, with 1–3 ostiolate necks; 

ostioles periphysate, with apical periphysoids present; wall 

consisting of 3–6 layers of dark brown textura angularis, textura 

epidermoidea in surface view. Asci fasciculate, bitunicate, 

subsessile, obovoid to broadly ellipsoid, straight to slightly curved, 

8-spored, 70–95 × 18–28 µm; with pseudoparenchymatal cells 

of the hamathecium persistent. Ascospores tri- to multiseriate, 

overlapping, hyaline, with irregular lumina, thick-walled, straight 

to slightly curved, fusoid-ellipsoidal with obtuse ends, widest near 

the middle of the apical cell, medianly 1-septate, not to slightly 

constricted at the septum, at times developing a second septum 

in each cell, several ascospores with persistent, irregular mucoid 

sheath, (22–)26–30(–35) × (7–)7.5–8(–9) µm. 

Mycelium immersed and superficial, irregularly branched, aerial 

mycelium twisted and spirally coiled, 1–3 µm wide, septate, 

sometimes with swellings or small lateral outgrowths, hyaline 

to subhyaline, smooth, walls unthickened, hyphae which give 

rise to conidiophores somewhat wider, 3–4.5 µm, subhyaline to 

pale brown, almost smooth to minutely verruculose, sometimes 

enveloped by a polysaccharide-like cover. Conidiophores usually 

macronematous, but also micronematous, arising terminally 

from ascending hyphae or laterally from plagiotropous hyphae. 

Macronematous conidiophores erect, more or less straight to 

flexuous, often distinctly geniculate-sinuous forming lateral 

shoulders or unilateral swellings, sometimes zigzag-like or 

somewhat coralloid, nodulose, swellings at first terminal, then 

becoming lateral due to sympodial proliferation, often as distinct 

lateral shoulders, unbranched, sometimes once branched, 6– 

180 × (2.5–)3–6 µm, swellings (3–)6–11 µm wide, septate, not 

constricted at the septa, pale to medium olivaceous-brown or 

brown, usually verruculose, walls somewhat thickened, about 1 µm 

thick, sometimes appearing to be two-layered. Conidiogenous cells 

integrated, terminal and intercalary, cylindrical, nodulose to nodose, 

152


with a single or two swellings per cell, swellings apart from each 

other or formed in short succession, loci confined to swellings, up to 

six per node, protuberant, 1–2 µm diam, thickened and darkenedrefractive. 

Micronematous conidiophores erect, straight to slightly 

flexuous, unbranched, usually without swellings, filiform to narrowly 

cylindrical, sometimes only as short lateral outgrowths of hyphae, 

often almost indistinguishable from hyphae, up to 50 µm long, 

1.5–2.5(–3) µm wide, longer ones pluriseptate, septa appear to be 

somewhat more darkened, with very short cells, 4–12 µm long, 

subhyaline to pale brown, smooth, walls unthickened or almost so. 

Conidiogenous cells integrated, usually terminal, rarely intercalary, 

cylindrical, non-nodulose, with a single, two or few conidiogenous 

loci at the distal end, protuberant, up to 2 µm diam, thickened 

and darkened-refractive. Conidia catenate, in branched chains, 

straight, subglobose, obovoid, oval, broadly ellipsoid to cylindrical, 

sometimes clavate, 4–26(–30) × (3.5–)5–9(–10) µm [av. ± SD, 

16.8 (± 6.9) × 6.5 (± 1.4) µm], 0–3-septate, usually not constricted 

at the septa, septa becoming sinuous with age, often appearing to 

be darkened, pale to medium or even dark brown or olivaceousbrown, 

verruculose to densely verrucose or echinulate (granulate 

under SEM), walls slightly to distinctly thickened in larger conidia, 

apex and base broadly rounded, sometimes broadly truncate or 

somewhat attenuated, apex and base often appear to be darkened 

or at least refractive, hila protuberant to somewhat sessile (within 

the outer wall ornamentation), (0.8–)1–2 µm diam, thickened and 

darkened-refractive; microcyclic conidiogenesis occurring. 


after 14 d at 25 ºC, olivaceous to olivaceous-grey or iron-grey, irongrey 

or olivaceous-grey reverse, velvety to powdery, margin regular, 

entire edge to fimbriate, almost colourless, aerial mycelium whitish 

turning olivaceous-grey, sometimes reddish, greyish rose, woollyfelty, 

growth flat with elevated colony centre, somewhat folded or 

radially furrowed, with age forming several very small but prominent 

exudates, sporulation profuse. Colonies on MEA attaining 27 mm 

diam after 14 d at 25 ºC, olivaceous-grey to iron-grey, white to pale 

olivaceous-grey in the centre due to abundant aerial mycelium, 

velvety, margin very narrow, colourless, more or less entire edge, 

radially furrowed, aerial mycelium fluffy to floccose, dense, growth 

low convex with wrinkled and folded centre, without exudates, 

sporulation profuse. Colonies on OA attaining 25 mm diam after 

14 d at 25 ºC, iron-grey or olivaceous, margin regular, entire edge, 

narrow, white, glabrous, aerial mycelium whitish, at first mainly in 

the colony centre, high, dense, floccose, growth flat, abundantly 

sporulating, no exudates. 

Specimens examined: Great Britain, Wales, Montgomeryshire, Welshpool, Forden 

Vicarage, on Spinacia oleracea (Chenopodiaceae), J.E. Vize, Cooke, Fungi Brit. Exs. 

Ser. I, No. 360, K, holotype. U.S.A., Washington, isolated from Spinacia oleracea, 1 

Jan. 2003, L. DuToit, CBS-H 19867, epitype designated here of C. variabile and D. 

variabile, cultures ex-epitype CBS 121635 = CPC 12753, CPC 12751. 

Substrate and distribution: Leaf-spotting fungus on Spinacia 

oleracea; Asia (China, India, Iraq, Pakistan), Europe (Austria, 

Belgium, Cyprus, Denmark, France, Germany, Great Britain, 

Hungary, Italy, Montenegro, Netherlands, Norway, Romania, Spain, 

Turkey), North America (U.S.A.). 

Literature: de Vries (1952: 85–88), Ellis (1971: 315), Ellis & Ellis 

(1985: 429), David (1995b; 1997: 94, 96–98), Ho et al. (1999: 

144). 

Notes: In vivo the conidia are usually longer, somewhat wider and 

more frequently septate, (6.5–)10–45(–55) × (4.5–)6–14(–17) µm, 


0–4(–5)-septate (Schubert 2005). In culture the dimensions tend to 

be smaller, which was already mentioned by de Vries (1952). 

This leaf-spotting fungus superficially resembles C. 

macrocarpum, but besides its pathogenicity to Spinacia, C. 

variabile differs from the latter species in having distinctly larger 

and more frequently septate conidia on the natural host, forming 

twisted and spirally coiled aerial mycelium in culture and in having 

lower growth rates in culture (29 mm after 14 d on PDA versus 

38 mm on average in C. macrocarpum). Furthermore, the conidial 

septa of C. variabile are often distinctly darkened, become sinuous 

with age and the apex and base of the conidia often appear to 

be distinctly darkened. A Davidiella teleomorph has not previously 

been reported for this species. 

The cladosporioides complex 

This species complex will be treated in an additional paper in 

this series, dealing with the epitypification of this common and 

widespread species, and with numerous isolates identified and 

deposited as C. cladosporioides. 

DISCUSSION 

In the present study, a multilocus genealogy supported by light 

and SEM microscopy, and cultural characteristics was used to 

redefine species borders within Cladosporium, especially within 

the C. herbarum complex. Most of the diagnostic features used 

for species delimitation on host material (Heuchert et al. 2005, 

Schubert 2005), proved to be applicable in culture. However, 

morphological features were often more pronounced in vivo than 

in vitro. For instance, conidiophore arrangement is not applicable 

to cultures, conidiophore and conidium widths were often 

narrower in culture than on the natural host, and macro- as well 

as microconidiophores were often observed in culture, but not on 

host material. All species belonging to the C. herbarum complex 

are characterised by possessing conidia which are ornamentated, 

the ornamentation ranging from minutely verruculose to verrucose, 

echinulate or spiny whereas in the C. sphaerospermum complex 

species with both smooth-walled as well as ornamented conidia 

are included (Zalar et al. 2007). The surface ornamentation varies 

based on the length of surface protuberances and in the density 

of ornamentation. Furthermore, the conidia are mainly catenate, 

formed in unbranched or branched chains. However, species 

previously referred to the genus Heterosporium, which usually 

produce solitary conidia or unbranched chains of two or three 

conidia at the most on the natural host, also belong to this species 

complex (e.g., C. iridis). In vitro these chains can become longer 

and may even be branched. The conidiophores formed in culture 

are mostly macro- but may also be micronematous, sometimes 

forming different types of conidia that vary in shape and size from 

each other. Most of the species possess nodulose conidiophores 

with the conidiogenesis confined to the usually lateral swellings. 

However, this phenetic trend is not consistently expressed in all 

of the species belonging to the C. herbarum complex. The various 

Cladosporium species within the C. herbarum complex were 

observed to have subtle differences in their phenotype which were 

visible via cryo-electron microscopy (cryoSEM), and are discussed 

below. 

Fungal colonies: CryoSEM provides the opportunity to study the 

organisation of the fungal colony at relatively low magnifications. 

Cladosporium tenellum proved to be the only fungus able to form 

153


aerial hyphal strands under the conditions studied. Cladosporium 

variabile formed abundant aerial hyphae, but in C. spinulosum 

these were sparse, and only conidiophores were observed on the 

agar surface. Three-day-old colonies of C. subinflatum formed 

numerous, long aerial hyphae, and no conidiophores could be 

discerned under the binocular. After 11 d the aerial hyphae seemed 

to have disappeared, giving rise to conidiophores. Cladosporium 

antarcticum, C. variabile and C. ramotenellum showed very large, 

swollen (> 10 μm) cells which gave rise to conidiophores. With C. 

variabile possible earlier stages of these cells were visible (Fig. 48), 

which gave rise to conidiophores. More than one conidiophore could 

be formed on such a structure (C. variabile and C. ramotenellum). 

Cladosporium herbarum has very wide hyphae on the agar surface, 

which gave rise to conidiophores as lateral branches. These wide 

hyphae were observed to anastomose, which may provide a firm 

interconnected supporting mycelium for these conidiophores. In 

C. herbaroides these wide hyphae could also be discerned, but 

conidiophore formation was less obvious. Similarily, C. tenellum 

has wide, parallel hyphae that gave rise to conidiophores. 

These observations reveal fungal structures in Cladosporium 

that have not previously been reported on, and that raise intriguing 

biological questions. For instance, why are hyphal strands observed 

in some species (C. tenellum), and not in others, and what happens 

to the aerial hyphae during incubation in some species such as C. 

subinflatum? Furthermore, these preliminary results suggest that 

CryoSEM provide additional features that can be used to distinguish 

the different species in the C. herbarum complex. 

Fine details of morphological stuctures: CryoSEM provides the 

opportunity to study fine details of the conidiophore, (ramo)conidia 

and scars. Samples can be studied at magnification up to 

× 8 000, revealing details at a refinement far above what is 

possible under the light microscope (LM) (Fig. 2). However, the LM 

micrographs provide information about the different compartments 

of ramoconidia, as well as the thickness and pigmentation of the 

cell wall of different structures. With other words, the different 

techniques are complementary, and both reveal fungal details that 

build up the picture that defines a fungal species. 

Conidiophores can vary with respect to their width and the 

length. Cladosporium ramotenellum, C. antarcticum and C. variabile 

have tapered conidiophores formed on large globoid “foot cells”. 

The conidiophore itself can be branched. Cladosporium spinulosum 

has conidiophores that rise from the agar surface, but can have 

a common point of origin. These conidiophores are not tapered, 

but parallel and slender. The conidiophores of C. bruhnei and C. 

herbaroides are rather long, and can appear as aerial hyphae. 

An important feature of the conidiophore is the location were the 

conidia are formed. Conidiophore ends can be simple and tubular, 

or rounded to more complex, several times geniculate, with several 

scars. Conidiophore ends become more elaborate over time. 

Cladosporium spinulosum and C. tenellum have nearly tubular 

conidiophore ends, with often very closely aggregated scars. The 

conidiophore ends of C. subinflatum are also near tubular with a 

hint of bulbousness. Cladosporium subtilissimum is similar, but with 

somewhat more elevated scars that look denticulate. Cladosporium 

variabile has nodulose, somewhat swollen apices with often sessile, 

almost inconspicuous scars. In the case of C. macrocarpum, these 

structures are also nodulose to nodose and somewhat bent, with 

only slightly protuberant loci. Cladosporium ramotenellum has 

tubular conidiophore ends with pronounced scars. Cladosporium 

antarcticum has very characteristic, tapered ends, and widely 

dispersed (5 μm) scars. More complex conidiophore ends are 

more irregular in shape, and have scars dispersed over a longer 

distance, such as observed in C. bruhnei, C. herbaroides, and C. 

herbarum. 

Secondary ramoconidia are usually the first conidia formed on 

a conidiophore. They are often multicellular, and have one basal 

cladosporioid hilum, and more at the apex. Few Cladosporium 

species additionally form true ramoconidia representing apical 

parts of the conidiophore which secede at a septum resulting in 

an undifferentiated non-coronate base and function as conidia. 

Ramification of conidial chains is realised through these conidia. 

They can occur in up to three stages, which results in elaborated 

spore structures. The basal secondary ramoconidium is invariably 

the largest, and cell size decreases through a series of additional 

secondary ramoconidia, intercalary conidia, and small, terminal 

conidia. The elongation of secondary ramoconidia varies among 

the different species. Cladosporium macrocarpum has broadly 

ellipsoid to cylindrical secondary ramoconidia usually with broadly 

rounded ends, like C. variabile, while C. spinulosum has secondary 

ramoconidia that can often hardly be discerned from the conidia 

that are formed at later stages. The conidia of the other species 

roughly fall between these species. The most notable structures on 

these conidia are their ornamentation, scar pattern and morphology. 

Cladosporium spinulosum forms numerous globose to subsphaerical 

spores with digitate, non-tapered surface ornamentation, which 

is unique for all the species discussed here. In his study on 

Cladosporium wall ornamentation, David (1997) recognised three 

classes of echinulate surfaces (aculeate, spinulose, digitate), and 

five classes of verrucose surfaces (muricate, granulate, colliculate, 

pustulate and pedicellate) (Fig. 2). The ornamentation particles vary 

in shape, width, height and density. The most strongly ornamented 

conidia of the species examined by SEM are formed by C. ossifragi, 

with the ornamentation both large (up to 0.5 μm wide) and high, 

and can be regarded as densely muricately ornamented. Strong 

ornamentation is also seen in C. herbaroides, which is mostly 

granulate. Cladosporium tenellum (with muricate, granulate and 

colliculate tendencies) and C. bruhnei (mostly granulate with some 

muricate projections) have relatively large ornamentation structures 

with slightly more space between the units than the other two 

species. Cladosporium antarcticum, C. ramotenellum, C. variabile 

and C. subtilissimum exhibit rather large granulate ornamentations 

that have a more irregular and variable shape. Cladosporium 

subinflatum shows the widest dispersed structures of the series, 

being muricate. In contrast, C. macrocarpum has a very neat and 

regular pattern of muricate ornamentation. The area of formation 

of new spores on conidia is invariably not ornamented, and hila all 

have the typical Cladosporium morphology with a central dome and 

a ring-like structure around it. 

Branching patterns: Spores usually show a “line of weakness” 

between them where the coronate scars form. It seems that scars at 

both sides of the line of weakness have the central dome structure, 

which appears to play a major role in the effective mechanism 

Cladosporium employs for spore dispersal, with the dome actively 

pushing the conidia apart. This mechanism is also illustrated in 

David (1997, fig. 2E). Indeed, conidia of Cladosporium are very 

easily dislodged; even snap freezing or the electrical forces inside 

the SEM often result in dislodgement of the spores in a powdery 

“wave”. It is no surprise, therefore, that Cladosporium conidia are 

to be found in most air samples. In Cladosporium, conidia are 

mostly formed in chains, with the size invariably decreasing from 

the base to the apex of the row. Upon formation each conidium is 

separated from the conidiophore, or previously formed conidium, 

and hence from its nutrients. The basal ramoconidium or secondary 

ramoconidia have the nutrients and metabolic power to produce 

154


a number of additional secondary ramoconidia that in turn could 

produce a chain of intercalary conidia, and finally, some small, 

single-celled, terminal conidia. Further research is still necessary 

to determine if specific branching patterns can be linked to different 

species. 

A surprising finding from the present study is the huge diversity 

in species and genotypes that exist in nature, be it in the indoor 

environment, on fruit surfaces, or in extreme ecological niches such 

as salterns, etc. It is clear that detailed studies would be required 

to find and characterise other species of Cladosporium and obtain 

a better understanding of their host ranges and ecology. A further 

surprise lay in the fact that several of these species are capable 

of sexual reproduction, and readily form Davidiella teleomorphs in 

culture. The Davidiella states induced here were all from homothallic 

species. Further attention now needs to be given to elucidating 

teleomorphs from other species which, as in Mycosphaerella 

(Groenewald et al. 2006, Ware et al. 2007) could be heterothallic, 

and experiencing clandestine sex. 

Despite the occurrence of many different genotypes in 

variable genes, the degree of diversity in the entire data set was 

low. For the majority of the species ITS was almost invariant, with 

only six genotypes in the entire dataset. This suggests a very 

recent evolution. The standardised index of association (I S A ) was 

high (0.3914), indicating an overabundance of clonality and / or 

inbreeding, the latter possibly matching with observed homothallism 

of Davidiella teleomorphs. Clonality was visualised with SplitsTree 

software, where star-shaped representations without any sign of 

reticulation were obtained for all genes, though at different branch 

lengths (Fig. 5). With Structure software an optimal subdivision 

was achieved at six putative groups. Some of them were distinctly 

separated, yielding a theta (θ) around 0.14, but in most cases there 

was considerable overlap in representation of motifs, with θ at 

significantly higher values. Results are difficult to interpret due to 

the small size of the data set compared to the number of predicted 

groups, and due to unknown but probably large sampling effects. 

With optimal subdivision of the 79 strains at a hypothesised value 

of K = 6 (Fig. 4), still a large degree of inter-group similarity was 

noted, as was the case at any other level of K. This was particularly 

obvious when data from the most variable genes (EF and ACT) are 

superimposed (Fig. 4). The ACT groups are further subdivided by EF 

data, but in many cases the same EF motif (indicated with arrows) 

was encountered in different (multilocus) species, for example in C. 

antarcticum, C. spinulosum, Davidiella sp., and the various clusters 

comprising Cladosporium strains which are phenotypically almost 

indistinguishable but genetically distinct from C. subtilissimum. A 

similar situation was found with the distribution of EF genotypes 

(indicated with doughnuts) in C. herbarum and C. macrocarpum. 

Nevertheless, the data set showed significant structuring, partly 

correlating with geography, e.g. the EF-determined cluster of C. 

bruhnei that contained isolates from different sources in The 

Netherlands. Differences may be over-accentuated by known 

sampling effects, particularly in C. herbarum and C. macrocarpum, 

where single-spore isolates from a single collection are included. 

Taken together the data suggest a recent, preponderantly clonal 

evolution, combined with limited natural selection at a low level of 

evolutionary pressure. As a result, many genotypes produced by 

hot spots in the genes analysed have survived, leading to nearly 

random variation in the data set. Many combinations of motifs that 

possibly could emerge have maintained in the course of time due 

to the absence of recombination. This indicates that the observed 

structure is that of populations within a single species, and 

consequently a distinction of clonal “species” could be redundant. 

This conclusion is underlined by the fact that a single source in 

a single location can be colonised by various genotypes, such 

as grapes in the U.S.A. containing three different, closely related 

genotypes. However, the phenomenon of co-inhabitation by 

different Mycosphaerella species on the same lesion of Eucalyptus 

has been described before (Crous 1998, Crous et al. 2004) and it 

is therefore not surprising that different genotypes occurring close 

together are also observed for the related genus Cladosporium. 

There is no obvious ecological difference between genotypes, and 

hence isolates seem to have equal fitness. 

However, in general we noticed a remarkable concordance 

of genetic and phenetic characters. The morphological study was 

done prior to sequencing, and nearly all morphotypes clustered 

in separate molecular entities. There are some exceptions, such 

as with C. antarcticum with striking morphology that was almost 

identical on the molecular level to Cladosporium spp. that resemble 

C. subtilissimum and would normally have been interpreted to 

be a mutant. Conversely, nearly all genetically distinguishable 

groups proved to be morphologically different, with the exception 

of members of the C. subtilissimum s. lat. complex (indicated as 

Cladosporium sp. in Fig. 3 and Table 1). The possibility remains 

that the found genetic parameters correlate with phenetic markers 

other than morphology, such as virulence, toxins or antifungal 

susceptibilities. For this reason we introduce the established 

entities here as formal species. They can be diagnosed by ACT 

sequencing or by phenetic characters provided in the key. For 

simple routine purposes, however, they can be seen and treated 

as the “C. herbarum complex”, based on their close phylogenetic 

relationships. 


Several colleagues from different countries provided material and valuable cultures 

without which this work would not have been possible. In this regard, we thank 

particularly Alina G. Greslebin (CIEFAP, Esquel, Chubut, Argentina), Frank Dugan 

and Lindsey DuToit (Washington State Univ., U.S.A.), and Annette Ramaley 

(Colorado, U.S.A.). Konstanze Schubert was financially supported by a Synthesys 

grant (No. 2559) which is gratefully acknowledged. We thank Marjan Vermaas for 

preparing the photographic plates, Rianne van Dijken for the growth studies, and 

Arien van Iperen for maintaining the cultures studied. Bert Gerrits van den Ende is 

acknowledged for assisting with the molecular data analyses. 

REFERENCES 



Arx JA von (1950). Über die Ascusform von Cladosporium herbarum (Pers.) Link. 

Sydowia 4: 320–324. 

Barr ME (1958). Life history studies of Mycosphaerella tassiana and M. typhae. 

Mycologia 50: 501–513. 

Braun U (2000). Miscellaneous notes on some micromycetes. Schlechtendalia 5: 

31–56. 



nov., the teleo-morph of Cladosporium s.str. Mycological Progress 2: 3–18. 

Braun U, Hill CF, Schubert K (2006). New species and new records of biotrophic 

micromycetes from Australia, Fiji, New Zealand and Thailand. Fungal Diversity 

22: 13–35. 




Clements FE, Shear CL (1931). The genera of fungi. New York: H.W. Wilson Co. 

Corda ACJ (1837). Icones fungorum hucusque cognitorum. Vol. 1. Praha. 




155


Crous PW, Aptroot A, Kang JC, Braun U, Wingfield MJ (2000). The genus 




Crous PW, Groenewald JZ, Groenewald M, Caldwell P, Braun U, Harrington TC 

(2006). Species of Cercospora associated with grey leaf spot of maize. Studies 

in Mycology 55: 189–197. 

Crous PW, Groenewald JZ, Mansilla JP, Hunter GC, Wingfield MJ (2004). 





1081–1101. 

David JC (1995a). Cladosporium magnusianum. Mycopathologia 129: 53–54. 

David JC (1995b). Cladosporium varibile. Mycopathologia 129: 57–58. 



Domsch KH, Gams W, Anderson TH (1980). Compendium of soil fungi. Vols 1 & 2. 

Acad. Press, London. 

Dugan FM, Roberts RG (1994). Morphological and reproductive aspects of 

Cladosporium macrocarpum and C. herbarum from bing cherry fruits. 

Mycotaxon 52: 513–522. 




from the Red Sea Coast of Egypt. Fungal Diversity 5: 43–54. 



Ellis MB (1972). Dematiaceous hyphomycetes. XI. Mycological Papers 131: 1–25. 

Ellis MB, Ellis JP (1985). Microfungi on land plants. An identification handbook. 

MacMillan, New York. 

Ellis MB, Ellis JP (1988). Microfungi on miscellaneous substrates. An identification 

handbook. Croom Helm, London, and Timber Press, Portland, Oregon. 

Ellis MB, Waller JM (1974). Mycosphaerella macrospora (conidial state: Cladosporium 

iridis). CMI Descriptions of Pathogenic Fungi and Bacteria No. 435. 

Falush D, Stephens M, Pritchard JK (2003). Inference of population structure: 

Extensions to linked loci and correlated allele frequencies. Genetics 164: 

1567–1587. 

Gams W, Verkleij GJM, Crous PW (2007). CBS Course of Mycology, 5 th ed. 

Centraalbureau voor Schimmelcultures, Utrecht, Netherlands. 

Groenewald M, Groenewald JZ, Harrington TC, Abeln ECA, Crous PW (2006). 

Mating type gene analysis in apparently asexual Cercospora species is 

suggestive of cryptic sex. Fungal Genetics and Biology 43: 813–825. 

Haubold B, Hudson RR (2000). LIAN 3.0: detecting linkage disequilibrium in 

multilocus data. Bioinformatics Applications Note 16: 847–848. 

Haubold B, Travisano M, Raibey PB, Hudson RR (1998). Detecting linkage 

disequilibrium in bacterial populations. Genetics 50: 1341–1348. 

Hawksworth DL (1979). The lichenicolous hyphomycetes. Bulletin of the British 

Museum (Natural History), Botany 6: 183–300. 

Heuchert B, Braun U (2006). On some dematiaceous lichenicolous hyphomycetes. 

Herzogia 19: 11–21. 



Ho MHM, Castañeda RF, Dugan FM, Jong SC (1999). Cladosporium and 


72: 115–157. 

Hoog GS de, Guarro J, Gené J, Figueras MJ (2000). Atlas of clinical fungi, 2nd ed. 

Centraalbureau voor Schimmelcultures, Utrecht and Universitat rovira I virgili, 

Reus. 



Huson DH, Bryant D (2006). Application of phylogenetic networks in evolutionary 

studies. Molecular Biology and Evolution 23: 254–267. 

Jaap O (1902). Abhandlungen. Schriften des Naturwissenschaftlichen Vereins 

Schleswig-Holstein 12: 346–347. 

Jolley KA, Feil EJ, Chan MS, Maiden MCJ (2001). Sequence type analysis and 

recombinational tests (START). Bioinformatics Applications Notes 17: 1230– 

1231. 

Kirk PM, Cannon PF, David JC, Stalpers JA (2001). Dictionary of the fungi, 9 th edn. 

CABI Biosciences, Egham, U.K. 

Lind J (1913). Danish fungi as represented in the herbarium of E. Rostrup. 

Copenhagen. 

Linder DH (1947). Botany of the Canadian Eastern Arctic, part II Thallophyta and 

Bryophyta. 4. Fungi. Bulletin of the National Museum of Canada 97, Biological 

Series 26: 234–297. 

Link HF (1809). Observationes in ordines plantarum naturales. Dissertatio I, 

complectens Anandrarum ordines Epiphytas, Mucedines, Gastromycos et 

Fungos. Der Gesellschaft naturforschender Freunde zu Berlin Magazin für die 

neuesten Entdeckungen in der gesammten Naturkunde 3: 3–42. 

Link HF (1816). Observationes in ordines plantarum naturales. Dissertatio II, sistens 

nuperas de Mucedinum et Gastromycorum ordinibus observationes. Der 

Gesellschaft naturforschender Freunde zu Berlin Magazin für die neuesten 

Entdeckungen in der gesammten Naturkunde 7: 25–45. 

Matsushima T (1985). Matsushima Mycological Memoirs No. 4. Kobe. 





lato V. Concerning the type species Cladosporium herbarum. Mycotaxon 42: 

307–317. 

Nylander JAA (2004). MrAic.pl. Program distributed by the author (http://www.abc. 

se/~nylander/). Evolutionary Biology Centre, Uppsala University. 

Persoon CH (1794). Neuer Versuch einer systematischen Eintheilung der 

Schwämme. Neues Magazin für die Botanik in ihrem ganzen Umfange 1: 

63–128. 

Persoon CH (1822). Mycologia europaea. Sectio prima. Erlangen. 



Pritchard JK, Stephens M, Donnelly P (2000). Inference of population structure 

using multilocus genotype data. Genetics 155: 945–959. 


Kew, Surrey, England. 

Riesen T, Sieber T (1985). Endophytic fungi in winter wheat (Triticum aestivum L.). 

Swiss Federal Institute of Technology, Zürich. 

Saccardo PA (1899). Sylloge Fungorum vol. 14 (Saccardo, P.A. & Sydow, P. eds.). 

Padova. 

Samson RA, Houbraken JAMP, Summerbell RC, Flannigan B, Miller JD (2001). 

Common and important species of Actinomycetes and fungi in indoor 

environments. In: Microorganisms in Home and Indoor Work Environments: 

Diversity, Health Impacts, Investigation and Control (Flannigan B, Samson RA, 

Miller JD, eds). Taylor & Francis, London: 287–473. 

Samson RA, Reenen-Hoekstra, ES van, Frisvad, JC, Filtenborg O (2000). 

Introduction to food- and airborne fungi, 6 th ed. Centraalbureau voor 

Schimmelcultures, Utrecht. 



Mycologia 98: 1041–1052. 

Schubert K (2005). Morphotaxonomic revision of foliicolous Cladosporium species 

(hyphomycetes). Ph.D. dissertation. Martin-Luther-University Halle-Wittenberg, 

Germany. http://sundoc.bibliothek.uni-halle.de/diss-online/05/05H208/index. 

htm. 

Schubert K, Braun U, Mułenko W (2006). Taxonomic revision of the genus 

Cladosporium s. lat. 5. Validation and description of new species. 


Shin HD, Lee HT, Im DJ (1999). Occurence of German Iris leaf spot caused by 

Cladosporium iridis in Korea. Plant Pathology Journal 15: 124–126. 

Sivanesan A (1984). The bitunicate Ascomycetes and their anamorphs. Cramer 

Verlag, Vaduz. 


ex Fr. CBS, Baarn. 

Wang CJK, Zabel RA (1990). Identification manual for fungi from utility poles in the 

eastern United States. ATCC, Rockville, MD. 

Ware SB, Verstappen ECP, Breeden J, Cavaletto JR, Goodwin SB, Waalwijk C, 

Crous PW, Kema GHJ (2007). Discovery of a functional Mycosphaerella 

teleomorph in the presumed asexual barley pathogen Septoria passerinii. 

Fungal Genetics and Biology 44: 389–397. 

Zalar P, Hoog GS de, Schroers HJ, Crous PW, Groenewald JZ & Gunde-Cimerman 

N (2007). Phylogeny and ecology of the ubiquitous saprobe Cladosporium 

sphaerospermum, with description of seven new species from hypersaline 

environments. Studies in Mycology 58: 157–183. 

156


doi:10.3114/sim.2007.58.06 


Phylogeny and ecology of the ubiquitous saprobe Cladosporium sphaerospermum, 

with descriptions of seven new species from hypersaline environments 

P. Zalar 1* , G.S. de Hoog 2,3 , H.-J. Schroers 4 , P.W. Crous 2 , J.Z. Groenewald 2 and N. Gunde-Cimerman 1 

1 

Biotechnical Faculty, Department of Biology, Večna pot 111, SI-1000 Ljubljana, Slovenia; 2 CBS Fungal Biodiversity Centre, P.O. Box 85167, 3508 AD Utrecht, The Netherlands; 

3 

Institute for Biodiversity and Ecosystem Dynamics, University of Amsterdam, Kruislaan 315, 1098 SM Amsterdam, The Netherlands; 4 Agricultural Institute of Slovenia, 

Hacquetova 17, p.p. 2553, 1001 Ljubljana, Slovenia 

*Correspondence: Polona Zalar, polona.zalar@bf.uni-lj.si 

Abstract: Saprobic Cladosporium isolates morphologically similar to C. sphaerospermum are phylogenetically analysed on the basis of DNA sequences of the ribosomal RNA 

gene cluster, including the internal transcribed spacer regions ITS1 and ITS2, the 5.8S rDNA (ITS) and the small subunit (SSU) rDNA as well as β-tubulin and actin gene introns 

and exons. Most of the C. sphaerospermum-like species show halotolerance as a recurrent feature. Cladosporium sphaerospermum, which is characterised by almost globose 

conidia, is redefined on the basis of its ex-neotype culture. Cladosporium dominicanum, C. psychrotolerans, C. velox, C. spinulosum and C. halotolerans, all with globoid conidia, 

are newly described on the basis of phylogenetic analyses and cryptic morphological and physiological characters. Cladosporium halotolerans was isolated from hypersaline 

water and bathrooms and detected once on dolphin skin. Cladosporium dominicanum and C. velox were isolated from plant material and hypersaline water. Cladosporium 

psychrotolerans, which grows well at 4 °C but not at 30 °C, and C. spinulosum, having conspicuously ornamented conidia with long digitate projections, are currently only known 

from hypersaline water. We also newly describe C. salinae from hypersaline water and C. fusiforme from hypersaline water and animal feed. Both species have ovoid to ellipsoid 

conidia and are therefore reminiscent of C. herbarum. Cladosporium langeronii (= Hormodendrum langeronii) previously described as a pathogen on human skin, is halotolerant 

but has not yet been recorded from hypersaline environments. 

Taxonomic novelties: Cladosporium dominicanum Zalar, de Hoog & Gunde-Cimerman, sp. nov., C. fusiforme Zalar, de Hoog & Gunde-Cimerman, sp. nov., C. halotolerans 

Zalar, de Hoog & Gunde-Cimerman, sp. nov., C. psychrotolerans Zalar, de Hoog & Gunde-Cimerman, sp. nov., C. salinae Zalar, de Hoog & Gunde-Cimerman, sp. nov., 

C. spinulosum Zalar, de Hoog & Gunde-Cimerman, sp. nov., C. velox Zalar, de Hoog & Gunde-Cimerman, sp. nov. 

Key words: Actin, β-tubulin, halotolerance, ITS rDNA, phylogeny, SSU rDNA, taxonomy. 

INTRODUCTION 

The halophilic and halotolerant mycobiota from hypersaline 

aqueous habitats worldwide frequently contain Cladosporium 

Link isolates (Gunde-Cimerman et al. 2000, Butinar et al. 2005). 

Initially, they were considered as airborne contaminants, but 

surprisingly, many of these Cladosporium isolates were identified 

as C. sphaerospermum Penz. because they formed globoid conidia 

(data unpublished). Cladosporium sphaerospermum, known as 

one of the most common air-borne, cosmopolitan Cladosporium 

species, was frequently isolated from indoor and outdoor air (Park 

et al. 2004), dwellings (Aihara et al. 2001), and occasionally from 

humans (Badillet et al. 1982) and plants (Pereira et al. 2002). 

Strains morphologically identified as C. sphaerospermum were 

able to grow at a very low water activity (a w 

0.816), while other 

cladosporia clearly preferred a higher, less extreme water activity 

(Hocking et al. 1994). This pronounced osmotolerance suggests 

a predilection for osmotically stressed environments although 

C. sphaerospermum is reported from a wide range of habitats 

including osmotically non-stressed niches. 

We therefore hypothesised that C. sphaerospermum 

represents a complex of species having either narrow or wide 

ecological amplitudes. The molecular diversity of strains identified 

as C. sphaerospermum has not yet been determined and isolates 

from humans have not yet been critically compared with those from 

environmental samples. Therefore, a taxonomic study was initiated 

with the aim to define phylogenetically and morphologically distinct 

entities and to describe their in vitro osmotolerance and their natural 

ecological preferences. 

MATERIALS AND METHODS 

Sampling 

Samples of hypersaline water were collected from salterns located 

at different sites of the Mediterranean basin (Slovenia, Bosnia and 

Herzegovina, Spain), different coastal areas along the Atlantic 

Ocean (Monte Cristy, Dominican Republic; Swakopmund, Namibia), 

the Red Sea (Eilat, Israel), the Dead Sea (Ein Gedi, Israel), and 

the salt Lake Enriquillio (Dominican Republic). Samples from the 

Sečovlje salterns (Slovenia) were collected once per month in 1999. 

Samples from the Santa Pola salterns and Ebre delta river saltern 

(Spain) were taken twice (July and November) in 2000. A saltern in 

Namibia and one in the Dominican Republic were sampled twice 

(August and October) in 2002. Various salinities, ranging from 15 to 

32 % NaCl were encountered in these ponds. 

Isolation and maintenance of fungi 

Strains were isolated from salterns using filtration of hypersaline 

water through membrane filters (pore diam 0.45 μm), followed by 

incubation of the membrane filters on different culture media with 

lowered water activity (Gunde-Cimerman et al. 2000). Only colonies 

of different morphology on one particular selective medium per 

sample were analysed further. Strains were carefully selected from 

different evaporation ponds, collected at different times, in order to 

avoid sampling of identical clones. Subcultures were maintained 

at the Culture Collection of Extremophilic Fungi (EXF, Biotechnical 

Faculty, Ljubljana, Slovenia), while a selection was deposited at 

the Centraalbureau voor Schimmelcultures (CBS, Utrecht, The 

Netherlands) and the Culture Collection of the National Institute 

of Chemistry (MZKI, Ljubljana, Slovenia). Reference strains were 

obtained from CBS, and were selected either on the basis of the 

strain history, name, or on the basis of their ITS rDNA sequence. 

Strains were maintained on oatmeal agar (OA; diluted OA, Difco: 

15 g of Difco 255210 OA medium, 12 g of agar, dissolved in 1 L 

of distilled water) with or without 5 % additional NaCl. They were 

preserved in liquid nitrogen or by lyophilisation. Strains studied are 

listed in Table 1. 

157

Zalar et al. 

Table 1. List of Cladosporium strains, with their current and original names, geography, GenBank accession numbers and references to earlier published sequences. 

Strain Nr. a Source Geography GenBank accession Nr. b 

ITS rDNA / 18S rDNA actin β-tubulin 


CBS 177.71 Thuja tincture The Netherlands, Amsterdam DQ780399 / DQ780938 EF101354 EF101451 

CBS 812.71 Polygonatum odoratum, leaf Czech Republic, Lisen DQ780401 / – – – 

Cladosporium cladosporioides 

CBS 170.54 NT Arundo, leaf U.K., England, Kew AY213640 / DQ780940 EF101352 EF101453 

EXF-321 Hypersaline water Slovenia, Sečovlje saltern DQ780408 / – – – 

EXF-780 DQ780409 / – – – 

EXF-946 Hypersaline water Bosnia and Herzegovina, DQ780410 / – – – 

Ston saltern 

Cladosporium dominicanum 

CPC 11683 Citrus fruit (orange) Iran DQ780357 / – EF101369 EF101419 

EXF-696 Hypersaline water Dominican Republic, saltern DQ780358 / – EF101367 EF101420 

EXF-718 Hypersaline water Dominican Republic, salt lake DQ780356 / – EF101370 EF101418 

Enriquilio 

EXF-720 Hypersaline water Dominican Republic, saltern DQ780355 / – – EF101417 

EXF-727 Hypersaline water Dominican Republic, saltern DQ780354 / – – EF101416 

EXF-732 T; CBS 119415 Hypersaline water Dominican Republic, salt lake DQ780353 / – EF101368 EF101415 

Enriquilio 

Cladosporium fusiforme 

CBS 452.71 Chicken food Canada DQ780390 / – EF101371 EF101447 

EXF-397 Hypersaline water Slovenia, Sečovlje saltern DQ780389 / – EF101373 EF101445 

EXF-449 T; CBS 119414 Hypersaline water Slovenia, Sečovlje saltern DQ780388 / DQ780935 EF101372 EF101446 


ATCC 66670, as Davidiella tassiana CCA-treated Douglas-fir pole U.S.A., New York, Geneva AY361959 2 & DQ780400 / DQ780939 AY752193 11 EF101452 

Cladosporium halotolerans 

ATCC 26362 Liver and intestine of diseased frog U.S.A., New Jersey AY361982 2 / – – – 

ATCC 64726 Peanut cell suspension tissue culture U.S.A., Georgia AY361968 2 / – – – 


Stem of Hypericum perforatum Switzerland, Glarus, 

DQ780369 / – EF101402 EF101432 

identified as Mycosphaerella hyperici Mühlehorn 

CBS 191.54 Laboratory air Great Britain – / – – – 

CBS 573.78 Aureobasidium caulivorum Russia, Moscow region – / – – – 

CBS 626.82 – Sweden, Stockholm – / – – – 

dH 12862; EXF-2533 Culture contaminant Brazil DQ780371 / – EF101400 EF101422 

dH 12941; EXF-2534 Culture contaminant Turkey – / – EF101421 

dH 12991; EXF-2535 Brain Turkey DQ780372 / – EF101423 

dH 13911; EXF-2422 Ice Arctics DQ780370 / – EF101401 EF101430 

EXF-228; MZKI B-840 Hypersaline water Slovenia, Sečovlje saltern DQ780365 / DQ780930 EF101393 EF101425 


EXF-564 Hypersaline water Namibia, saltern DQ780363 / – EF101395 EF101433 

EXF-565 Hypersaline water Namibia, saltern – / – – – 



EXF-572 T; CBS 119416 Hypersaline water Namibia, saltern DQ780364 / – EF101397 EF101424 

EXF-646 Hypersaline water Spain, Santa Pola saltern DQ780366 / – EF101398 EF101428 

EXF-698 Hypersaline water Dominican Republic, saltern – / – – – 

EXF-703 Hypersaline water Dominican Republic, salt lake DQ780367 / – EF101392 EF101426 

Enriquilio 

EXF-944 Hypersaline water Bosnia and Herzegovina, – / – – – 

Ston saltern 

EXF-972 Bathroom Slovenia – / – – – 

EXF-977 Bathroom Slovenia DQ780362 / – EF101396 EF101431 

158

Cladosporium sphaerospermum species complex 



EXF-1072 Hypersaline water Israel, Dead Sea DQ780373 / – EF101399 EF101428 

EXF-2372 Hypersaline water Slovenia, Sečovlje saltern – / – – – 

UAMH 7686 

Indoor air ex RCS strip, from Apis 

mellifera overwintering facility 

U.S.A., Alta, Clyde Corner AY625063 5 / – – – 

– rDNA from bottlenose dolphin skin U.S.A., Texas AF035674 6 / – – – 

infected with Loboa loboi 

– Microcolony, on rock Turkey, Antalya AJ971409 7 / – – – 

– Microcolony, on rock Turkey, Antalya AJ971408 7 / – – – 

– Tomato leaves – L25433 8 / – – – 

Cladosporium langeronii 

CBS 189.54 NT Mycosis Brazil DQ780379 / DQ780932 EF101357 EF101435 

CBS 601.84 Picea abies, wood Germany, Göttingen DQ780382 / – EF101360 EF101438 

CBS 101880 Moist aluminium school window frame Belgium, Lichtervoorde DQ780380 / – EF101359 EF101440 

CBS 109868 Mortar of Muro Farnesiano Italy, Parma DQ780377 / – EF101362 EF101434 

dH 11736 Biomat in a lake Antarctics DQ780381 / – EF101363 EF101436 

dH 12459 Orig. face lesion Brazil DQ780378 / – EF101358 EF101439 

dH 13833 Ice Arctics DQ780383 / – EF101361 EF101437 

– Nasal mucus – AF455525 4 / – – – 

– Nasal mucus – AY345352 4 / – – – 

– Mycorrhizal roots – DQ068982 9 / – – – 

Cladosporium oxysporum 

ATCC 66669 Creosote-treated southern pine pole U.S.A., New York, 

AF393689 10 / DQ780395 AY752192 11 EF101454 

Binghamton 

ATCC 76499 Decayed leaf, Lespedeza bicolor – AF393720 – – 

CBS 125.80 Cirsium vulgare, seedcoat The Netherlands AJ300332 12 / DQ780941 EF101351 EF101455 

EXF-697 Hypersaline water Dominican Republic, salt lake DQ780392 / – – – 

Enriquilio 

EXF-699 Hypersaline water Dominican Republic, saltern DQ780394 / – – – 



Cladosporium psychrotolerans 

EXF-326 Hypersaline water Slovenia, Sečovlje saltern DQ780387 / DQ780934 – EF101444 

EXF-332 Hypersaline water Slovenia, Sečovlje saltern DQ780385 / DQ780933 EF101364 EF101441 

EXF-391 T; CBS 119412 Hypersaline water Slovenia, Sečovlje saltern DQ780386 / – EF101365 EF101442 

EXF-714 Hypersaline water Dominican Republic DQ780384 / – EF101366 EF101443 


EXF-454 T; CPC 12043 Hypersaline water Slovenia, Sečovlje saltern DQ780403 / – – – 

Cladosporium salinae 

EXF-322 Hypersaline water Slovenia, Sečovlje DQ780375 / – EF101391 EF101403 

EXF-335 T; CBS 119413 Hypersaline water Slovenia, Sečovlje DQ780374 / DQ780931 EF101390 EF101405 

EXF-604 Hypersaline water Spain, Santa Pola DQ780376 / – EF101389 EF101404 

Cladosporium sp. 

CBS 300.96 Soil along coral reef coast Papua New Guinea, Madang, DQ780352 / – EF101385 – 

Jais Aben 

EXF-595 Hypersaline water Spain, Santa Pola saltern DQ780402 / – – – 

Cladosporium sphaerospermum 

ATCC 12092 Soil Canada AY361988 2 / – – – 

ATCC 200384 Compost biofilter The Netherlands AY361991 2 / – – – 

CBS 109.14; ATCC 36950 Carya illinoensis leaf scale U.S.A. DQ780350 / – EF101384 EF101410 

CBS 122.47; IFO 6377; IMI 49640; Decaying stem of Begonia sp., The Netherlands, Aalsmeer AJ244228 1 / – – – 

VKM F-772; ATCC 11292 

with Thielaviopsis basicola 

CBS 188.54; ATCC 11290; IMI de Vries (Engelhardt strain) – AY361990 2 & AY251077 3 / – – – 

049638 

CBS 190.54; ATCC 11293; IFO 6380; – – AY361992 2 / – – – 

IMI 49641 

CBS 192.54; ATCC 11288; IMI 49636 Nail of man – AY361989 2 / – – – 


159

Zalar et al. 



CBS 193.54 NT; ATCC 11289; IMI 

49637 


Human nails – DQ780343 & AY361958 2 / DQ780925 EF101380 EF101406 

CBS 122.63 Plywood of Betula sp. Finland, Helsinki – / – – – 

CBS 102045; EXF-2524; MZKI B- Hypersaline water 

Spain, Barcelona, Salines de DQ780351 / – EF101378 EF101411 

1066 

la Trinitat 

CBS 114065 Outdoor air Germany, Stuttgart – / – – – 

CPC 10944 Gardening peat substrate Russia, Kaliningrad DQ780350 / – – – 

EXF-131; MZKI B-1005 Hypersaline water Slovenia, Sečovlje saltern AJ238670 1 / – – – 







EXF-464 Hypersaline water Slovenia, Sečovlje saltern – / DQ780927 – – 


EXF-598 Hypersaline water Spain, Santa Pola – / – EF101377 – 

EXF-644 Hypersaline water Spain, Santa Pola – / – – – 



EXF-715 Hypersaline water Dominican Republic, saltern – / – – – 



EXF-781; MZKI B-899 Hypersaline water Slovenia, Sečovlje – / – – – 

EXF-788 Hypersaline water Slovenia, Sečovlje – / – – – 


EXF-965 Bathroom Slovenia – / – – – 

EXF-1069 Hypersaline water Israel, Eilat saltern – / – EF101376 – 

EXF-1061 Hypersaline water Israel, Dead Sea DQ780346 / – EF101379 EF101408 

EXF-1726 Hypersaline water Israel, Dead Sea – / – – – 

EXF-1732 Hypersaline water Israel, Eilat saltern – / DQ780928 – – 

– Bryozoa sp. – AJ557744 / – – – 

– Nasal mucus – AF455481 4 / – – – 



EXF-334 T Hypersaline water Slovenia, Sečovlje saltern DQ780406 / – EF101355 EF101450 

EXF-382 Hypersaline water Slovenia, Sečovlje saltern DQ780407 / DQ780936 EF101356 EF101449 

Cladosporium subinflatum 

EXF-343 T; CPC 12041 Hypersaline water Slovenia, Sečovlje saltern DQ780405 / – EF101353 EF101448 

Cladosporium tenuissimum 

ATCC 38027 Soil New Caledonia AF393724 / – – – 

EXF-324 Hypersaline water Slovenia, Sečovlje saltern – / DQ780926 – – 



EXF-563 Hypersaline water Namibia, saltern DQ780398 / – – – 

Cladosporium velox 

CBS 119417 T; CPC 11224 Bamboo sp. India, Charidij DQ780361 / DQ780937 EF101388 EF101456 

EXF-466 Hypersaline water Slovenia, Sečovlje saltern DQ780359 / – EF101386 – 

EXF-471 Hypersaline water Slovenia, Sečovlje saltern DQ780360 / – EF101387 – 

160


Cultivation and microscopy 

For growth rate determination and phenetic description of colonies, 

strains were point inoculated on potato-dextrose agar (PDA, Difco), 

OA and Blakeslee malt extract agar (MEA, Samson et al. 2002) 

and incubated at 25 °C for 14 d in darkness. Surface colours were 

rated using the colour charts of Kornerup & Wanscher (1967). For 

studies of microscopic morphology, strains were grown on synthetic 

nutrient agar (SNA, Gams et al. 2007) in slide cultures. SNA blocks 

of approximately 1 × 1 cm were cut out aseptically, placed upon 

sterile microscope slides, and inoculated at the upper four edges 

by means of a conidial suspension (Pitt 1979). Inoculated agar 

blocks were covered with sterile cover slips and incubated in moist 

chambers for 7 d at 25 °C in darkness. The structure and branching 

pattern of conidiophores were observed at magnifications × 100, 

× 200 and × 400 in intact slide cultures under the microscope 

without removing the cover slips from the agar blocks. For higher 

magnifications (× 400, × 1 000) cover slips were carefully removed 

and mounted in lactic acid with aniline blue. 

Morphological parameters 

Morphological terms follow David (1997), Kirk et al. (2001) and 

Schubert et al. (2007 – this volume). Conidiophores in Cladosporium 

are usually ascending and sometimes poorly differentiated. Though 

the initiation point of conidiophore stipes could sometimes be 

determined only approximately, their lengths were in some cases 

useful for distinguishing morphologically similar species when 

observed in slide cultures. The branching patterns can be rotationally 

symmetric or unilateral. Characters of conidial scars were studied 

by light and scanning electron microscopy (SEM). Conidial chains 

show different branching patterns, determined by the numbers of 

conidia in unbranched parts, the nature of ramoconidia as well as 

their distribution in conidial chains. Measurements are given as (i) 

n 1 

–n 2 

or (ii) (n 1 

–)n 3 

–n 4 

(–n 2 

), with n 1 

= minimum value observed; n 2 

= maximum value observed; n 3 

/n 4 

= first/third quartile. For conidia 

and ramoconidia also average values and standard deviations are 

listed. The values provided are based on at least 25 measurements 

for the conidiophores of each strain, and at least 50 measurements 

for conidia. 

Ecophysiology 

To determine the degree of halotolerance, strains were pointinoculated 

on MEA without and with additional NaCl at concentrations 

of 5, 10, 17 and 20 % NaCl (w/v) and incubated at 25 °C for 14 d. 

To determine cardinal temperature requirements for growth, plates 

were incubated at 4, 10, 25, 30 and 37 °C, and colony diameters 

measured after 14 d of incubation. 

DNA extraction, sequencing and analysis 

For DNA isolation strains were grown on MEA for 7 d. DNA was 

extracted according to Gerrits van den Ende & de Hoog (1999) 

by mechanical lysis of approx. 1 cm 2 of mycelium. A fragment of 

the rDNA including the Internal Transcribed Spacer region 1, 5.8S 

rDNA and the ITS 2 (ITS) was amplified using the primers V9G 

(de Hoog & Gerrits van den Ende 1998) and LS266 (Masclaux 

et al. 1995). Sequence reactions were done using primers ITS1 

and ITS4 (White et al. 1990). For amplification and sequencing 

of the partial actin gene, primers ACT-512F and ACT-783R were 

applied according to Carbone & Kohn (1999). For amplification and 

sequencing of the β-tubulin gene primers T1 and T22 were used 

according to O’Donnell & Cigelnik (1997). A BigDye terminator 

cycle sequencing kit (Applied Biosystems, Foster City, CA, U.S.A.) 

was used in sequence reactions. Sequences were obtained with 

an ABI Prism 3700 DNA Analyzer (Applied Biosystems). They 

were assembled and edited using SeqMan v. 3.61 (DNAStar, 

Inc., Madison, U.S.A.). Sequences downloaded from GenBank 

are indicated in the trees by their GenBank accession numbers; 

newly generated sequences are indicated by strain numbers 

(see also Table 1). Sequences were automatically aligned using 

ClustalX v. 1.81 (Jeanmougin et al. 1998). The alignments were 

adjusted manually using MEGA3 (Kumar et al. 2004). Phylogenetic 

relationships of the taxa were estimated from aligned sequences 

by the maximum parsimony criterion as implemented in PAUP v. 

4.0b10 (Swofford 2003). Data sets of the SSU rDNA, ITS rDNA 

and the β-tubulin and actin genes are analysed separately. Species 

of Cladosporium s. str. were compared with various taxa of the 

Mycosphaerellaceae using SSU rDNA sequences and Fusicladium 

effusum G. Winter (Venturiaceae) as outgroup. The other data sets 

focus on Cladosporium s. str., using Cladosporium salinae Zalar, 

de Hoog & Gunde-Cimerman as an outgroup, because this species 

was most deviant within Cladosporium in the SSU rDNA analysis 

(see below). Heuristic searches were performed on all characters, 

which were unordered and equally weighted. Gaps were treated 

as missing characters. Starting tree(s) were obtained via stepwise, 

random, 100 times repeated sequence addition. Other parameters 

included a “MaxTrees” setting to 9 000, the tree-bisectionreconnection 

as branch-swapping algorithm, and the “MulTrees” 

option set to active. Branch robustness was tested in the parsimony 

analysis by 10 000 search replications, each on bootstrapped data 

sets using a fast step-wise addition bootstrap analysis. Bootstrap 

values larger than 60 are noted near their respective branches. 

Newly generated sequences were deposited in GenBank (www. 

ncbi.nlm.nih.gov); their accession numbers are listed in Table 1. 

Alignments and trees were deposited in TreeBASE (www.treebase. 

org). 

Table 1. (Page 158–160). 

a 

Abbreviations used: ATCC = American Type Culture Collection, Virginia, U.S.A.; CBS = Centraalbureau voor Schimmelcultures, Utrecht, The Netherlands; CPC = Culture 

Collection of Pedro Crous, housed at CBS, Utrecht, The Netherlands; dH = de Hoog Culture Collection, housed at CBS, Utrecht, The Netherlands; EXF = Culture Collection 

of Extremophilic Fungi, Ljubljana, Slovenia; IFO = Institute for Fermentation, Culture Collection of Microorganizms, Osaka, Japan; IMI = The International Mycological 

Institute, Egham, Surrey, U.K.; MZKI = Microbiological Culture Collection of the National Institute of Chemistry, Ljubljana, Slovenia; UAMH = University of Alberta Microfungus 

Collection, Alberta, Canada; VKM = All-Russian Collection of Microorganisms, Russian Academy of Sciences, Institute of Biochemistry and Physiology of Microorganisms, 

Pushchino, Russia; NT = ex-neotype strain; T = ex-type strain. 

b 

Reference: 1 de Hoog et al. 1999; 2 Park et al. 2004; 3 Braun et al. 2003; 4 Buzina et al. 2003; 5 Meklin et al. 2004; 6 Haubold et al. 1998; 7 Sert & Sterflinger, unpubl.; 8 Curtis et 

al. 1994; 9 Menkis et al. 2005; 10 Managbanag et al. unpubl.; 11 Crous et al. 2004; 12 Wirsel et al. 2002. All others are newly reported here. 


161

Zalar et al. 

AY251097 Cladosporium uredinicola 

Cladosporium oxysporum CBS 125.80 

Cladosporium herbarum ATCC 66670 

Cladosporium bruhnei CBS 177.71 

Cladosporium spinulosum EXF-382 

Cladosporium psychrotolerans EXF-326 

Cladosporium psychrotolerans EXF-332 

Cladosporium langeronii CBS 189.54 T 

Cladosporium tenuissimum EXF-324 

Cladosporium cladosporioides CBS 170.54 NT 

AY251092 Cladosporium colocasiae 

AY251096 Cladosporium herbarum 

98 

Cladosporium halotolerans CBS 119416 T 

Cladosporium halotolerans EXF-228 

Cladosporium dominicanum CBS 119415 T 

Cladosporium velox CBS 119417 T 

66 

Cladosporium sphaerospermum EXF-464 

Cladosporium fusiforme CBS 119414 T 

Cladosporium sphaerospermum CBS 193.54 NT 

Cladosporium salinae CBS 119413 T 

AY251107 Pseudocercospora protearum 

80 AY251106 Pseudocercospora angolensis 

64 

AY251105 Cercospora cruenta 

AY251104 Cercospora zebrina 

AY251109 Passalora fulva 

79 

AY251108 Passalora dodonaeae 

AY251117 Mycosphaerella graminicola 

AY251110 Ramulispora sorghi 

100 

AY251114 Mycosphaerella latebrosa 

AY251102 Batcheloromyces proteae 

AY251101 Mycosphaerella lateralis 

76 AY251120 Teratosphaeria nubilosa 

AY251118 Catenulostroma macowanii 

AY251121 Devriesia staurophorum 

97 U42474 Dothidea insculpta 

93 AY016343 Dothidea ribesia 

AY016342 Discosphaerina fagi 

AY251126 Fusicladium effusum 

AY251122 Fusicladium amoenum Venturiaceae 

Dothioraceae & Dothideaceae 

10 steps 

Fig. 1. One of 30 equally most parsimonious and equally looking phylogenetic trees based on a heuristic tree search using aligned small subunit ribosomal DNA sequences. 

The tree was randomly selected. Support based on 10 000 replicates of a fast step-wise addition bootstrap analysis is indicated near the branches. Species of Cladosporium s. 

str., including the seven newly described species, form a strongly supported monophyletic group among other taxa of the Mycosphaerellaceae (Dothideomycetes) (CI = 0.631, 

RI = 0.895, PIC = 50). 




Table 2. Statistical parameters describing phylogenetic analyses performed on sequence alignments of four different loci. 

Parameter SSU rDNA ITS rDNA 1 β-tubulin 2 Actin 3 

Number of alignment positions 1031 498 654 210 

Number of parsimony informative characters (PIC) 50 68 220 103 

Length of tree / number of steps 103 102 714 338 

Consistency Index (CI) 0.631 0.804 0.538 0.586 

Retention Index (RI) 0.895 0.975 0.883 0.885 

Rescaled Consistency Index (RC) 0.565 0.784 0.475 0.518 

Homoplasy index (HI) 0.369 0.196 0.462 0.414 

Number of equally parsimonious trees retained 30 600 90 32 

1 

Including the internal transcribed spacer region 1 and 2 and the 5.8S rDNA. 

2 

Including partial sequences of 4 exons and complete sequences of 3 introns. 

3 

Including partial sequences of 3 exons and 2 introns. 

162


69 

EXF-322 


EXF-604 

10 steps 

100 

85 

C. salinae 

62 

64 

79 

98 

EXF-780 

EXF-321 

EXF-946 


ATCC 76499 

ATCC 66669 

EXF-699 

EXF-710 

EXF-697 

EXF-711 


EXF-563 

EXF-452 

EXF-371 

Moricca et al. 

ATCC 38027 

EXF-714 

EXF-332 

EXF-391 

EXF-326 

87 CBS 109868 

dH 13833 


dH 11736 



dH 12459 

100 EXF-449 


EXF-397 

EXF-466 


EXF-471 

EXF-739 



EXF-455 

EXF-738 

EXF-962 

EXF-1061 


EXF-458 

CBS 300.96 Cladosporium sp. 

EXF-732 

80 

90 

62 

100 

CPC 11683 

EXF-718 

EXF-720 

EXF-727 

72 

63 

79 

99 

EXF-977 

EXF-1072 

dH 12991 

dH 12862 

dH 13911 

EXF-564 C. halotolerans 

EXF-380 

EXF-703 

EXF-646 

EXF-228 

EXF-572 

CBS 280.49 “Mycosphaerella” hyperici 

CBS 177.71 C. bruhnei 


EXF-595 Cladosporium sp. 

EXF-454 C. ramotenellum 

ATCC 66670 Davidiella tassiana 

EXF-333 

EXF-382 

EXF-334 

EXF-343 C. subinflatum 

C. spinulosum 

C. cladosporioides 

C. oxysporum 

C. tenuissimum 

C. sphaerospermum 

C. dominicanum 

C. psychrotolerans 

C. langeronii 

C. fusiforme 

C. velox 

Fig. 2. One of 600 equally most parsimonious and equally looking phylogenetic trees based on a heuristic tree search using aligned sequences of the internal transcribed 

spacer regions 1 and 2 and the 5.8S rDNA. The tree was randomly selected. Support based on 10 000 replicates of a fast step-wise addition bootstrap analysis is indicated near 

the branches. Trees were rooted with the strains of Cladosporium salinae. Most monophyletic species clades received high, but some deeper branches moderate, bootstrap 

support (CI = 0.804, RI = 0.975, PIC = 68). 

RESULTS 

Descriptive statistical parameters of phylogenetic analyses and 

calculated tree scores for each analysed sequence locus are 

summarised in Table 2. Mainly reference material such as extype 

or ex-neotype strains was analysed on the level of SSU 

rDNA sequences. Downloaded and newly generated SSU rDNA 

sequences of members of Cladosporium s. str. were compared 

with related taxa of the Mycosphaerellaceae, Dothioraceae and 

Dothideaceae. The somewhat more distantly related Fusicladium 

effusum (Venturiaceae) (Braun et al. 2003: Fig. 2) was selected 

as outgroup. Anungitopsis amoena R.F. Castañeda & Dugan (now 

placed in Fusicladium Bonord., see Crous et al. 2007b), also a 

member of the Venturiaceae, was included in the analyses. All taxa 

included in the SSU rDNA analysis belong to the Dothideomycetes 


(Schoch et al. 2006), within which the ingroup is represented by 

the orders Capnodiales (Davidiellaceae, Mycosphaerellaceae, 

Teratosphaeriaceae) and Dothideales (Dothioraceae, Dothideaceae) 

(see also Schoch et al. 2006). The genus Cladosporium, of which 

some species are linked to Davidiella Crous & U. Braun teleomorphs 

(Braun et al. 2003), forms a statistically strongly supported 

monophyletic group (Davidiellaceae). It also accommodates species 

newly described in this paper, namely, C. halotolerans Zalar, de 

Hoog & Gunde-Cimerman, C. fusiforme Zalar, de Hoog & Gunde- 

Cimerman, C. dominicanum Zalar, de Hoog & Gunde-Cimerman, C. 

salinae, C. psychrotolerans Zalar, de Hoog & Gunde-Cimerman, C. 

velox Zalar, de Hoog & Gunde-Cimerman and C. spinulosum Zalar, 

de Hoog & Gunde-Cimerman (Fig. 1). A sister group relationship 

of Cladosporium s. str. with a clade of taxa characterised, 

among others, by Mycosphaerella Johanson teleomorphs, 

containing various anamorphic genera such as Septoria Sacc., 

163

Zalar et al. 

97 EXF-322 


EXF-604 

10 steps 

89 

C. salinae 

96 

61 

87 

93 

91 

98 



dH 12459 



100 

dH 13833 

dH 11736 

EXF-332 

EXF-326 

EXF-391 

EXF-714 

100 CBS 177.71 C. bruhnei 

ATCC 66670 Davidiella tassiana 

CBS 170.54 C. cladosporioides 

EXF-343 C. subinflatum 

EXF-382 

C. spinulosum 

EXF-334 

ATCC 66669 


CBS 119417 C. velox 

C. oxysporum 

EXF-572 

EXF-564 

EXF-1072 

EXF-380 

C. halotolerans 

EXF-703 

EXF-228 

EXF-646 

EXF-977 


98 

dH 12862 

100 

63 

84 

100 

95 93 100 

100 

dH 12941 

dH 13911 

dH 12991 

EXF-727 

90 8862 

100 

99 

EXF-458 

98 

EXF-455 


EXF-1061 


EXF-738 


EXF-739 

EXF-962 

EXF-397 

EXF-449 


78 

99 

EXF-720 

EXF-732 

EXF-718 

CPC 11683 

EXF-696 

C. halotolerans 


C. fusiforme 

C. langeronii 



Fig. 3. One of 90 equally most parsimonious and equally looking phylogenetic trees based on a heuristic tree search using aligned exons and introns of a part of the β-tubulin 

gene. The tree was randomly selected. Support based on 10 000 replicates of a fast step-wise addition bootstrap analysis is indicated near the branches. Trees were rooted with 

the strains of Cladosporium salinae. Most monophyletic species clades received high, but deeper branches weak or no, bootstrap support (CI = 0.538, RI = 0.883, PIC = 220). 

Ramularia Unger, Cercospora Fresen., Pseudocercospora Speg., 

“Trimmatostroma” Corda (now Catenulostroma Crous & U. Braun) 

(see Crous et al. 2004, 2007a – this volume) and the somewhat 

cladosporium-like genus Devriesia Seifert & N.L. Nick. (Seifert et al. 

2004), was statistically only moderately supported (Fig. 1), whereas 

in an analogous analysis by Braun et al. (2003: Fig. 2) it was highly 

supported. These data also support the conclusion by Braun et al. 

(2003) and Crous et al. (2006) that Cladosporium is not a member 

of the distantly related Herpotrichiellaceae (Chaetothyriomycetes), 

which is also rich in cladosporium-like taxa (Crous et al. 2006). 

None of the fungi isolated from hypersaline environments belonged 

to the Herpotrichiellaceae. The SSU rDNA sequences do not 

resolve a phylogenetic structure within Cladosporium s. str. Only 

a moderately supported clade comprising C. halotolerans, C. 

dominicanum, C. velox, C. sphaerospermum and C. fusiforme is 

somewhat distinguished from a statistically unsupported clade with 

C. herbarum (Pers. : Fr.) Link, C. cladosporioides (Fresen.) G.A. 

de Vries, C. oxysporum Berk. & Broome, C. spinulosum, and C. 

psychrotolerans, etc. Because C. salinae appeared most distinct 

within the genus Cladosporium in analyses of the SSU rDNA (Fig. 

1), it was used as outgroup in analyses of the ITS rDNA and the 

β-tubulin and actin genes. 

Analyses of the more variable ITS rDNA and partial β-tubulin 

and actin gene introns and exons supported the species clades of 

C. halotolerans, C. dominicanum, C. sphaerospermum, C. fusiforme 

and C. velox (Figs 2–4), of which C. velox was distinguished in 

the β-tubulin tree by a particular long terminal branch of the only 

sequenced strain (Fig. 3). Cladosporium salinae also clustered as a 

well-supported species clade in preliminary analyses using various 

Mycosphaerella species as outgroup (not shown). All strains of C. 

langeronii (Fonseca, Leão & Nogueira) Vuill. are particularly well 

distinguishable from other Cladosporium species by strikingly slow- 

164


EXF-322 


EXF-604 

60 

100 

C. salinae 




dH 12459 

dH 13833 

96 CBS 109868 

dH 11736 

62 EXF-332 

EXF-391 

77 EXF-714 

CBS 177.71 C. bruhnei 

66 89 ATCC 66670 Davidiella tassiana 

67 EXF-343 C. subinflatum 

85 


87 EXF-696 

EXF-718 

100 

EXF-732 


CPC 11683 

EXF-598 

100 


EXF-458 

EXF-1069 

EXF-455 

72 

EXF-1061 

77EXF-739 

76 

EXF-962 


81 EXF-738 



CBS 300.96 Cladosporium sp. 

EXF-449 

100 90 

EXF-397 C. fusiforme 


100 

EXF-466 

EXF-471 


C. velox 

84 98 EXF-334 

EXF-382 

C. spinulosum 

CBS 170.54 C. cladosporioides 

89 


ATCC 66669 

C. oxysporum 

dH 12862 

dH 13911 

EXF-646 

EXF-703 

EXF-228 

EXF-1072 C. halotolerans 

EXF-572 

EXF-977 

EXF-564 

EXF-380 


C. langeronii 

10 steps 

Fig. 4. One of 32 equally most parsimonious and equally looking phylogenetic trees based on a heuristic tree search using aligned exons and introns of the partial actin gene. 

The tree was randomly selected. Support based on 10 000 replicates of a fast step-wise addition bootstrap analysis is indicated near the branches. Trees were rooted with 

the strains of Cladosporium salinae. Most monophyletic species clades received high, but deeper branches weak or no, bootstrap support (CI = 0.586, RI = 0.885, PIC = 103). 

growing colonies at all tested temperatures and relatively large, 

oblong conidia. However, phylogenetic analyses of the β-tubulin and 

actin gene indicate that C. langeronii presents two cryptic species 

(Figs 3–4). The species clade of C. psychrotolerans is moderately 

supported in analyses of the actin gene but highly by means of 

the β-tubulin gene. It is evident from all three analyses (Figs 2– 

4) that C. langeronii and C. psychrotolerans are closely related 

species. The species node of Cladosporium spinulosum, which is 

morphologically clearly distinguished from all other species by its 

conspicuous ornamentation consisting of digitate projections (Fig. 

5), is supported by β-tubulin (Fig. 3) and actin (Fig. 4) sequence 

data but not by those of the ITS rDNA (Fig. 2). Analyses of all loci, 

however, indicate that it is a member of the C. herbarum complex. 

The analyses of sequences of the ITS and the β-tubulin 

and actin gene introns and exons (Figs 2–4) do not allow the 

full elucidation of phylogenetic relationships among these 

Cladosporium species. Statistical support of the interior tree 

branches resulting from analyses of the β-tubulin and actin genes 

is low (bootstrap values mostly < 50 %). While the sister group 

relationship of C. sphaerospermum and C. fusiforme is highly 

supported in the analysis based on the β-tubulin gene, analysis 

of the ITS rDNA indicate that these two species are unrelated, and 

that C. sphaerospermum is closely related to C. dominicanum. It is 

clear from the data that the species morphologically resembling C. 

sphaerospermum are not phylogenetically closely related and that 

the data we present here do not allow their classification in natural 

subgroups of the genus Cladosporium. Only C. spinulosum was 

placed in all analyses among species of the C. herbarum complex 


165

Zalar et al. 

Fig. 5. Conidial scars and surface ornamentation of ramoconidia and conidia (SEM). A. C. dominicanum (strain EXF-732). B. C. fusiforme (strain EXF-449). C. C. halotolerans 

(strain EXF-572). D. C. langeronii (strain CBS 189.54). E. C. psychrotolerans (strain EXF-391). F. C. salinae (strain EXF-335 = CBS 119413). G. C. sphaerospermum (strain 

CBS 193.54). H. C. spinulosum (strain EXF-334). I. C. velox (strain CBS 119417). Scale bars = 5 µm. (Photos: K. Drašlar). 

and all analyses supported close relatedness of C. langeronii and 

C. psychrotolerans. 

The majority of species described here have slightly 

ornamented conidia ranging from minutely verruculose (C. 

fusiforme, C. langeronii, C. psychrotolerans, C. sphaerospermum, 

C. velox) to verrucose (C. halotolerans) (Fig. 5). The verrucose 

conidia of C. halotolerans can be recognised also under the 

light microscope and used as a distinguishing character. Almost 

smooth to minutely verruculose conidia are encountered in C. 

dominicanum and C. salinae (Fig. 5). Cladosporium spinulosum, 

a member of the C. herbarum species complex, has conidia with 

a digitate ornamentation that can appear spinulose under the light 

microscope; however, when using the SEM it became clear that its 

projections have parallel sides and a blunt end (Fig. 5). 

DISCUSSION 

The genus Cladosporium was established by Link (1816) who 

originally included four species, of which C. herbarum is the 

type species of the genus (Clements & Shear 1931). In 1950, 

von Arx reported a teleomorph connection for this species with 

Mycosphaerella tassiana (De Not.) Johanson. Based on SSU 

rDNA data the majority of Mycosphaerella species, including 

the type species of the genus, M. punctiformis (Pers.) Starbäck, 

clustered within the Mycosphaerellaceae, a family separated 

from M. tassiana (Braun et al. 2003). Therefore, Mycosphaerella 

tassiana was reclassified as Davidiella tassiana (De Not.) Crous 

& U. Braun, the type of the new genus Davidiella. All anamorphs 

with a cladosporium- and heterosporium-like appearance and with 

a supposed Dothideomycetes relationship were maintained under 

the anamorph name Cladosporium, morphologically characterised 

by scars with a protuberant hilum consisting of a central dome 

surrounded by a raised rim (David 1997). 

The concept of distinguishing ramoconidia from secondary 

ramoconidia has been adopted from Schubert et al. (2007). In 

the species described here, ramoconidia have been observed 

often in C. sphaerospermum, sometimes in C. psychrotolerans, 

C. langeronii and C. spinulosum, and only sporadically in all other 

species. Therefore, ramoconidia can be seen as important for 

distinguishing species although sometimes, they can be observed 

only with difficulty. When using ramoconidia as a diagnostic 

criterion, colonies only from SNA and not older than 7 d should be 

taken into account. 

Cladosporium sphaerospermum was described by Penzig 

(1882) from decaying Citrus leaves and branches in Italy. He 

described C. sphaerospermum as a species with (i) branched, 

septate and dark conidiophores having a length of 150–300 µm and 

a width of the main conidiophore stipe of 3.5–4 µm, (ii) spherical to 

ellipsoid, acrogenously formed conidia of 3.4–4 µm diam, and (iii) 

ramoconidia of 6–14 × 3.5–4 µm. Penzig’s original material is not 

known to be preserved. Later, a culture derived from CBS 193.54, 

originating from a human nail, was accepted as typical of C. 

sphaerospermum. However, de Vries (1952), incorrectly cited it as 

“lectotype”, and thus the same specimen is designated as neotype 

in this study (see below), with the derived culture (CBS 193.54) 

166


used as ex-neotype strain. Numerous strains with identical or very 

similar ITS rDNA sequences as CBS 193.54 were isolated from 

hypersaline water or organic substrata including plants or walls of 

bathrooms. It is not clear yet whether surfaces in bathrooms and of 

plants, colonised by C. sphaerospermum, can have a similar low 

water activity as salterns. In our experiments, the strains of this 

species, however, grew under in vitro conditions at a water activity 

of up to 0.860, while Hocking et al. (1994) and Aihara et al. (2002) 

reported that it can grow even at 0.815. Therefore, we consider 

C. sphaerospermum as halo- or osmotolerant. Hardly any reports 

are available unambiguously proving that C. sphaerospermum is 

a human pathogen. It is therefore possible that CBS 193.54 was 

not involved in any disease process but rather occurred as a 

contaminant on dry nail material. Cladosporium sphaerospermum 

is a phylogenetically well-delineated species (Figs 2–4). 

Strains of C. halotolerans were isolated sporadically from 

substrata such as peanut cell suspension, tissue culture, bathroom 

walls and as culture contaminants. This surprising heterogeneity 

of substrata suggests that C. halotolerans is distributed by air and 

that it can colonise whatever substrata available, although it may 

have its natural niche elsewhere. We have recurrently isolated it 

from hypersaline water of salterns and other saline environments 

and it was also detected with molecular methods (but not isolated) 

from skin of a salt water dolphin. There are only few reports of 

this species from plants (Table 1). It is therefore possible that C. 

halotolerans is a species closely linked to salty or hypersaline 

environments although additional sampling is necessary to prove 

that. Cladosporium halotolerans is morphologically recognisable 

by relatively oblong to spherical, coarsely rough-walled conidia. 

The ITS rDNA sequence of a fungus in the skin of a bottlenose 

dolphin, suffering from lobomycosis, is identical to the sequences 

of C. halotolerans. This sequence was deposited as Lacazia 

loboi Taborda, V.A. Taborda & McGinnis (GenBank AF035674) by 

Haubold et al. (1998), who apparently concluded wrongly that a 

fungus with a cladosporium-like ITS rDNA sequence similar to that 

of C. halotolerans can be the agent of lobomycosis. Later, Herr 

et al. (2001) showed that Lacazia loboi phylogenetically belongs 

to the Onygenales on the basis of amplified SSU rDNA and chitin 

synthase-2 gene sequences generated from tissue lesions. By 

this, they confirmed an earlier supposition by Lacaz (1996) who 

reclassified the organism as Paracoccidioides loboi O.M. Fonseca 

& Silva Lacaz (Onygenales). It is therefore possible that C. 

halotolerans was not the main etiologic agent for the lobomycosis 

and it was colonising the affected dolphin skin secondarily while 

inhabiting other seawater habitats. 

Cladosporium langeronii and C. psychrotolerans are closely 

related but C. langeronii is particularly well distinguishable from 

all other Cladosporium species by its slow growing colonies 

(1–7 mm diam / 14 d) and relatively large conidia (4–5.5 × 3–4 

μm). Cladosporium psychrotolerans has smaller conidia (3–4 × 

2.5–3 μm) but a similar length : width ratio and faster expanding 

colonies (8–18 mm diam / 14 d). Cladosporium langeronii is most 

likely a complex of at least two species. Strains isolated from the 

Arctic and the Antarctic may need to be distinguished from C. 

langeronii s. str. on species level. This inference is particularly 

supported by analyses of the β-tubulin and actin genes (Figs 3–4). 

Cladosporium langeronii s. str., represented by an authentic strain 

of Hormodendrum langeronii Fonseca, Leão & Nogueira, CBS 

189.54 (Trejos 1954), has been isolated from a variety of substrata 

but is tolerating only up to 10 % NaCl. It was originally described 

by da Fonseca et al. (1927a, b) and subsequently reclassified 

as Cladosporium langeronii by Vuillemin (1931). The authentic 


strain derived from an ulcerating nodular lesion on the arm of a 

human patient. Because other strains of this species are ubiquitous 

saprobes originating from various substrata, we suspect that C. 

langeronii is not an important human pathogen. Cladosporium 

psychrotolerans has been isolated from hypersaline environments 

only, and tolerates up to 20 % NaCl in culture media. 

In general, the human- or animal-pathogenic role of the C. 

sphaerospermum-like species described here seems to be limited. 

It is possible that pathogenic species of Cladophialophora Sacc. 

have been misidentified as C. sphaerospermum or as other 

species of Cladosporium (de Hoog et al. 2000). Alternatively, true 

Cladosporium species isolated as clinical strains could have been 

secondary colonisers since they are able to dwell on surfaces poor 

in nutrients, possibly in an inconspicuous dormant phase and may 

then be practically invisible. More likely, they could be air-borne 

contaminations of lesions, affected nails etc. (Summerbell et al. 

2005) or are perhaps disseminated by insufficiently sterilised medical 

devices, as melanised fungi can be quite resistant to disinfectants 

(Phillips et al. 1992). They can easily be isolated and rapidly 

become preponderant at isolation and thus difficult to exclude as 

etiologic agents of a disease. For example, in 2002, a case report 

on an intrabronchial lesion by C. sphaerospermum in a healthy, 

non-asthmatic woman was described (Yano et al. 2002), but we 

judge the identification of the causal agent to remain uncertain, as 

it was based on morphology alone and no culture is available. The 

present authors have the opinion that all clinical cases ascribed to 

Cladosporium species need careful re-examination. 

General characteristics and description of Cladosporium 

sphaerospermum-like species 

The present paper focuses on Cladosporium strains isolated from 

hypersaline environments. Comparison of data from deliberate 

sampling and analysis of reference strains from culture collections 

inevitably leads to statistical bias, and therefore a balanced 

interpretation of ecological preferences of the species presented is 

impossible. Nevertheless, some species appeared to be consistent 

in their choice of habitat, and for this reason we summarise 

isolation data for all species described. Strains belonging to a single 

molecular clade proved to have similar cultural characteristics and 

microscopic morphology. Although within most of the species there 

was some molecular variation noted (particularly when intron-rich 

genes were analysed), some consistent phenetic trends could be 

observed. 

Conidiophores of all C. sphaerospermum-like species lack 

nodose inflations (McKemy & Morgan-Jones 1991). They are 

usually ascending and can sometimes be poorly differentiated from 

their supporting hyphae. Though the initiation point of conidiophore 

stipes could sometimes be determined only approximately, 

their lengths were in some cases useful for distinguishing 

morphologically similar species when observed in slide cultures. 

Generally, the branched part of a conidiophore forms a complex 

tree-like structure. The number and orientation of early formed 

secondary ramoconidia, however, determines whether it is 

rotationally symmetric or unilateral. 

The variability in ITS rDNA sequences observed in all C. 

sphaerospermum-like species (about 10 %) spans the variation 

observed in all members of the genus Cladosporium sequenced 

to date. Thus, the C. sphaerospermum-like species described here 

may not present a single monophyletic group but may belong to 

various species complexes within Cladosporium. Verifying existing 

literature with sequence data of these species (Wirsel et al. 2002, 

Park et al. 2004), we noticed that names of the common saprobes 

seem to be distributed nearly at random over phylogenetic trees. 

167

Zalar et al. 

For most commonly used names, no type material is available for 

sequencing. Also verification of published reports is difficult without 

available voucher strains. 

Cladosporium cladosporioides was incorrectly lectotypified 

based on CBS 170.54 (de Vries 1952), which Bisby considered a 

standard culture of C. herbarum. The C. cladosporioides species 

complex requires revision, and will form the basis of a future study. 

Cladosporium herbarum is maintained as a dried specimen in the 

Leiden herbarium; Prasil & de Hoog (1988) selected CBS 177.71 

as a representative living strain. Strains, earlier accepted as living 

representatives of C. herbarum, CBS 177.71 and CBS 812.71 

(Prasil & de Hoog 1988, Wirsel et al. 2002) and ATCC 66670 

(Braun et al. 2003, as Davidiella tassiana) have been re-identified 

as C. bruhnei Linder by Schubert et al. (2007 – this volume). Ho 

et al. (1999) used strain ATCC 38027 as a representative of C. 

tenuissimum Cooke and this strain has identical ITS sequences as 

the non-deposited C. tenuissimum material used by Moricca et al. 

(1999). We tentatively accept this concept although we could not 

include ATCC 38027 in our analyses. The ITS sequence of strain 

CBS 125.80, identified by Wirsel et al. (2002) as C. oxysporum, 

is identical to the sequence of ATCC 38027. Strain ATCC 76499, 

published by Ho et al. (1999) as C. oxysporum, appears to be 

identical to a number of currently unidentified Cladosporium strains 

from Slovenian salterns that compose a cluster separate from all 

remaining species. Strains of this cluster, represented in Fig. 2 by 

strain ATCC 76499, morphologically resemble C. oxysporum. 

Strain CBS 300.96 has not been identified to species level 

in the present study. It clusters outside the species clade of 

C. sphaerospermum, with the latter being its nearest relative. 

CBS 300.96 differs from C. sphaerospermum by having smaller 

structures: conidiophore stipes [(5–)20–80(–150) × (2–)2.5–3(–4) 

μm], 0–1 septate ramoconidia [(13–)19–27(–32) × 2–2.5 μm], 

conidia [(2.5–)3–3.5(–4) × (2–)2–2.5(–3) μm] and secondary 

ramoconidia [(5–)9–18(–30) × (2–)2.5–2.5(–3) μm]. However, 

based on a single isolate, we currently refrain from describing it as 

a new species. 

Key to species treated in this study 

Macro-morphological characters used in the key are from colonies grown on PDA and MEA 14 d at 25 °C, if not stated otherwise; microscopical 

characters are from SNA slide cultures grown for 7 d at 25 °C. 

1. Conidial ornamentation conspicuously echinulate / digitate because of up to 1.3 µm long projections that have more or less parallel sides 

............................................................................................................................................................................................. C. spinulosum 

1. Conidial ornamentation verruculose to verrucose or smooth, not conspicuously echinulate or digitate .................................................... 2 

2. Conidiophores micronematous, poorly differentiated, once or several times geniculate-sinuous, short, up to 60 µm long; terminal conidia 

obovoid ....................................................................................................................................................................................... C. salinae 

2. Conidiophores micro- or macronematous, not geniculate or only slightly so, usually up to 100 µm or 220 µm long or even longer; terminal 

conidia globose, subglobose to ovoid or fusiform ...................................................................................................................................... 3 

3. Secondary ramoconidia 0–3(–4)-septate; septa of conidiophores and conidia darkened and thickened .................................................. 4 

3. Secondary ramoconidia 0–1(–2)-septate; septa neither darkened nor thickened ..................................................................................... 5 

4. Conidiophores (5–)10–50(–300) × (2–)2.5–3(–5.5) μm; terminal conidia (2–)3–4(–6) × (2–)2.5–3(–5) μm; secondary ramoconidia (5–)7– 

12(–37.5) × (2–)2.5–3(–6.5) μm; ramoconidia sporadically formed ................................................................................... C. halotolerans 

4. Conidiophores mostly longer and somewhat wider, (10–)45–130(–300) × (2.5–)3–4(–6) μm; terminal conidia mostly wider, (2.5–)3–4(–7) 

× (2–)3–3.5(–4.5) μm; secondary ramoconidia (4–)8.5–16(–37.5) × (2–)3–3.5(–5) μm; ramoconidia often formed, up to 40 µm long, with 

up to 5 septa ............................................................................................................................................................. C. sphaerospermum 

5. Terminal conidia usually fusiform ............................................................................................................................................ C. fusiforme 

5. Terminal conidia globose, subglobose or ovoid ......................................................................................................................................... 6 

6. Conidia and secondary ramoconidia irregularly verruculose to sometimes loosely verrucose; radial growth on PDA at 25 °C after 14 d 

typically less than 5 mm ......................................................................................................................................................... C. langeronii 

6. Conidia and secondary ramoconidia smooth to minutely verruculose; radial growth on PDA at 25 °C after 14 d typically more than 10 mm 

.................................................................................................................................................................................................................... 7 

7. Conidiophores (3–)3.5–4(–7.5) μm wide, thick-walled; conidiogenous loci and conidial hila 0.5–2 μm diam; ramoconidia sometimes 

formed with a broadly truncate, up to 2 µm wide non-cladosporioid base; no growth observed after 14 d at 30 °C on MEA 

..................................................................................................................................................................................... C. psychrotolerans 

7. Conidiophores mostly narrower, 2–4 μm wide, only with slightly thickened walls; conidiogenous loci and conidial hila narrower, 0.5–1.5 

μm diam; ramoconidia rarely formed; colony showing at least weak growth after 14 d at 30 °C on MEA ................................................. 8 

8. Secondary ramoconidia (4–)6.5–13(–24.5) μm long; no visible colony growth after 14 d at 10 °C on MEA.................... C. dominicanum 

8. Secondary ramoconidia mostly longer, (3.5–)5.5–19(–42) μm; radial growth of colonies after 14 d at 10 °C on MEA more than 5 mm 

........................................................................................................................................................................................................ C. velox 

168


Description of Cladosporium species 

Cladosporium dominicanum Zalar, de Hoog & Gunde-Cimerman, 

sp. nov. MycoBank MB510995. Fig. 6. 

Etymology: Refers to the the Dominican Republic, where most 

strains were encountered. 

Conidiophora lateralia vel terminalia ex hyphis rectis oriunda; stipes longitudine 

variabili, (5–)10–100(–200) × (1.5–)2–2.5(–3.5) μm, olivaceo-brunneus, levis vel 

leniter verruculosus, tenuitunicatus, plerumque unicellularis, simplex vel ramosus. 

Conidiorum catenae undique divergentes, ad 8 conidia in parte continua continentes. 

Cellulae conidiogenae indistinctae. Conidia levia vel leniter verruculosa, dilute 

brunnea, unicellularia, plerumque breviter ovoidea, utrinque angustata, (2.5–) 

3–3.5(–5.5) × (2–)2–2.5(–2.5) μm, long.: lat. 1.4–1.6; ramoconidia secundaria 

cylindrica vel quasi globosa, 0–1-septata, (4–)6.5–13(–24.5) × (2–)2.5–3(–4.5) μm, 

ad 4 cicatrices terminales ferentia; cicatrices inspissatae, protuberantes, 0.5–1.2 μm 

diam. Hyphae vagina polysaccharidica carentes. 

Mycelium without extracellular polysaccharide-like material. 

Conidiophores arising laterally and terminally on erect hyphae, 

micronematous and semimacronematous, stipes of variable length, 

(5–)10–100(–200) × (1.5–)2–2.5(–3.5) μm, olivaceous-brown, 

smooth to minutely verruculose, thin-walled, almost non-septate, 

unbranched or branched. Conidial chains branching in all directions, 

up to eight conidia in the unbranched parts. Conidiogenous cells 

undifferentiated. Ramoconidia rarely formed. Conidia smooth 

to minutely verruculose, subhyaline to light brown, non-septate, 

usually short-ovoid, narrower at both ends, length : width ratio = 

1.4–1.6; (2.5–)3–3.5(–5.5) × (2–)2–2.5(–2.5) μm [av. (± SD) 3.4 

(± 0.6) × 2.2 (± 0.2)]; secondary ramoconidia cylindrical to almost 

spherical, 0–1-septate, (4–)6.5–13(–24.5) × (2–)2.5–3(–4.5) μm 

[av. (± SD) 10.3 (± 5.2) × 2.7 (± 0.6)], with up to four distal scars. 

Conidiogenous scars thickened and conspicuous, protuberant, 

0.5–1.2 μm diam. 


diam, olive-yellow (2D6), hairy granular, flat or slightly furrowed, 

with flat margin. Droplets of light reseda-green (2E6) exudate 

sometimes present. Reverse dark green to black. Colonies on OA 

reaching 19–34 mm diam, olive (2F5), loosely powdery with raised 

central part due to fasciculate bundles of conidiophores. Reverse 

dark green. Colonies on MEA reaching 30–32 mm diam, reseda 

green (2E6), velvety, furrowed, with undulate margin. Reverse dark 

green-brown. Colonies on MEA + 5 % NaCl reaching 37–41 mm 

diam, reseda-green (2E6), radially furrowed, velvety, sporulating in 

the central part or all over the colony, margin white and regular. 

Reverse brownish green. 

Maximum tolerated salt concentration: 75 % of tested strains 

develop colonies at 20 % NaCl after 7 d, while after 14 d all strains 

grow and sporulate. 

Cardinal temperatures: No growth at 4 and 10 °C, optimum 25 °C 

(30–32 mm diam), maximum 30 °C (2–15 mm diam), no growth at 

37 °C. 

Specimen examined: Dominican Republic, from hypersaline water of salt lake 

Enriquillo, coll. Nina Gunde-Cimerman, Jan. 2001, isol. P. Zalar 25 Feb. 2001, CBS 

H-19733, holotype, culture ex-type EXF-732 = CBS 119415. 

Habitats and distribution: Fruit surfaces; hypersaline waters in 

(sub)tropical climates. 

Differential parameters: No growth at 10 °C, ovoid conidia, large 

amounts of sterile mycelium. 

Strains examined: CPC 11683, EXF-696, EXF-718, EXF-720, EXF- 

727, EXF-732 (= CBS 119415; ex-type strain). 


Note: Cultures of C. dominicanum sporulate less abundantly than 

C. sphaerospermum and C. halotolerans and tend to lose their 

ability to sporulate with subculturing. 

Cladosporium fusiforme Zalar, de Hoog & Gunde-Cimerman, sp. 


Etymology: Refers to its usually fusiform conidia. 

Conidiophora erecta, lateralia vel terminalia ex hyphis rectis oriunda; stipes 

longitudine variabili, (10–)25–50(–100) × (2–)2–3.5(–4) μm, olivaceo-brunneus, 

levis, crassitunicatus, compluries septatus (cellulis 9–23 μm longis), plerumque 

simplex. Conidiorum catenae undique divergentes, in parte continua ad 5 conidia 

continentes. Cellulae conidiogenae indistinctae. Conidia leniter verruculosus, dilute 

brunnea, unicellularia, plerumque fusiformia, utrinque angustata, (2.5–)3.5–5(– 

6.5) × (2–)2–2.5(–3) μm, long. : lat. 1.8–2.0; ramoconidia secundaria cylindrica, 

0(–1)-septata, (5–)6–11(–22) × (2.5–)2.5–3(–3) μm, ad 4 cicatrices terminales 

ferentia; cicatrices inspissatae, conspicuae, 0.7–1.0 μm diam. Hyphae vagina 

polysaccharidica carentes. 

Mycelium without extracellular polysaccharide-like material. 

Conidiophores erect, arising laterally and terminally from straight 

hyphae, stipes of variable length, (10–)25–50(–100) × (2–)2–3.5(– 

4) μm, olivaceous-brown, smooth- and thick-walled, regularlyseptate 

(cell length 9–23 μm), mostly unbranched. Conidial chains 

branching in all directions, up to 5 conidia in the unbranched parts. 

Conidiogenous cells undifferentiated. Ramoconidia rarely formed. 

Conidia minutely verruculose, light brown, aseptate, usually fusiform 

and narrower at both ends, length : width ratio = 1.8–2.0; (2.5–)3.5– 

5(–6.5) × (2–)2–2.5(–3) μm [av. (± SD) 4.4 (± 0.8) × 2.2 (± 0.2)]; 

secondary ramoconidia cylindrical, 0(–1)-septate, (5–)6–11(–22) × 

(2.5–)2.5–3(–3) μm [av. (± SD) 9.0 (± 4.7) × 2.6 (± 0.3)], with up 

to 4 distal scars. Conidiogenous scars thickened and conspicuous, 

protuberant, 0.7–1.0 μm diam. 

Cultural characteristics: Colonies on PDA reaching 20–26 mm diam, 

dull green (30E3), granular due to profuse sporulation, flat, with flat 

margin. Sterile mycelium absent. Reverse blackish green. Colonies 

on OA reaching 24–28 mm diam, olive (3F3), granular in concentric 

circles, consisting of two kinds of conidiophores (low and high), flat, 

with flat margin. Reverse black. Colonies on MEA reaching 23–28 

mm diam, olive (3E5), deeply furrowed, velvety (sporulating all over) 

with undulate, white margin. Reverse brownish green. Colonies on 

MEA + 5 % NaCl reaching 28–43 mm diam, olive (3E6), granular 

due to profuse sporulation, slightly furrowed with flat, olive-grey 

(3F2) margin. Reverse dark green. 

Maximum tolerated salt concentration: Only one of three strains 

tested (CBS 452.71) developed colonies at 17 % NaCl after 14 d, 

the other two strains grew until 10 % NaCl. 

Cardinal temperatures: For one of three strains (CBS 452.71) the 

minimum temperature of growth was 4 °C (6 mm diam), for the 

other two 10 °C (8–9 mm diam); optimum 25 °C (23–28 mm diam), 

maximum 30 °C (only strain CBS 452.71 grew 5 mm diam), no 

growth at 37 °C. 

Specimen examined: Slovenia, from hypersaline water of Sečovlje salterns, coll. 

and isol. L. Butinar, Dec. 1999, CBS H-19732, holotype, culture ex-type EXF-449 

= CBS 119414. 

Habitats and distribution: Osmotic environments worldwide. 

Differential parameters: Oblong conidia, relatively low degree of 

halotolerance. 

Strains examined: CBS 452.71, EXF-397, EXF-449 (= CBS 119414; 

ex-type strain). 

169

Zalar et al. 

Fig. 6. Cladosporium dominicanum. Macro- and micromorphological characters. A–D. Colony surface grown on PDA (A), OA (B), MEA (C) and MEA plus 5 % NaCl (D) of strains 

incubated for 14 d at 25 ºC in darkness. E–F. Habit of conidiophores. G. Conidiophore. H–I. Secondary ramoconidia and conidia. E–I. All from 7-d-old SNA slide cultures. A, D, 

F–H, from EXF-2519; B, C, E from EXF-727; I, EXF-732 (ex-type strain). Scale bars A–D = 10 mm, E = 100 µm, F = 30 µm, G–I = 10 µm. 

170


Fig. 7. Cladosporium fusiforme. Macro- and micromorphological characters. A–D. Colony surface grown on PDA (A), OA (B), MEA (C) and MEA plus 5 % NaCl (D) of strains 

incubated for 14 d at 25 ºC in darkness. E–G. Habit of conidiophores. H–I. Ramoconidia and conidia. E–I. All from 7-d-old SNA slide cultures. A–H, from EXF-449 (ex-type strain); 

I, from CBS 452.71. Scale bars A–D = 10 mm, E = 100 µm, F–G = 30 µm, H–I = 10 µm. 


171

Zalar et al. 

Fig. 8. Cladosporium halotolerans. Macro- and micromorphological characters. A–D. Colony surface grown on PDA (A), OA (B), MEA (C) and MEA plus 5 % NaCl (D) of strains 

incubated for 14 d at 25 ºC in darkness. E–H. Habit of conidiophores. I. Conidiophore. J. Succession of secondary ramoconidia. K. Conidia. E–K. All from 7-d-old SNA slide 

cultures. A–B, from EXF-572 (ex-type strain); C–D, from EXF-977; E, G, from EXF-972; F, from EXF-564; H, I, K, from EXF-1072; J, from dH 12862. Scale bars A–D = 10 mm, 

E = 100 µm, F–G = 50 µm, H = 30 µm, I–K = 10 µm. 

Cladosporium halotolerans Zalar, de Hoog & Gunde-Cimerman 


Etymology: Refers to its halotolerant habit. 

Conidiophora erecta, lateralia vel terminalia ex hyphis rectis oriunda; stipes 

longitudine variabili, (5–)10–50(–300) × (2–)2.5–3(–5.5) μm, pallide olivaceobrunneus, 

levis vel leniter verruculosus, tenuitunicatus, 0–3-septatus, interdum 

pluriseptatus, simplex, denticulatus. Conidiorum catenae undique divergentes, 

terminales ad 9 conidia continentes. Cellulae conidiogenae indistinctae. Conidia 

verrucosa, brunnea vel fusca, unicellularia, plerumque subglobosa vel globosa, 

raro breviter ovoidea, utrinque angustata, (2–)3–4(–6) × (2–)2.5–3(–5) μm, long. 

: lat. 1.2–1.5; ramoconidia secundaria cylindrica vel quasi globosa, 0(–1)-septata, 

(5–)7–12(–37.5) × (2–)2.5–3(–6.5) μm , ad 4 cicatrices terminales ferentia; cicatrices 

inspissatae, conspicuae, protuberantes, 0.7–1.0(–1.5) μm diam. Hyphae vagina 

polysaccharidica carentes. 

172


Mycelium partly submerged, partly superficial; hyphae without 

extracellular polysaccharide-like material. Conidiophores erect, 

arising laterally and terminally from straight hyphae, stipes of 

variable length, (5–)10–50(–300) × (2–)2.5–3(–5.5) μm, pale 

olivaceous-brown, smooth to minutely verruculose, thin-walled, 0– 

3-septate, unbranched, with pronounced denticles. Conidial chains 

branching in all directions, terminal chains with up to 9 conidia. 

Conidiogenous cells undifferentiated. Ramoconidia rarely formed. 

Conidia verrucose, brown to dark brown, non-septate, usually 

subglobose to globose, less often short-ovoid, narrower at both 

ends, length : width ratio = 1.2–1.5; (2–)3–4(–6) × (2–)2.5–3(–5) 

μm [av. (± SD) 3.5 (± 0.7) × 2.7 (± 0.5)]; secondary ramoconidia 

cylindrical to almost spherical, 0–1-septate, (5–)7–12(–37.5) × 

(2–)2.5–3(–6.5) μm [av. (± SD) 10.3 (± 4.8) × 2.9 (± 0.6)], with up 


protuberant, 0.7–1.0(–1.5) μm diam. 


olive (2F5), slightly furrowed, often covered with grey secondary 

mycelium, except at the marginal area where only sporulating 

structures can be observed. Margin white and regular, with 

submerged hyphae. Reverse pale green to black. Colonies on OA 

reaching 29–40 mm diam, olive (2F6), flat, uniform, granular due 

to profuse sporulation and fasciculate bundles of conidiophores, 

without sterile mycelium. Reverse dark green to black. Colonies on 

MEA reaching 18–44 mm diam, highly variable in colour, but mainly 

olive (2E5), and from flat with regular margin to deeply furrowed 

with undulate margin. Colony centre wrinkled with crater-shaped 

appearance. Reverse pale to dark green. Colonies on MEA + 5 % 

NaCl reaching 24–48 mm diam, olive (3E8), furrowed, velvety, with 

more pale, undulate margins. Reverse dark green to black. 

Maximum tolerated salt concentration: Only 15 % of tested strains 

develop colonies at 20 % NaCl after 7 d, whereas after 14 d all 

cultures grow and sporulate. 

Cardinal temperatures: No growth at 4 °C, optimum 25 °C (18–44 

mm diam), maximum 30 °C (6–23 mm diam). No growth at 37 °C. 

Specimen examined: Namibia, from hypersaline water of salterns, coll. Nina Gunde- 

Cimerman, 1 Sep. 2000, isol. P. Zalar, 1 Oct. 2000, CBS H-19734, holotype, culture 

ex-type EXF-572 = CBS 119416. 

Habitats and distribution: Hypersaline water in subtropical climates; 

indoor environments; Arctic ice; contaminant in lesions of humans 

and animals; plant phyllosphere; rock. 

Literature: Haubold et al. (1998), Meklin et al. (2004). 

Differential parameters: Verrucose conidia, short unbranched and 

non-septate conidiophores which arise laterally alongside erect 

hyphae. 

Strains examined: CBS 191.54, CBS 573.78, CBS 626.82, dH 

12862, dH 12991, dH 13911, EXF-228, EXF-380, EXF-565, EXF- 

567, EXF-571, EXF-572 (= CBS 119416; ex-type strain), EXF-646, 

EXF-698, EXF-703, EXF-944, EXF-972, EXF-977, EXF-1072, 

EXF-2372. 

Notes: Cladosporium halotolerans strongly resembles C. 

sphaerospermum. Several strains of this species such as dH 

12862, dH 12941, CBS 191.54 and UAMH 7686 have been 

isolated sporadically from various indoor habitats in Europe, Brazil 

and the U.S.A. and repeatedly from bathrooms in Slovenia (Table 

1). Probably sometimes as uncertain culture contaminations, it 

has been isolated from plants (GenBank accession no. L25433), 


inner organs of a diseased frog (AY361982) and human brain 

(Kantarcioglu et al. 2002). The presence of C. halotolerans 

species in gypsum sediments entrapped in Arctic ice, the fact that 

it was repeatedly isolated from hypersaline water and possibly its 

presence in dolphin skin (see Discussion) suggest that it has a 

clear preference for (hyper)osmotic habitats. This is supported by 

its ability to grow at 20 % NaCl. 

The teleomorph of C. halotolerans is predicted to be a 

Davidiella species. Strain CBS 280.49 was isolated by J.A. von Arx 

from teleomorphic material of a fungus labelled as Mycosphaerella 

hyperici (Auersw.) Starbäck on Hypericum perforatum in 

Switzerland. According to Aptroot (2006) this species may belong 

in Davidiella and produces a Septoria anamorph. In the original 

herbarium specimen, CBS H-4867, a Mycosphaerella teleomorph 

was present, but no sign of a Cladosporium anamorph. We assume 

that CBS 280.49 was a culture contaminant. 

Cladosporium langeronii (Fonseca, Leão & Nogueira) Vuill., 

Champ. Paras.: 78. 1931. Fig. 9. 

Basionym: Hormodendrum langeronii Fonseca, Leão & Nogueira, 

Sci. Med. 5: 563. 1927. 

≡ Cladosporium langeronii (Fonseca, Leão & Nogueira) Cif., Manuale di 

Micologia Medica, ed. 2: 488 (1960), comb. superfl. 

Mycelium partly submerged, partly superficial; hyphae sometimes 

enveloped in polysaccharide-like material. Conidiophores erect or 

ascending, micronematous and macronematous, stipes of variable 

length, (20–)50–130(–200) × (3–)3.5–4.5(–6.5) μm, dark brown, 

rough- and thick-walled, regularly septate (cell length 9–22 μm), 

arising laterally and terminally from submerged or aerial hyphae, 

branched. Conidial chains dichotomously branched, up to 6 conidia 

in the unbranched parts. Conidiogenous cells undifferentiated, 

sometimes seceding and forming ramoconidia. Ramoconidia 

cylindrical, 0–1 septate, (10–)11–22(–42) × (3–)3.5–4.5(–5) µm, 

base broadly truncate, 2–3.5 µm wide, slightly thickened and 

somewhat darkened. Conidia irregularly verruculose to sometimes 

loosely verrucose, dark brown, non-septate, usually ovoid, length 

: width ratio = 1.3–1.5; conidial size (3–)4–5.5(–8) × (2–)3–4(–5) 


cylindrical to almost spherical, mostly 0–1(–2)-septate, (5.5–)7.5– 

12.5(–35.5) × (2.5–)3–4.5(–5.5) μm [av. (± SD) 10.7 (± 4.7) × 3.6 (± 

0.8)], with 2, rarely 3 distal scars. Conidiogenous scars thickened 

and conspicuous, protuberant, 0.9–1.5(–2.3) μm diam. 

Cultural characteristics: Colonies on PDA, OA and MEA with 

restricted growth, attaining 2.5–4.5, 1.5–7.0 and 1.0–5.5 mm 

diam, respectively. Colonies flat or heaped (up to 3 mm), dark 

green (30F4), with black reverse and slightly undulate margin with 

immersed mycelium. Sporulating on all media. On MEA + 5 % NaCl 

growth is faster, colonies attaining 8.5–12.0 mm diam, sporulating 

and growing deeply into the agar. 

Maximum tolerated salt concentration: All strains develop colonies 

at 17 % NaCl after 14 d. 

Cardinal temperatures: No growth at 4 °C, optimum / maximum 25 

°C (1.0–5.5 mm diam), no growth at 30 °C. 

Specimen examined: Brazil, from man ulcero-nodular mycosis of hand and arm, 

1927, coll. and isol. da Fonseca, CBS H-19737, holotype, culture ex-type CBS 

189.54. 

Habitats and distribution: Polar ice and biomats; conifer wood and 

window frame in Europe; humans; strains originating from nasal 

mucus (Buzina et al. 2003) have 100 % sequence homology with 

173

Zalar et al. 

Fig. 9. Cladosporium langeronii. Macro- and micromorphological characters. A–D. Colony surface grown on PDA (A), OA (B), MEA (C) and MEA plus 5 % NaCl (D) of strains 

incubated for 14 d at 25 ºC in darkness. E–F. Habit of conidiophores. G–I. Ramoconidia and conidia. E–I. All from 7-d-old SNA slide cultures. A–D, from CBS 189.54 (ex-type 

strain); E, from CBS 109868; F–I, from EXF-999. Scale bars A, C–D = 10 mm, B = 5 mm, E = 100 µm, F = 30 µm, G–I = 10 µm. 

the strains studied, as well as with a clone from mycorrhizal roots 

(Menkis et al. 2005). The species is distributed worldwide, without 

any apparent predilection for a particular habitat. The strains from 

clinical cases probably were culture contaminants. 

Literature: da Fonseca et al. (1927a, b). 

Differential parameters: Restricted growth; lowest salt halotolerance 

taxon of all C. sphaerospermum-like species. 

Strains examined: CBS 189.54 (ex-type strain), CBS 601.84, CBS 

101880, CBS 109868, dH 11736, dH 12459 = EXF-999, dH 13833 

= EXF-1933. 

Notes: De Vries (1952) synonymised the isolate identified as 

Hormodendrum langeronii with C. sphaerospermum. Strains of this 

species have often been identified as C. cladosporioides (Buzina 

et al. 2003, Menkis et al. 2005) although it has slightly longer 

conidia. 

174


Cladosporium psychrotolerans Zalar, de Hoog & Gunde- 

Cimerman, sp. nov. MycoBank MB492428. Fig. 10. 

Etymology: Refers to its ability to grow at low temperatures. 

Mycelium partim submersum; hyphae vagina polysaccharidica carentes. 

Conidiophora erecta vel adscendentia; stipes (10–)50–100(–150) × (3–)3.5–4(–7.5) 

μm, olivaceo-brunneus, levis, crassitunicatus, compluries regulariter septatus 

(cellulis 10–40 μm longis), identidem dichotome ramosus. Conidiorum catenae 

undique divergentes, terminales partes simplices ad 4 conidia continentes. Cellulae 

conidiogenae indistinctae. Ramoconidia primaria cylindrica, (18–)19–22(–43) × 

(2.5)3–3.5(–4.5) µm, 0(–1)-septata. Conidia leves vel leniter verruculosa, dilute 

brunnea, unicellularia, globosa vel ovoidea, (2.5–)3–4(–4.5) × (2–)2.5–3(–3) μm, 

long.: lat. 1.3–1.4; ramoconidia secundaria cylindrica, 0–1(–2)-septata, (5–)8–16(– 

36) × (2–)2.5–3(–5) μm, ad 4 cicatrices terminales ferentia; cicatrices inspissatae, 

conspicuae, 0.5–2 μm diam. 

Mycelium partly superficial partly submerged; hyphae without 

extracellular polysaccharide-like material. Conidiophores erect or 

ascending, macronematous, stipes (10–)50–100(–150) × (3–)3.5– 

4(–7.5) μm, olivaceous-brown, smooth or almost so, thick-walled, 

regularly septate (cell length 10–40 μm), arising laterally from 

aerial hyphae, repeatedly dichotomously branched. Conidial chains 


Ramoconidia sometimes formed, cylindrical, (18–)19–22(–43) × 

(2.5)3–3.5(–4.5) µm, aseptate, rarely 1-septate, with a broadly 

truncate base, up to 2 µm wide, unthickened or slightly thickened, 

somewhat darkened-refractive. Conidia smooth to minutely 

verruculose, light brown, non-septate, spherical to ovoid, length : 

width ratio = 1.3–1.4; conidial size (2.5–)3–4(–4.5) × (2–)2.5–3(–3) 


cylindrical, 0–1(–2)-septate, (5–)8–16(–36) × (2–)2.5–3(–5) μm 

[av. (± SD) 12.7 (± 6.5) × 3.0 (± 0.5)], with up to 4 distal scars. 

Conidiogenous scars thickened and conspicuous, protuberant, 

0.5–2 μm diam. 


diam, velvety, olive (3F4) due to profuse sporulation, flat with 

straight margin. Reverse dark green. Colonies on OA reaching 13– 

15 mm diam, olive (2F8), of granular appearance due to profuse 

sporulation; aerial mycelium sparse. Margin regular. Reverse black. 

Colonies on MEA reaching 8–15 mm diam, olive (2F4), velvety, 

radially furrowed with undulate white margin. Colonies on MEA 

with 5 % NaCl growing faster than on other media, reaching 25–27 

mm diam, olive (3E6) and granular due to profuse sporulation, 

either slightly furrowed or heavily wrinkled with regular or undulate 

margin. Reverse dark green. 

Maximum tolerated salt concentration: 17 % NaCl after 14 d. 

Cardinal temperatures: Minimum at 4 °C (5 mm diam), optimum 

and maximum at 25 °C (8–15 mm diam). 


and isol. S. Sonjak, May 1999, CBS H-19730, holotype, culture ex-type EXF-391 


Habitats and distribution: Hypersaline water in the Mediterranean 

basin. 

Differential parameters: Growth at 4 °C; maximal NaCl 

concentration 17 % NaCl, which differentiates it from other species 

with similar conidia, like C. sphaerospermum, C. halotolerans and 

C. dominicanum. 

Strains examined: EXF-326, EXF-332, EXF-391 (= CBS 119412; 

ex-type strain), EXF-714. 

Cladosporium salinae Zalar, de Hoog & Gunde-Cimerman, sp. 


Etymology: Refers to salterns (= Latin salinae) as the habitat of 

this species. 

Mycelium partim submersum; hyphae multa rostra lateralia ferentes, hyphae vagina 

polysaccharidica involutae. Conidiophora vix distincta, lateralia vel terminalia ex 

hyphis aeriis oriunda; stipes longitudine variabili, (5–)25–50(–60) × (2–)2.5–3(–4) 

μm, olivaceo-brunneus, levis vel leniter verruculosus, crassitunicatus, irregulariter 

dense septatus (cellulis 6–29 μm longis), simplex, interdum ramosus. Conidiorum 

catenae undique divergentes, terminales ad 6 conidia continentes. Cellulae 

conidiogenae nonnumquam integratae, in summo sequentiam sympodialem 

denticulorum formantes. Conidia levia, interdum leniter verruculosa, dilute brunnea, 

unicellularia, plerumque fusiformia, (4.5–)5.5–7.5(–10) × (2–)2.5–3(–3.5) μm, 

long. : lat. 1.9–2.4; ramoconidia secundaria cylindrica, 0–1(–2)-septata, (7.5–)9.5– 

13.5(–19) × (2.5–)2.5–3.5(–4.5) μm, ad 5 cicatrices terminales ferentia; cicatrices 

inspissatae, conspicuae, protuberantes, 0.7–1.8 μm diam. 

Mycelium partly superficial partly submerged, with numerous 

lateral pegs, consistently enveloped in polysaccharide-like 

material. Conidiophores poorly differentiated, micronematous, 

stipes (5–)25–50(–60) × (2–)2.5–3(–4) μm, olivaceous-brown, 

smooth to often minutely verruculose or irregularly rough-walled, 

thick-walled, irregularly densely septate (length of cells 6–29 μm), 

arising laterally and terminally from aerial hyphae, unbranched, 

occasionally branched. Conidial chains branching in all directions, 

terminal chains with up to 6 conidia. Conidiogenous cells sometimes 

integrated, producing sympodial clusters of pronounced denticles 

at their distal ends. Conidia usually smooth, occasionally minutely 

verruculose, light brown, aseptate, usually oblong ellipsoidal to 

fusiform, length : width ratio = 1.9–2.4; (4.5–)5.5–7.5(–10) × (2–) 

2.5–3(–3.5) μm [av. (± SD) 6.7 (± 1.3) × 2.9 (± 0.4)]; secondary 

ramoconidia cylindrical, 0–1(–2)-septate, (7.5–)9.5–13.5(–19) 

× (2.5–)2.5–3.5(–4.5) μm [av. (± SD) 12.1 (± 3.3) × 3.2 (± 0.6)], 

with up to 5 distal scars. Conidiogenous scars thickened and 

conspicuous, protuberant, 0.7–1.8 μm diam. 


diam, granular, olive (2E4) due to profuse sporulation, with white 

undulate margin. Aerial mycelium absent. Colonies either heaped 

or radially furrowed, in the marginal area growing deeply into the 

agar. Reverse dark brown to dark green. Colonies on OA reaching 

7–20 mm diam, olive (3E6), of granular appearance due to profuse 

sporulation, aerial mycelium present. Margin either undulate or 

arachnoid, deeply furrowed. Reverse pale brown to dark green. 

Colonies on MEA reaching 8–19 mm diam, velvety, reseda-green 

(2E6), heaped. Margin furrowed, growing deeply into the agar. 

Colonies on MEA with 5 % NaCl growing much faster than on 

other media, reaching 25–38 mm diam, of different colours, mostly 

reseda-green (2E6) and granulate due to profuse sporulation, 

margin olive-yellow (2D6). Reverse yellow to dark green. 

Maximum tolerated salt concentration: MEA + 17 % NaCl after 14 d. 

Cardinal temperatures: No growth at 4 °C, optimum and maximum 

temperature at 25 °C (8–19 mm diam), no growth at 30 °C. 


and isol. S. Sonjak, Feb. 1999, CBS H-19731, holotype, culture ex-type EXF-335 


Habitats and distribution: Hypersaline water in the Mediterranean 

basin. 

Differential parameters: Sympodial conidiogenous cells with 

pronounced denticles, narrow temperature amplitude. 


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Zalar et al. 

Fig. 10. Cladosporium psychrotolerans. Macro- and micromorphological characters. A–D. Colony surface grown on PDA (A), OA (B), MEA (C) and MEA plus 5 % NaCl (D) of 

strains incubated for 14 d at 25 ºC in darkness. E–F. Conidiophores. G. Apical part of a conidiophore. H–I. Secondary ramoconidia and conidia. E–I. All from 7-d-old SNA slide 

cultures. All but C, from EXF-391 (ex-type strain); C, from EXF-714. Scale bars A–D = 10 mm, E = 100 µm, F = 50 µm, G–I = 10 µm. 

Strains examined: EXF-322, EXF-335 (= CBS 119413; ex-type 

strain), EXF-604. 

Notes: Cladosporium salinae morphologically resembles species 

of the genus Fusicladium because its conidia are oblong ellipsoidal 

to fusiform and conidiogenous loci of ramoconidia are placed 

closely together. As any other Cladosporium species, its conidia 

show typical cladosporioid scar structures, however. Cladosporium 

salinae seems to have a separate position within the genus 

Cladosporium since it seems to be distantly related to any other 

described Cladosporium species or currently known species 

complex within Cladosporium. 

176


Fig. 11. Cladosporium salinae. Macro- and micromorphological characters. A–D. Colony surface grown on PDA (A), OA (B), MEA (C) and MEA plus 5 % NaCl (D) of strains 

incubated for 14 d at 25 ºC in darkness. E–F. Habit of conidiophores. G. Conidiophore. H. Detail of apical part of conidiophore. I. Conidia. J. Secondary ramoconidia and conidia. 

E–J. All from 7-d-old SNA slide cultures. A–D, from EXF-604; E–J, from EXF-335 (ex-type strain). Scale bars A–C = 5 mm, D = 10 mm, E = 100 µm, F = 50 µm, G = 30 µm, 

H–J = 10 µm. 

Cladosporium sphaerospermum Penzig, Michelia 2(8): 473. 

1882. Fig. 12. 

Mycelium partly submerged, partly superficial; hyphae thick, darkly 

pigmented and densely septate in submerged mycelium, not 

enveloped in polysaccharide-like material. Conidiophores erect or 

ascending, micronematous and macronematous, stipes of variable 

length, (10–)45–130(–300) × (2.5–)3–4(–6) μm, olivaceous-brown, 

smooth to minutely verruculose, thick-walled, with relatively 

dense septation (cells mostly 4.5–23 long), septa darkened and 


somewhat thickened, arising laterally and terminally from immersed 

or aerial hyphae, either unbranched or branched. Conidial chains 


Conidiogenous cells not differentiated. Ramoconidia often formed, 

cylindrical, (11.5–)20.5–40(–48) × (2.5–)3(–3.5) µm, with up to 5 

septa, base broadly truncate, 2 µm wide, slightly thickened and 

somewhat darkened-refractive. Conidia verruculose, brown to 

dark brown, non-septate, usually subspherical to spherical, less 

often short-ovoid, narrower at both ends, with length : width ratio 

= 1.1–1.5; conidial size (2.5–)3–4(–7) × (2–)3–3.5(–4.5) μm [av. (± 

177

Zalar et al. 

Fig. 12. Cladosporium sphaerospermum. Macro- and micromorphological characters. A–D. Colony surface grown on PDA (A), OA (B), MEA (C) and MEA plus 5 % NaCl (D) of 

strains incubated for 14 d at 25 ºC in darkness. E–F. Habit of conidiophores. G–I. Ramoconidia and conidia. E–I. All from 7-d-old SNA slide cultures. A, C–D, F–H, from CBS 

193.54 (ex-neotype strain); B, from EXF-738; E, EXF-455; I, EXF-458. Scale bars A–D = 10 mm; E = 100 µm; F = 50 µm; G–I = 10 µm. 

SD) 3.8 (± 0.8) × 3.1 (± 0.4)]; secondary ramoconidia cylindrical to 

almost spherical, 0–3(–4) septate, (4–)8.5–16(–37.5) × (2–)3–3.5(– 

5) μm [av. (± SD) 13.1 (± 6.3) × 3.2 (± 0.5)], with up to 4, rarely up 


protuberant, 0.9–1.1(–1.4) μm diam. 


velvety, olive (2F5) due to profuse sporulation, either with white 

and regular, or exceptionally undulate margin. Aerial mycelium 

sparse. Colonies flat or rarely radially furrowed with elevated 

colony centre. Exudates not prominent, some strains release 

green soluble pigments into the agar. Reverse blackish blue to pale 

green. Growth deep into the agar. Colonies on OA reaching 21– 

38 mm diam, olive (2F8), of granular appearance due to profuse 

and uniform sporulation, almost no aerial mycelium. Margin either 

regular or arachnoid, deeply radially furrowed. Reverse black. 

Colonies on MEA reaching 15–35 mm diam, velvety, linden-green 

(2C5), radially furrowed. Colony centre wrinkled, forming a craterlike 

structure; margin furrowed, lighter in colour, consisting of 

178


Fig. 13. Cladosporium spinulosum. Macro- and micromorphological characters. A–D. Colony surface grown on PDA (A), OA (B), MEA (C) and MEA plus 5 % NaCl (D) of strains 

incubated for 14 d at 25 ºC in darkness. E. Habit of conidiophores. F–J. Conidiophores. K–L. Conidia (also visible in I–J). E–L. All from 7-d-old SNA slide cultures. A–L, from 

EXF-334 (ex-type strain). Scale bars A–D = 10 mm, E = 100 µm, F = 30 µm, G–L = 10 µm. 

submerged mycelium. Reverse pale to dark brown. Colonies on 

MEA with 5 % NaCl growing faster than on other media, reaching 

31–60 mm diam, mainly olive (2D4), either being almost flat or 

radially furrowed, with margin of superficial mycelium; sporulation 

dense. Reverse ochraceous or dark green. 

Maximum tolerated salt concentration: On MEA + 20 % NaCl 89 % 

of all strains tested develops colonies after 7 d, 96 % after 14 d. 

Cardinal temperatures: No growth at 4 °C, optimum 25 °C (15–35 

mm diam), maximum 30 °C (2–15 mm diam). No growth at 37 °C. 

Specimen examined: Netherlands, from nail of man, 1949, coll. and isol. R.W. 

Zappey, CBS H-19738, neotype designated here, incorrectly selected by de Vries 

(1952) as “lectotype”, culture ex-neotype CBS 193.54 = ATCC 11289 = IMI 049637. 

Habitats and distribution: Hypersaline water in mediterranean 

and tropics; soil and plants in temperate climates; indoor wet 

cells; humans. The species does not seem to have any particular 

preference. Human isolates were probably culture contaminants. 

Literature: Penzig (1882), de Vries (1952), Ellis (1971), de Hoog et 

al. (2000), Samson et al. (2002). 

Diagnostic parameters: Thick-walled, melanised, densely septate 

mycelium, almost spherical, verruculose to verrucose terminal 

conidia, growth on 20 % NaCl after 7 d. 

Strains examined: CBS 109.14, CBS 122.63, CBS 190.54, CBS 

192.54, CBS 193.54 (ex-neotype strain), CBS 102045, CPC 10944, 

EXF-131, EXF-328, EXF-385, EXF-446, EXF-455, EXF-458, EXF- 

461, EXF-464, EXF-465, EXF-598, EXF-644, EXF-645, EXF-649, 

EXF-715, EXF-738, EXF-739, EXF-781, EXF-962, EXF-965, EXF- 

1061, EXF-1726, EXF-1732. 


179

Zalar et al. 

Fig. 14. Cladosporium velox. Macro- and micromorphological characters. A–D. Colony surface grown on PDA (A), OA (B), MEA (C) and MEA plus 5 % NaCl (D) of strains 

incubated for 14 d at 25 ºC in darkness. E–F. Habit of conidiophores. G. Conidiophore. H–I. Secondary ramoconidia and conidia. E–I. All from 7-d-old SNA slide cultures. A–D, 

G, from CBS 119417 (ex-type strain); E–F, H–I, from EXF-466. Scale bars A–D = 10 mm, E = 100 µm, F = 50 µm, G = 30 µm, H–I = 10 µm. 

Cladosporium spinulosum Zalar, de Hoog & Gunde-Cimerman, 


Etymology: Refers to its conspicuously digitate conidia. 

Conidiophora erecta vel adscendentia; stipites longitudine variabili, (15–)25–50(– 

155) × (2.5–)3–4(–5) μm, olivaceo-brunneus, levis, crassitunicatus, 0–6(–9)-septatus 

(cellulis 6–20 μm longis), ex hyphis submersis vel aeriis lateraliter vel terminaliter 

oriundus, simplex vel ramosus. Conidiorum catenae undique ramosae, ad 4 conidiis 

in partibus linearibus continuis cohaerentibus. Cellulae conidiogenae integratae vel 

discretae, acervos distales denticulorum conspicuorum sympodialium proferentes. 

Conidia echinulata vel digitata, brunnea vel fusca, continua, vulgo subglobosa vel 

globosa, (4.5–)5.5–7(–8) × (3–)4–4.5(–5) μm, long.: lat. = 1.1–1.6, digiti 0.6–1.3 

μm longi; ramoconidia secundaria etiam digitata, cylindrica vel subglobosa, 0(–1)- 

septata, (6–)6.5–8(–18) × (4–)4.5–5(–5.5) μm, 1–3 cicatrices distales ferentia. 

Cicatrices inspissatae, conspicuae, protuberantes, 0.8–1.2 μm diam. Hyphae 

nonnumquam polysaccharido circumdatae. 

Hyphae sometimes enveloped in polysaccharide-like material. 

Conidiophores erect or ascending, stipes of variable length, (15–) 

180


25–50(–155) × (2.5–)3–4(–5) μm, olivaceous-brown, smooth, 

sometimes irregularly rough-walled to verrucose near the base, 

thick-walled, 0–6(–9)-septate (cells mostly 6–20 μm long), arising 

laterally and terminally from immersed or aerial hyphae, either 

unbranched or branched, somewhat tapering towards the apex. 

Conidial chains branching in all directions, up to 4 conidia in the 

unbranched parts. Conidiogenous cells sometimes integrated, 

producing sympodial clusters of pronounced denticles at their distal 

ends. Ramoconidia rarely formed. Conidial wall ornamentation 

conspicuously digitate, with up to 1.3 μm long projections having 

parallel sides and blunt ends. Conidia brown to dark brown, 

aseptate, usually subspherical to spherical, length : width ratio = 

1.1–1.6; conidial size (4.5–)5.5–7(–8) × (3–)4–4.5(–5) μm [av. (± 

SD) 6.2 (± 1.0) × 4.2 (± 0.5)]; secondary ramoconidia ornamented 

as conidia, cylindrical to almost spherical, 0(–1)-septate, (6–)6.5– 

8(–18) × (4–)4.5–5(–5.5) μm [av. (± SD) 8.6 (± 4.0) × 4.8 (± 0.4)], 

with up to 3 distal scars. Conidiogenous scars thickened and 

conspicuous, protuberant, 0.8–1.2 μm diam. 


diam, velvety, dull green (29E4) to dark green (29F6) due to profuse 

sporulation, either with white and regular, or undulate margin. Aerial 

mycelium sparse. Colonies flat or radially furrowed with elevated 

colony centre. Growth deep into the agar. Exudates not prominent. 

Colonies on OA reaching 20–25 mm diam, dull green (29E4) to dark 

green (29F6), sometimes olive (3D4), of granular appearance due 

to profuse and uniform sporulation; almost without aerial mycelium. 

Margin arachnoid. Reverse pale brown to black. Colonies on MEA 

reaching 17–28 mm diam, velvety, dull green (29E4) to dark green 

(29F6), either flat or radially furrowed. Colony centre wrinkled, 

forming a crater-like structure; margin furrowed, paler in colour, 

consisting of submerged mycelium only. Reverse pale to dark 

green. Colonies on MEA with 5 % NaCl reaching 12–18 mm diam, 

of different colours, greenish grey (29D2), greyish green (29D5) 

to dark green (29F6); colony appearance variable, mostly either 

being almost flat with immersed colony centre or radially furrowed, 

with white to dark green margin consisting of superficial mycelium; 

sporulation dense. Reverse pale to dark green. 

Maximum tolerated salt concentration: On MEA + 17 % NaCl, two 

of three strains tested developed colonies after 14 d. 

Cardinal temperatures: Growth at 4 °C, optimum and maximum at 

25 °C (17–28 mm). No growth at 30 °C. 


and isol. S. Sonjak, Feb. 1999, CBS H-19796, holotype, culture ex-type EXF-334 


Habitats and distribution: Hypersaline water in temperate climate. 

Diagnostic parameters: Conidia and ramoconidia with a digitate 

ornamentation. 

Strains examined: EXF-334 (= CBS 119907; ex-type strain), EXF- 

382. 

Notes: Cladosporium spinulosum is a member of the C. herbarum 

species complex (Figs 2–4) although its globoid conidia are 

reminiscent of C. sphaerospermum. Within Cladosporium, the 

species is unique in having conspicuously digitate conidia and 

ramoconidia. The two strains are differing in the size of conidia. 

The average size of conidia in EXF-334 is 6.2 (± 0.9) × 4.2 (± 0.5) 

μm, and in EXF-382 it is 3.9 (± 0.6) × 3.3 (± 0.4) μm. 

Cladosporium velox Zalar, de Hoog & Gunde-Cimerman, sp. nov. 


Etymology: Refers to the quick growth of strains of this species. 

Mycelium partim submersum; hyphae vagina polysaccharidica carentes. 

Conidiophora erecta, lateralia vel terminalia ex hyphis aeriis oriunda; stipes (10–) 

25–150(–250) × (2.5–)3–4(–4.5) μm, olivaceo-brunneus, levis, crassitunicatus, ad 

7–septatus (cellulis 10–60 μm longis), identidem dichotome ramosus. Conidiorum 

catenae undique divergentes, terminales partes simplices ad 5 conidia continentes. 

Cellulae conidiogenae indistinctae. Conidia levia vel leniter verruculosa, dilute 

brunnea, unicellularia, ovoidea, (2–)3–4(–5.5) × (1.5–)2–2.5(–3) μm, long. : lat. 

1.4–1.7; ramoconidia secundaria cylindrica, 0–1-septata, (3.5–)5.5–19(–42) × 

(2–)2.5–3(–4.5) μm, ad 4(–5) cicatrices terminales ferentia; cicatrices inspissatae, 

protuberantes, conspicuae, 0.5–1.5 μm diam. 

Mycelium partly superficial partly submerged; hyphae without 

extracellular polysaccharide-like material. Conidiophores erect, 

stipes (10–)25–150(–250) × (2.5–)3–4(–4.5) μm, slightly attenuated 

towards the apex, olivaceous-brown, smooth- and thick-walled, 

arising terminally and laterally from aerial hyphae, dichotomously 

branched [up to 5(–7)-septate, cell length 10–60 μm]. Ramoconidia 

rarely formed. Conidial chains branching in all directions, terminal 

chains with up to 5 conidia. Conidia smooth to very finely 

verruculose, pale brown, non-septate, ovoid, length : width ratio 

= 1.4–1.7; (2–)3–4(–5.5) × (1.5–)2–2.5(–3) μm [av. (± SD) 3.6 (± 

0.6) × 2.3 (± 0.2)]; secondary ramoconidia cylindrical, 0–1-septate, 

(3.5–)5.5–19(–42) × (2–)2.5–3(–4.5) μm [av. (± SD) 13.4 (± 10.2) 

× 2.8 (± 0.5)], with up to 4(–5) distal scars. Conidiogenous scars 

thickened and conspicuous, protuberant, 0.5–1.5 μm diam. 


diam, velvety, dark green due to profuse sporulation, on some parts 

covered with white sterile mycelium, flat with straight white margin. 

Reverse dark green to black. Colonies on OA reaching 30–43 mm 

diam, dark green, mycelium submerged, aerial mycelium sparse. 

Margin regular. Reverse black. Colonies on MEA reaching 30–42 

mm diam, pale green, radially furrowed, with raised, crater-shaped 

central part, with white, undulate, submerged margin. Sporulation 

poor. Colonies on MEA with 5 % NaCl reaching 35–45 mm diam, 

pale green, velvety, flat with regular margin. Reverse pale green. 

Sporulation poor. 

Maximum tolerated salt concentration: 20 % NaCl after 14 d. 

Cardinal temperatures: Minimum at 10 °C (9 mm diam), optimum at 

25 °C (30–42 mm diam) and maximum at 30 °C (5–18 mm diam). 

Specimen examined: India, Charidij, isolated from Bambusa sp., W. Gams, CBS 

H-19735, holotype, culture ex-type CBS 119417. 

Habitats and distribution: Hypersaline water in Slovenia; bamboo, 

India. 

Strains examined: CBS 119417 (ex-type strain), EXF-466, EXF- 

471. 


The authors are grateful to Walter Gams for his comments on the manuscript 

and for providing Latin diagnoses, and to Konstanze Schubert for contributions to 

species descriptions. We thank Kazimir Drašlar and Marko Lutar for preparing SEM 

illustrations, and Kasper Luijsterburg, Špela Štrekelj and Barbara Kastelic-Bokal for 

their technical assistance. The research was partially supported by the European 

Union-funded Integrated Infrastructure Initiative grant SYNTHESYS Project NL- 

TAF-1070. 


181

Zalar et al. 

REFERENCES 

Aihara M, Tanaka T, Ohta T, Takatori K (2002). Effect of temperature and water 

activity on the growth of Cladosporium sphaerospermum and Cladosporium 

cladosporioides. Biocontrol Science 7: 193–196. 

Aihara M, Tanaka T, Takatori K (2001). Cladosporium as the main fungal contaminant 

of locations in dwelling environments. Biocontrol Science 6: 49–52. 



Arx JA von (1950). Über die Ascusform von Cladosporium herbarum (Pers.) Link. 

Sydowia 4: 320–324. 

Badillet G, Bièvre C de, Spizajzen S (1982). Isolement de dématiées à partir d’ongles 

et de squames. Bulletin de la Société Française de Mycologie 11: 69–72. 




Butinar L, Sonjak S, Zalar P, Plemenitaš A, Gunde-Cimerman N (2005). Melanized 

halophilic fungi are eukaryotic members of microbial communities in hypersaline 

waters of solar salterns. Botanica Marina 48: 73–79. 

Buzina W, Braun H, Freudenschuss K, Lackner A, Stammberger H (2003). Fungal 

biodiversity as found in nasal mucus. Medical Mycology 41: 149–161. 

Carbone I, Kohn LM (1999). A method for designing primer sets for speciation 

studies in filamentous ascomycetes. Mycologia 91: 553–556. 

Clements FE, Shear CL (1931). The genera of fungi. New York: H.W. Wilson Co. 

Crous PW, Braun U, Groenewald JZ (2007a). Mycosphaerella is polyphyletic. 

Studies in Mycology 58: 1-32. 

Crous PW, Groenewald JZ, Pongpanich K, Himaman W, Arzanlou M, Wingfield MJ 

(2004). Cryptic speciation and host specificity among Mycosphaerella spp. 

occurring on Australian Acacia species grown as exotics in the tropics. Studies 

in Mycology 50: 457–469. 


JZ (2007b). Opportunistic, human-pathogenic species in the Herpotrichiellaceae 


Venturiaceae. Studies in Mycology 58: 185-217. 

Crous PW, Verkley GJM, Groenewald JZ (2006). Eucalyptus microfungi known from 

culture. 1. Cladoriella and Fulvoflamma genera nova, with notes on some other 

poorly known taxa. Studies in Mycology 55: 53–63. 

Curtis MD, Gore J, Oliver RP (1994). The phylogeny of the tomato leaf mould fungus 

Cladosporium fulvum syn. Fulvia fulva by analysis of rDNA sequences. Current 

Genetics 25: 318–322. 





Fonseca FOOR da, Area Leão AE de, Peñido NJJC (1927a). Mycose de type 

ulcéro-nodulaire, semblable à la sporotrichose et produite par Hormodendrum 

langeronii. Comptes-rendus des Séances de la Société de Biologie 97: 1772– 

1774. 

Fonseca FOOR da, Area Leão AE de, Peñido NJJC (1927b). Mycose de type 

ulcéro-nodulaire, semlable à la sporotrichose et produite par Hormodendrum 

langeronii. Sciencia Medica 5: 563–573. 

Gams W, Verkleij GJM, Crous PW (2007). CBS Course of Mycology, 5 th ed. 

Centraalbureau voor Schimmelcultures, Utrecht, Netherlands. 


of the neurotropic species Cladophialophora bantiana. Studies in Mycology 43: 

151–162. 

Gunde-Cimerman N, Zalar P, Hoog GS de, Plemenitaš A (2000). Hypersaline 

water in salterns – natural ecological niches for halophilic black yeasts. FEMS 

Microbiology Ecology 32: 235–240. 

Haubold EM, Aronson JF, Cowan DF, McGinnis MR, Cooper CR (1998). Isolation 

of fungal rDNA from bottlenose dolphin skin infected with Loboa loboi. Medical 

Mycology 36: 263–267. 

Herr RA, Tarcha EJ, Taborda PR, Taylor JW, Ajello L, Mendoza L (2001). 

Phylogenetic analysis of Lacazia loboi places this previously uncharacterised 

pathogen within the dimorphic Onygenales. Journal of Clinical Microbiology 

39: 309–314. 

Ho MHM, Castañeda RF, Dugan FM, Jong SC (1999). Cladosporium and 


72: 115–157. 

Hocking AD, Miscamble BF, Pitt J (1994). Water relations of Alternaria alternata, 

Cladosporium cladosporioides, Cladosporium sphaerospermum, Curvularia 

lunata and Curvularia pallescens. Mycological Research 98: 91–94. 



Hoog GS de, Guarro J, Gené J, Figueras MA (2000). Atlas of Clinical Fungi, 2 nd ed. 

Centraalbureau voor Schimmelcultures, Utrecht / Universitat Rovira i Virgili, 

Reus. 

Hoog GS de, Zalar P, Urzì C, Leo F de, Yurlova NA, Sterflinger K (1999). Relationships 

of dothideaceous black yeasts and meristematic fungi based on 5.8S and ITS2 

rDNA sequence comparison. Studies in Mycology 43: 31–37. 

Jeannmougin F, Thompson JD, Gouy M, Higgins DG, Gibson TJ (1998). Multiple 

sequence alignment with ClustalX. Trends in Biochemical Sciences 23: 403– 

405. 

Kantarcioglu AS, Yucel A, Hoog GS de (2002). Case report. Isolation of Cladosporium 

cladosporioides from cerebrospinal fluid. Mycoses 45: 500–503. 

Kirk P, Cannon P, David J, Stalpers J (2001). Ainsworth and Bisby’s Dictionary of the 

Fungi. 9 th edn. Wallingford, UK. CAB International. 

Kornerup A, Wanscher JH (1967). Methuen Handbook of Colour. 2 nd edn. Methuen 

Co., London. 

Kumar S, Tamura K, Nei M (2004). MEGA3: integrated software for molecular 

evolutionary genetics analysis and sequence alignment. Briefings in 

Bioinformatics 5: 150–163. 

Lacaz CS (1996). Paracoccidioides loboi (Fonseca Filho et Arêa Leão, 1940) 

Almeida et Lacaz, 1948–1949. Description of the fungus in Latin. Revista 

Instituto Medicina Tropical São Paulo 38: 229–231. 

Link HF (1816). Observationes in ordines plantarum naturales III. Magazin der 

Gesellschaft Naturforschender Freunde Berlin 7: 37–38. 

Masclaux F, Guého E, Hoog GS de, Christen R (1995). Phylogenetic relationships of 

human-pathogenic Cladosporium (Xylohypha) species inferred from partial LS 

rRNA sequences. Journal of Medical and Veterinary Mycology 33: 327–338. 

McKemy JM & Morgan-Jones G (1991). Studies in the genus Cladosporium sensu 

lato. V. Concerning the type species, Cladosporium herbarum. Mycotaxon 42: 

307–317. 

Meklin T, Haugland RA, Reponen T, Varma M, Lummus Z, Bernstein D, Wymer 

LJ, Vesper SJ (2004). Quantitative PCR analysis of house dust can reveal 

abnormal mold conditions. Journal of Environmental Monitoring 6: 615–620. 

Menkis A, Vasiliauskas R, Taylor AFS, Stenlid J, Finlay R (2005). Fungal 

communities in mycorrhizal roots of conifer seedlings in forest nurseries under 

different cultivation systems, assessed by morphotyping, direct sequencing 

and mycelial isolation. Mycorrhiza 16: 33–41. 

Moricca S, Ragazzi A, Mitchelson KR (1999). Molecular and conventional detection 

and identification of Cladosporium tenuissimum on two-needle pine rust 

aeciospores. Canadian Journal of Botany 77: 339–347. 

O’Donnell K, Cigelnik E (1997). Two divergent intragenomic rDNA ITS2 types within 

a monophyletic lineage of the fungus Fusarium are nonorthologous. Molecular 

Phylogenetic Evolution 7: 103–116. 

Park HG, Managbanag JR, Stamenova EK, Jong SC (2004). Comparative analysis 

of common indoor Cladosporium species based on molecular data and conidial 

characters. Mycotaxon 89: 441–451. 

Penzig AJO (1882). Fungi Agromiculi. Michelia 2: 385–503. 

Pereira PT, Carvalho MM de, Girio FM, Roseiro JC, Amaral-Collaco MT (2002). 

Diversity of microfungi in the phylloplane of plants growing in a Mediterranean 

ecosystem. Journal of Basic Microbiology 42: 396–407. 

Phillips G, Hudson S, Stewart WK (1992). Microbial growth and blockage of subfloor 

drains in a renal dialysis centre: a problem highlighted. Journal of Hospital 

Infection 21: 193–198. 

Pitt JI (1979). The Genus Penicillium and its teleomorphic states Eupenicillium and 

Talaromyces. Academic Press, London. 



Samson RA, Hoekstra ES, Frisvad JC, Filtenborg O (2002). Introduction to Foodand 

Airborne Fungi, 6 th ed. Centraalbureau voor Schimmelcultures, Utrecht. 

Schoch C, Shoemaker RA, Seifert, KA, Hambleton S, Spatafora JW, Crous PW 


Mycologia 98: 1043–1054. 



(Davidiellaceae, Capnodiales), with standardisation of methods for 





Summerbell RC, Cooper E, Bunn U, Jamieson F, Gupta AK (2005). Onychomycosis: 

a critical study of techniques and criteria confirming the etiologic significance of 

nondermatophytes. Medical Mycology 43: 39–59. 

Swofford DL (2003). PAUP*. Phylogenetic Analysis Using Parsimony (*and Other 


Trejos A (1954). Cladosporium carrionii n. sp. and the problem of cladosporia 

isolated from chromoblastomycosis. Revista de Biología Tropical 2: 75–112. 


ex Fr. Baarn, 121 pp. Dissertation, University of Utrecht. 

Vuillemin P (1931). Les champignons parasites et les mycoses de l’homme. 

Encyclopédie Mycologique 2: 1–290. 

182






Wirsel SGR, Runge-Froböse C, Ahren DG, Kemen E, Oliver RP, Mendgen, 

KW (2002). Four or more species of Cladosporium sympatrically colonize 

Phragmites australis. Fungal Genetics and Biology 25: 99–113. 

Yano S, Koyabashi K, Kato K (2002). Intrabronchial lesion due to Cladosporium 

sphaerospermum in a healthy, non-asthmatic woman. Mycoses 46: 348–350. 


183


doi:10.3114/sim.2007.58.07 


Opportunistic, human-pathogenic species in the Herpotrichiellaceae are 

phenotypically similar to saprobic or phytopathogenic species in the 

Venturiaceae 

P.W. Crous 1* , K. Schubert 2 , U. Braun 3 , G.S. de Hoog 1 , A.D. Hocking 4 , H.-D. Shin 5 and J.Z. Groenewald 1 

1 

CBS Fungal Biodiversity Centre, P.O. Box 85167, 3508 AD, Utrecht, The Netherlands; 2 Botanische Staatssammlung München, Menzinger Strasse 67, D-80638 München, 

Germany; 3 Martin-Luther-Universität, Institut für Biologie, Geobotanik und Botanischer Garten, Herbarium, Neuwerk 21, D-06099 Halle, Germany; 4 CSIRO Food Science 

Australia, 11 Julius Avenue, North Ryde, NSW 2113, Australia; 5 Division of Environmental Science & Ecological Engineering, Korea University, Seoul 136-701, Korea 


Abstract: Although morphologically similar, species of Cladophialophora (Herpotrichiellaceae) were shown to be phylogenetically distinct from Pseudocladosporium 

(Venturiaceae), which was revealed to be synonymous with the older genus, Fusicladium. Other than being associated with human disorders, species of Cladophialophora 

were found to also be phytopathogenic, or to occur as saprobes on organic material, or in water, fruit juices, or sports drinks, along with species of Exophiala. Caproventuria 

and Metacoleroa were confirmed to be synonyms of Venturia, which has Fusicladium (= Pseudocladosporium) anamorphs. Apiosporina, based on A. collinsii, clustered 

basal to the Venturia clade, and appears to represent a further synonym. Several species with a pseudocladosporium-like morphology in vitro represent a sister clade to the 

Venturia clade, and are unrelated to Polyscytalum. These taxa are newly described in Fusicladium, which is morphologically close to Anungitea, a heterogeneous genus with 

unknown phylogenetic affinity. In contrast to the Herpotrichiellaceae, which were shown to produce numerous synanamorphs in culture, species of the Venturiaceae were 

morphologically and phylogenetically more uniform. Several new species and new combinations were introduced in Cladophialophora, Cyphellophora (Herpotrichiellaceae), 

Exophiala, Fusicladium, Venturia (Venturiaceae), and Cylindrosympodium (incertae sedis). 

Taxonomic novelties: Cladophialophora australiensis Crous & A.D. Hocking, sp. nov., Cladophialophora chaetospira (Grove) Crous & Arzanlou, comb. nov., Cladophialophora 

hostae Crous, U. Braun & H.D. Shin, sp. nov., Cladophialophora humicola Crous & U. Braun, sp. nov., Cladophialophora potulentorum Crous & A.D. Hocking, sp. nov., 

Cladophialophora scillae (Deighton) Crous, U. Braun & K. Schub., comb. nov., Cladophialophora sylvestris Crous & de Hoog, sp. nov., Cylindrosympodium lauri Crous & 

R.F. Castañeda, sp. nov., Cyphellophora hylomeconis Crous, de Hoog & H.D. Shin, sp. nov., Exophiala eucalyptorum Crous, sp. nov., Fusicladium africanum Crous, sp. nov., 

Fusicladium amoenum (R.F. Castañeda & Dugan) Crous, K. Schub. & U. Braun, comb. nov., Fusicladium brevicatenatum (U. Braun & Feiler) Crous, U. Braun & K. Schub., 

comb. nov., Fusicladium fagi Crous & de Hoog, sp. nov., Fusicladium intermedium (Crous & W.B. Kendr.) Crous, comb. nov., Fusicladium matsushimae (U. Braun & C.F. Hill) 

Crous, U. Braun & K. Schub., comb. nov., Fusicladium pini Crous & de Hoog, sp. nov., Fusicladium ramoconidii Crous & de Hoog, sp. nov., Fusicladium rhodense Crous & M.J. 

Wingf., sp. nov., Venturia hystrioides (Dugan, R.G. Roberts & Hanlin) Crous & U. Braun, comb. nov. 

Key words: Anungitea, Anungitopsis, Cladophialophora, Exophiala, Fusicladium, phylogeny, Pseudocladosporium, systematics, Venturia. 

Introduction 

Species of Cladophialophora Borelli are relatively simple 

hyphomycetes with brown hyphae that give rise to branched chains 

of pale brown conidia. Phylogenetically they are defined to belong to 

the Chaetothyriales (Haase et al. 1999, Untereiner 2000), an order 

containing numerous opportunists (de Hoog et al. 2000); teleomorph 

relationships are with Capronia Sacc. in the Herpotrichiellaceae. 

In several cases cladophialophora-like synanamorphs are found 

accompanying black yeasts of the genus Exophiala J.W. Carmich. 

(de Hoog et al. 1995). Braun & Feiler (1995) placed several saprobic 

hyphomycetes in Cladophialophora, and described Capronia 

hanliniana U. Braun & Feiler as teleomorph of Cladophialophora 

brevicatenata U. Braun & Feiler. This work was continued by Dugan 

et al. (1995), who described an additional teleomorph, Capronia 

hystrioides Dugan, R.G. Roberts & Hanlin for Cladophialophora 

hachijoensis (Matsush.) U. Braun & Feiler. Untereiner (1997) 

reduced Capronia hystrioides to synonymy with C. hanliniana, 

and placed them in Venturia Sacc. (Venturiaceae, Pleosporales). 

The concept of Cladophialophora hachijoensis, which is based 

on Phaeoramularia hachijoensis Matsush. (Matsushima 1975) is 

confused, however, and phylogenetic studies have revealed that 

isolates attributed to this name in recent studies, were in fact 

representatives of three different species in phylogenetically distinct 

genera (Braun et al. 2003). The separation of Cladophialophora 

with Capronia teleomorphs (Herpotrichiellaceae, Chaetothyriales; 

commonly isolated as human pathogens), from predominantly 

saprobic or phytopathogenic isolates in the Dothideomycetes 

was recognised by Braun (1998). Recently the cactus endophyte 

Cladophialophora yegresii de Hoog was reported to be the nearest 

neighbour of C. carrionii (Trejos) de Hoog et al., a major agent 

of human chromoblastomycosis (de Hoog et al. 2007), so that 

the main distinction between the two anamorph genera remains 

in their phylogenetic positions. Capronia hanliniana and C. 

hystrioides were again recognised as distinct species, and placed 

in a new genus, Caproventuria U. Braun (Venturiaceae), while their 

anamorphs were accommodated in Pseudocladosporium U. Braun. 

Caproventuria was primarily distinguished from Venturia based on 

its distinct Pseudocladosporium anamorphs. Recently, Crous et al. 

(2007b) introduced a third genus, namely Sympoventuria Crous & 

Seifert, which produces a sympodiella-like anamorph in culture. To 

complicate matters further, Beck et al. (2005) concluded, based on 

an ITS DNA phylogeny, that the morphology attributed to the form 

genera Spilocaea Fr., Pollaccia E. Bald. & Cif., and Fusicladium 

Bonord. has evolved several times within Venturia, and that a single 

anamorph genus should be used for Venturia, namely Fusicladium 

(see Schubert et al. 2003 for additional generic synonyms). 

In their treatment of Venturia anamorphs, Schubert et al. 

(2003) excluded Pseudocladosporium, and stated that its status 

needs to be confirmed along with that of other genera such as 

Anungitea B. Sutton, Fusicladium and Polyscytalum Riess. In the 

study by Beck et al. (2005) an isolate of Caproventuria hystrioides 

(Pseudocladosporium sp.) was included to confirm the link to 

the Venturiaceae, though this was not well resolved, nor was the 

status of the older generic names mentioned above addressed. 

185

Crous et al. 

The aim of the present study, therefore, was to use DNA sequence 

comparisons in conjunction with morphology in an attempt to clarify 

these generic issues, as well as to determine which morphological 

characters could be used to distinguish Pseudocladosporium from 

Cladophialophora. 


Isolates 

Cultures were obtained from the Centraalbureau voor 

Schimmelcultures (CBS) in Utrecht, the Netherlands, or isolated 

from plant material incubated in moist chambers to promote 

sporulation. Isolates were cultured on 2 % malt extract plates 

(MEA; Gams et al. 2007), by obtaining single conidial colonies as 

explained in Crous (2002). Colonies were subcultured onto fresh 

MEA, oatmeal agar (OA), potato-dextrose agar (PDA) and synthetic 

nutrient-poor agar (SNA) (Gams et al. 2007), and incubated at 25 

°C under continuous near-ultraviolet light to promote sporulation. 

DNA extraction, amplification and phylogeny 


DNA was isolated following the CTAB-based protocol described 

in Gams et al. (2007). The primers V9G (de Hoog & Gerrits van 

den Ende 1998) and LR5 (Vilgalys & Hester 1990) were used to 

amplify part (ITS) of the nuclear rDNA operon spanning the 3’ end 

of the 18S rRNA gene (SSU), the first internal transcribed spacer 

(ITS1), the 5.8S rRNA gene, the second ITS region and the 5’ 

end of the 28S rRNA gene (LSU). Four internal primers, namely 

ITS4 (White et al. 1990), LR0R (Rehner & Samuels 1994), LR3R 

(www.biology.duke.edu/fungi/mycolab/primers.htm), and LR16 

(Moncalvo et al. 1993), were used for sequencing to ensure good 

quality overlapping sequences were obtained. The PCR conditions, 

sequence alignment and subsequent phylogenetic analysis followed 

the methods of Crous et al. (2006a). The ITS1, ITS2 and 5.8S rRNA 

gene were only sequenced for isolates of which these data were 

not available. The ITS data were not included in the analyses but 

deposited in GenBank where applicable. Gaps longer than 10 


the remaining gaps were treated as missing data. Sequence data 

were deposited in GenBank (Table 1) and alignments in TreeBASE 

(www.treebase.org). 

Taxonomy 

Structures were mounted in lactic acid, and 30 measurements 

(× 1 000 magnification) determined wherever possible, with the 

extremes of spore measurements given in parentheses. Colony 

colours (surface and reverse) were assessed after 2–4 wk on OA 

and PDA at 25 °C in the dark, using the colour charts of Rayner 

(1970). All cultures obtained in this study are maintained in the CBS 

collection (Table 1). Nomenclatural novelties and descriptions were 

deposited in MycoBank (www.MycoBank.org). 

Results 

DNA phylogeny 

Amplicons of approximately 1 700 bases were obtained for the 

isolates listed in Table 1. These sequences were used to obtain 

additional sequences from GenBank which were added to the 

alignment. The manually adjusted LSU alignment contained 116 

sequences (including the two outgroup sequences) and 1 157 

characters including alignment gaps (available in TreeBASE). 

Of the 830 characters used in the phylogenetic analysis, 326 

were parsimony-informative, 79 were variable and parsimonyuninformative, 

and 425 were constant. Neighbour-joining analyses 

using three substitution models on the sequence data yielded trees 

with identical topologies to one another. The neighbour-joining trees 

support the same clades as obtained from the parsimony analysis, 

but with a different arrangement at the deep nodes, for example, 

the clade containing Protoventuria alpina (Sacc.) M.E. Barr (CBS 

140.83) is placed as sister to the Venturiaceae using parsimony but 

basal to the Herpotrichiellaceae using neighbour-joining. Because 

of the large number of different strain associations in the Venturia 

clade (see the small number of strict consensus branches for this 

clade in Fig. 1), only the first 5 000 equally most parsimonious trees 

(TL = 1 752 steps; CI = 0.392; RI = 0.849; RC = 0.333) were saved, 

one of which is shown in Fig. 1. 

Bayesian analysis was conducted on the same aligned LSU 

data set using a general time-reversible (GTR) substitution model 

with inverse gamma rates and dirichlet base frequencies. The 

Markov Chain Monte Carlo (MCMC) analysis of 4 chains started 

from a random tree topology and lasted 2 000 000 generations. 

Trees were saved each 1 000 generations, resulting in 2 000 saved 

trees. Burn-in was set at 500 000 generations after which the 

likelihood values were stationary, leaving 1 500 trees from which 

the consensus tree (Fig. 2) and posterior probabilities (PP’s) were 

calculated. The average standard deviation of split frequencies 

was 0.06683 at the end of the run. The same overall topology as 

that observed using parsimony was obtained, with the exception 

of the position of Anungitopsis speciosa R.F. Castañeda & W.B. 

Kendr., which is placed between the Leotiomycetes and the 

Sordariomycetes based on the Bayesian analysis. Also, similar to 

the results obtained using neighbour-joining, the clade containing 

Protoventuria alpina (CBS 140.83) is placed as sister to the 

Herpotrichiellaceae and not to the Venturiaceae. The phylogenetic 

affinity of specific genera or species are discussed below. 

Taxonomy 

Several collections represented novel members of the 

Herpotrichiellaceae and Venturiaceae, and these are described 

below. Taxa that were cladophialophora- or pseudocladosporiumlike, 

but that clustered elsewhere, are treated under excluded 

species. 

Members of Chaetothyriales, Herpotrichiellaceae 

Cladophialophora australiensis Crous & A.D. Hocking, sp. nov. 


Etymology: Named after its country of origin, Australia. 

Cladophialophorae carrionii similis, sed conidiis secundis majoribus, (7–)8–12(–15) 

× 3–4 µm. 

In vitro: Mycelium consisting of branched, septate, smooth, pale 

brown, guttulate, 2–3 µm wide hyphae; hyphal coils not seen. 

Conidiophores dimorphic; macroconidiophores mononematous, 

subcylindrical, multi-septate, straight to curved, up to 150 µm long 

(including conidiogenous cells), and 4 µm wide, pale to medium 

brown, smooth, guttulate; microconidiophores integrated with 

hyphae, which terminate in subcylindrical conidiogenous cells that 

give rise to branched chains of conidia; conidiophores (including 

186

Herpotrichiellaceae and Venturiaceae 

conidiogenous cells) up to 5-septate, 50 µm long, with terminal and 

lateral conidiogenous cells. Conidiogenous cells pale to medium 

brown, smooth, guttulate, terminal and lateral, subcylindrical, 

20–35 × 2–3.5 µm, or reduced to indistinct subtruncate to truncate 

loci, scars up to 2 µm wide, mono- to polyblastic, proliferating 

sympodially, scars neither darkened, thickened, nor refractive. 

Conidia pale to medium brown, guttulate, smooth; ramoconidia 

subcylindrical, 0–1-septate, 20–35 × 2–3 µm, hila subtruncate, 

inconspicuous, up to 2 µm wide, giving rise to branched chains 

of conidia; conidia ellipsoid, pale brown, but becoming dark brown 

and thick-walled in older cultures, guttulate, tapering towards 

subtruncate terminal loci, 0–1-septate, occurring in chains of up to 

20 conidia, (7–)8–12(–15) × 3–4 µm (older, dark brown conidia are 

ellipsoid, up to 5 µm wide). 

Cultural characteristics: Colonies erumpent, somewhat spreading, 

margins crenate, feathery, aerial mycelium sparse; colonies on 

PDA olivaceous-grey to iron-grey (surface); reverse iron-grey; on 

OA and SNA olivaceous-grey. Colonies reaching 5 mm diam after 2 

wk at 25 °C in the dark; colonies fertile. Not able to grow at 37 °C. 

Specimen examined: Australia, isolated from apple juice, Dec. 1986, A.D. Hocking, 

holotype CBS H-19899, culture ex-type CBS 112793 = CPC 1377. 

Notes: Cladophialophora australiensis is one of two novel species 

of Cladophialophora originally isolated from sports drinks in 

Australia. Cladophialophora spp. are commonly associated with 

human disorders (Honbo et al. 1984, de Hoog et al. 2000, Levin 

et al. 2004), and thus their occurrence in sports drinks is cause for 

concern. However, none of the new species described here had 

the ability to grow at 37 °C, and therefore it is not expected that 

they could pose a danger to humans. Comparing ITS diversity, the 

species shows more than 12 % difference to established pathogens 

such as C. carrionii and C. bantiana (Sacc.) de Hoog et al. 

Cladophialophora chaetospira (Grove) Crous & Arzanlou, comb. 


Basionym: Septocylindrium chaetospira Grove, J. Bot. Lond. 24: 

199. 1886. 

≡ Septonema chaetospira (Grove) S. Hughes, Naturalist, London 840: 9. 

1952. 

≡ Heteroconium chaetospira (Grove) M.B. Ellis, in Ellis, More Dematiaceous 


In vitro: Mycelium consisting of branched, septate, smooth, 

medium brown hyphae, 2–3.5 µm wide. Conidiophores reduced 

to conidiogenous cells, or a single supporting cell, 20–40 × 3–4 

µm. Conidiogenous cells subcylindrical, erect, straight to irregularly 

curved, medium brown, smooth, 15–30 × 3–4 µm. Conidia in 

branched, acropetal chains with up to 30 conidia; subcylindrical to 

fusiform, medium brown, smooth, tapering slightly at subtruncate 

ends, 1(–3)-septate, thin-walled, becoming slightly constricted at 

septa of older conidia, (20–)25–30(–45) × 3–4(–5) µm; conidia 

remaining attached in long chains; hila neither thickened, nor 

darkened-refractive. 

Cultural characteristics: Colonies erumpent, convex, spreading, 

with sparse to dense aerial mycelium; margins smooth, undulate; 

on PDA iron-grey (surface), margins olivaceous-black; reverse 

olivaceous-black; on OA olivaceous-grey in the middle due to fluffy 

aerial mycelium, iron-grey in wide outer margin; on SNA olivaceousgrey. 

Colonies reaching 12 mm diam after 2 wk on PDA at 25 °C in 

the dark. Not able to grow at 37 °C. 

Specimens examined: China, Yunnan, Yiliang, isolated from Phyllostachys 

bambusoides (Gramineae), decaying bamboo, freshwater, 6 Jul. 2003, L. Cai, CBS 

114747; China, Yunnan, stream in Kunming, isolated from bamboo wood, 15 Jun. 

2003, C. Lei, CBS 115468. Denmark, isolated from roots of Picea abies (Pinaceae), 

isol. by D.S. Malla, CBS 491.70. Germany, Schleswig-Holstein, Kiel-Kitzeberg, 

isolated from wheat field soil, isol. by W. Gams, CBS 514.63 = ATCC 16274 = MUCL 

8310. 

Notes: Two cultures of Heteroconium chaetospira were originally 

deposited as Spadicoides minuta L. Cai, McKenzie & K.D. Hyde (Cai 

et al. 2004), but later found to represent Heteroconium chaetospira, 

a species commonly found on rotting wood in Europe (Ellis 1976). 

The genus Heteroconium Petr. has in recent years been used 

to name leaf spotting fungi with chains of brown, disarticulating 

conidia (Crous et al. 2006b), which have phylogenetic affinities 

to several orders, obviously being polyphyletic. The type species 

of Heteroconium, H. citharexyli Petr., is a plant pathogen on 

Cytharexylum (Petrak 1949) with hitherto unknown phylogenetic 

position. The fact that H. chaetospira is linked to the Chaetothyriales, 

was rather unexpected. The species appears to be similar to others 

placed in Cladophialophora by having short, lateral conidiogenous 

cells, and long chains of branched subcylindrical conidia that largely 

remain attached. It is, however, quite distinct from other members of 

Cladophialophora in having medium brown conidia, and in lacking 

the ellipsoid conidia observed in several species. 

Cladophialophora hostae Crous, U. Braun & H.D. Shin, sp. nov. 


Etymology: Epithet derived from the host genus, Hosta. 

Cladophialophorae scillae similis, sed conidiophoris in vitro brevioribus et leniter 

angustioribus, 10–15 × 1.5–2 µm, conidiis brevioribus, (7–)10–15(–20) µm. 

In vivo: Leaf spots amphigenous, subcircular to somewhat angularirregular, 

1–5 mm wide, scattered to aggregated, sometimes 

confluent, pale to medium brown or with a reddish brown tinge, 

later greyish brown, margin indefinite or on the upper leaf surface 

with a narrow slightly raised marginal line or very narrow lighter 

halo, yellowish, ochraceous to brownish. Caespituli epiphyllous, 

punctiform to confluent, dingy greyish brown. Mycelium immersed, 

forming fusicladium-like hyphal strands or plates; hyphae septate, 

sometimes with constrictions at the septa, thin-walled, pale 

olivaceous, 1.5–7 µm wide. Stromata immersed, small, 10–40 µm 

diam, composed of swollen hyphal cells, subcircular to somewhat 

angular-irregular in outline, 2–8 µm diam, wall somewhat 

thickened, brown. Conidiophores in small to moderately large 

fascicles, loose, divergent to moderately dense, rarely solitary, 

arising from stromatic hyphal aggregations, erumpent, erect, 

usually unbranched, rarely branched, straight, subcylindrical to 

distinctly geniculate-sinuous, 5–40 × 2–5 µm, 0–6-septate, pale 

to medium olivaceous to olivaceous-brown, thin-walled, up to 0.5 

µm, smooth. Conidiogenous cells integrated, terminal, 5–15(–20) 

µm long, sympodial, conidiogenous loci rather inconspicuous 

to subdenticulate, flat-tipped, 1–1.5 µm diam, unthickened or 

almost so, not to slightly darkened-refractive. Conidia in simple or 

branched chains, narrowly ellipsoid-subcylindrical, 10–15 × 1.5– 

3.5 µm, 0–1-septate, subhyaline to pale olivaceous, thin-walled, 

smooth, ends truncate or with two denticle-like hila in ramoconidia, 

(0.75–)1–1.5(–2) µm diam, unthickened or almost so, at most 

slightly darkened-refractive. 

In vitro: Mycelium composed of branched, smooth, pale olivaceous 

to medium brown hyphae, frequently forming hyphal coils, guttulate, 

septa inconspicuous, not constricted, hyphae somewhat irregular 

in width, 1–2 µm wide. Conidiophores reduced to conidiogenous 

cells, integrated in hyphae, terminal, subcylindrical, pale olivaceous 

to pale brown, smooth, 0–1-septate, proliferating sympodially at 


187

Crous et al. 

10 changes 



100 


Fasciatispora petrakii AY083828 

Pidoplitchkoviella terricola AF096197 

Anungitopsis speciosa CBS 181.95 

100 

94 100 

98 

81 97 

90 

90 

87 

53 

100 

94 

100 

Polyscytalum fecundissimum CBS 100506 

Phlogicylindrium eucalypti DQ923534 

Cyphellophora laciniata CBS 190.61 

Cladophialophora proteae CBS 111667 

100 

84 

82 

67 

100 

Phaeococcomyces catenatus AF050277 

Exophiala sp. 3 CPC 12171 



Glyphium elatum AF346420 

67 100 

“Cladosporium” adianticola DQ008144 


Metulocladosporiella musae DQ008162 

Metulocladosporiella musicola DQ008159 

Thysanorea papuana EU041871 

Veronaea botryosa EU041874 

Ceramothyrium carniolicum AY004339 

Cyphellophora hylomeconis CBS 113311 

Exophiala eucalyptorum CPC 11261 

98 

100 

100 

100 

Cladophialophora humicola AF050263 

Cladophialophora sylvestris CBS 350.83 

Cladophialophora hostae CPC 10737 

Cladophialophora scillae CBS 116461 





Capronia munkii AF050250 

Ramichloridium mackenziei AF050288 

Capronia pilosella AF050254 

Exophiala sp. 1 CBS 115142 

Exophiala sp. 2 CBS 115143 

53 

68 

90 

Exophiala pisciphila DQ823101 


Ramichloridium anceps AF050285 

Capronia acutiseta AF050241 


Cladophialophora australiensis CBS 112793 

Cladophialophora potulentorum CBS 115144 


85 89 

98 

100 


Cladophialophora carrionii AF050262 

Phialophora americana AF050283 


Cladophialophora chaetospira CBS 114747 

Cladophialophora chaetospira CBS 514.63 



Sordariomycetes, 

Xylariomycetidae, 

Xylariales 

Chaetothyriomycetes, Chaetothyriales, Herpotrichiellaceae 

Fig. 1. (Page 188–189). One of 5 000 equally most parsimonious trees obtained from a heuristic search with 100 random taxon additions of the LSU sequence alignment using 

PAUP v. 4.0b10. The scale bar shows 10 changes, and bootstrap support values from 1 000 replicates are shown at the nodes. Thickened lines indicate the strict consensus 

branches and ex-type sequences are printed in bold face. The tree was rooted to two sequences obtained from GenBank (Athelia epiphylla AY586633 and Paullicorticium 

ansatum AY586693). 

apex via 1–2(–3) flat-tipped, minute, denticle-like loci, 1–1.5 µm 

wide, 10–15 × 1.5–2 µm; scars minutely darkened and thickened, 

but not refractive. Conidia in extremely long chains (–60), simple 

or branched, subcylindrical, or narrowly ellipsoid, smooth, pale 

olivaceous, 0–1-septate, (7–)10–15(–20) × (1.5–)2(–2.5) µm, 

hila truncate, 1–1.5 µm wide, minutely thickened and darkenedrefractive. 


with smooth, undulate margins and dense aerial mycelium; surface 

hazel (middle), outer zone isabelline; reverse fuscous-black in 

middle, isabelline in outer zone. Colonies reaching 25 mm diam 

on SNA, and 40 mm diam on PDA after 1 mo at 25 °C in the dark; 


Specimen examined: Korea, Pyongchang, Hosta plantaginea (Hostaceae), 20 Sep. 

2003, H.D. Shin, HAL 2030 F, holotype, culture ex-type SMK 19664, CPC 10737 = 

CBS 121637, CPC 10738–10739. 

Notes: Although this species is morphologically similar to 

Cladophialophora scillae (Deighton) Crous, U. Braun & K. 

Schub. described below in this paper, C. hostae is treated as a 

separate taxon due to the differences in the length and width of its 

conidiophores and conidia in vitro, as well as 17 bp differences in 

the ITS DNA sequence data and a distinct ecology causing leafspots 

on a different, unrelated host. Based on disease symptoms 

caused on the living host leaves, C. hostae is a very unusual, 

unexpected member of the genus Cladophialophora. In vivo, the 

mycelium forms obvious hyphal strands and plates which are 

characteristic for Fusicladium species. The conidiophores and 

conidia are also fusicladium-like. Nevertheless, this species clusters 

within the Herpotrichiellaceae, i.e., it has to be placed in the genus 

Cladophialophora. Biotrophic species like C. hostae and C. scillae 

without phialidic synanamorphs render the differentiation between 

Cladophialophora and Fusicladium (incl. Pseudocladosporium) 

almost impossible without sequence data. Furthermore, the 

188


10 changes 

Satchmopsis brasiliensis DQ195797 

Neofabraea malicorticis AY544662 

Protoventuria alpina CBS 140.83 

Clathrosporium intricatum AY616235 

Zeloasperisporium hyphopodioides CBS 218.95 

96 72 

94 

99 

86 

71 

65 

88 

59 

71 

73 

91 

66 

100 

99 

81 

95 

100 

100 

80 

Veronaeopsis simplex EU041877 

Fusicladium africanum CPC 12828 


Sympoventuria capensis DQ885906 



100 

Fusicladium amoenum CBS 254.95 

Fusicladium intermedium CBS 110746 

Fusicladium rhodense CPC 13156 

Fusicladium pini CBS 463.82 

Fusicladium ramoconidii CBS 462.82 

90 

100 

Cylindrosympodium lauri CBS 240.95 

Venturia fraxini CBS 374.55 

Venturia macularis CBS 477.61 

Venturia maculiformis CBS 377.53 


Venturia alpina CBS 373.53 

Fusicladium fagi CBS 621.84 

Venturia sp. CBS 681.74 

Caproventuria hanliniana AF050290 

Venturia hystrioides CBS 117727 

Venturia lonicerae CBS 445.54 

Venturia chlorospora CBS 466.61 

Fusicladium catenosporum CBS 447.91 

Venturia helvetica CBS 474.61 

Venturia minuta CBS 478.61 

Venturia polygoni-vivipari CBS 114207 

Venturia atriseda CBS 378.49 

Venturia viennotii CBS 690.85 

Venturia anemones CBS 370.55 

Venturia aceris CBS 372.53 

Venturia cephalariae CBS 372.55 

Venturia tremulae var. tremulae CBS 257.38 

Venturia ditricha CBS 118894 

Venturia populina CBS 256.38 


Venturia inaequalis CBS 535.76 

Fusicladium pomi CBS 309.31 


Fusicladium mandshuricum CBS 112235 

Venturia tremulae var. grandidentatae CBS 695.85 

Venturia tremulae var. populi-albae CBS 694.85 

Fusicladium radiosum CBS 112625 

Venturia saliciperda CBS 214.27 


Fusicladium convolvularum CBS 112706 

Fusicladium oleagineum CBS 113427 

Fusicladium phillyreae CBS 113539 

Venturia nashicola CBS 793.84 

Venturia pyrina CBS 120825 

Venturia pyrina CBS 331.65 

Venturia aucupariae CBS 365.35 

Venturia crataegi CBS 368.35 

Apiosporina collinsii CPC 12229 

Fusicladium effusum CPC 4524 


60 

91 


Fusicladium carpophilum CBS 497.62 

Venturia cerasi CBS 444.54 

Leotiomycetes, Helotiales 

Dothideomycetes, Pleosporales, Venturiaceae 

Fig 1. (Continued). 

morphology of C. hostae in vivo and in vitro shows remarkable 

differences in conidiophore morphology, i.e., the growth in vivo is 

characteristically fusicladium-like (conidiophores macronematous, 

long, septate), whereas habit in vitro is rather pseudocladosporiumlike 

(conidiophores less developed, usually reduced to conidiogenous 

cells, short). However, several Fusicladium species have also been 

observed to exibit a Pseudocladosporium growth habit in culture, 

suggesting this growth plasticity to be rather common, and strongly 

influenced by growth conditions. 

Cladophialophora humicola Crous & U. Braun, sp. nov. 



Etymology: Named after its ecology, namely occurring in soil. 

Cladophialophorae bantianae similis, sed conidiis majoribus, (8–)11–14(–17) × 

(1.5–)2(–2.5) µm, locis conidiogenis et hilis angustioribus, 1–1.5 µm latis. 

In vitro: Mycelium composed of branched, smooth, pale 

olivaceous to pale brown hyphae, frequently forming hyphal coils, 

prominently guttulate, not to slightly constricted at the septa, 1–2 

µm wide, cells somewhat uneven in width. Conidiophores solitary, 

mostly inconspicuous and integrated in hyphae, varying from 

inconspicuously truncate lateral loci on hyphal cells, 1–1.5 µm wide, 

to occasionally terminal conidiophores, 0–3-septate, subcylindrical, 

proliferating sympodially, 10–30 × 1.5–3 µm, pale brown, smooth. 

189

Crous et al. 

Table 1. Isolates used for sequence analysis. 


Anungitopsis speciosa CBS 181.95*; INIFAT C94/135 

(ITS, LSU) 

Leaf litter of Buchenavia 

capitata Cuba R.F. Castañeda EU035401, EU035401 

Cladophialophora australiensis CBS 112793*; CPC 1377 Sports drink Australia – EU035402, EU035402 

Cladophialophora chaetospira CBS 114747 Phyllostachys bambusoides China L. Cai EU035403, EU035403 

CBS 115468; HKUCC 10147 Bamboo China – EU035404, EU035404 

CBS 491.70 Roots of Picea abies Denmark – EU035405, EU035405 

CBS 514.63; ATCC 16274; MUCL 8310 Soil, wheat field Germany – EU035406, EU035406 

Cladophialophora hostae CPC 10737* Hosta plantaginea Korea H.D. Shin EU035407, EU035407 

Cladophialophora humicola CBS 117536*; BBA 65570 Soil, arable Germany Z. Zaspel & H. Nirenberg EU035408, AF050263 

Cladophialophora potulentorum CBS 112222; CPC 1376; FRR 4946 Sports drink Australia N.J. Charley EU035409, EU035409 

CBS 114772; CPC 1375; FRR 4947 Sports drink Australia N.J. Charley EU035410, EU035410 

CBS 115144*; CPC 11048; FRR 3318 Apple juice – – DQ008141, DQ008141 

Cladophialophora proteae CBS 111667*; CPC 1514 Protea cynaroides South Africa L. Viljoen EU035411, EU035411 

Cladophialophora scillae CBS 116461* Scilla peruviana New Zealand C.F. Hill EU035412, EU035412 

Cladophialophora sylvestris CBS 350.83 Pinus sylvestris Netherlands – EU035413, EU035413 

“Cladosporium” adianticola CBS 735.87*; ATCC 200931; INIFAT C87/40 Adiantum sp. Cuba R.F. Castañeda & G. Arnold DQ008125, DQ008144 

Cylindrosympodium lauri CBS 240.95*; INIFAT C95/3-2 Laurus sp. 

Spain, Canary 

Islands R.F. Castañeda EU035414, EU035414 

Cyphellophora hylomeconis CBS 113311* Helomeco velane Korea H.D. Shin EU035415, EU035415 

Cyphellophora laciniata CBS 190.61*; ATCC 14166; MUCL 9569 Man, skin Switzerland K.M. Wissel EU035416, EU035416 

Exophiala eucalyptorum CPC 11261* Eucalyptus sp. New Zealand J. Stalpers EU035417, EU035417 

Exophiala sp. 1 CBS 115142; CPC 11044; FRR 5582 Fruit-based drink – – DQ008139, EU035418 

Exophiala sp. 2 CBS 115143*; CPC 11047; FRR 5599 Bottled spring water – – DQ008140, EU035419 

Exophiala sp. 3 CPC 12171 Prunus sp. Canada K.A. Seifert EU035420, EU035420 

CPC 12172 Prunus sp. Canada K.A. Seifert EU035421, EU035421 

CPC 12173 Prunus sp. Canada K.A. Seifert EU035422, EU035422 

Fusicladium africanum CPC 12828* Eucalyptus sp. South Africa P.W. Crous EU035423, EU035423 

CPC 12829 Eucalyptus sp. South Africa P.W. Crous EU035424, EU035424 

Fusicladium amoenum CBS 254.95*; ATCC 200947; CPC 3681; IMI 367525; Leaf litter of Eucalyptus Cuba R.F. Castañeda 

INIFAT C94/155; MUCL 39143 

grandis 

EU035425, EU035425 

Fusicladium carpophilum Venturia carpophila CBS 497.62; ETH 4568 Prunus sp. Switzerland – EU035426, EU035426 

190


Fusicladium catenosporum CBS 447.91* Salix triandra Germany H. Butin EU035427, EU035427 

Fusicladium convolvularum CBS 112706*; CPC 3884; IMI 383037 Convolvulus arvensis New Zealand C.F. Hill AY251082, EU035428 

Fusicladium effusum CPC 4524 Carya illinoinensis U.S.A. K. Stevenson AY251084, EU035429 

CPC 4525 Carya illinoinensis U.S.A. K. Stevenson AY251085, EU035430 

Fusicladium fagi CBS 621.84*; ATCC 200937 Fagus sylvatica Netherlands G.S. de Hoog EU035431, EU035431 

Fusicladium intermedium CBS 110746*; CPC 778; IMI 362702 Eucalyptus sp. Madagascar P.W. Crous EU035432, EU035432 

Fusicladium mandshuricum Venturia mandshurica CBS 112235*; CPC 3639 Populus simonii China – EU035433, EU035433 

Fusicladium oleagineum CBS 113427 Olea europaea New Zealand C.F. Hill EU035434, EU035434 

Fusicladium phillyreae CBS 113539; UPSC 1329 – Portugal B. d’Oliveira EU035435, EU035435 

Fusicladium pini CBS 463.82* Pinus sylvestris Netherlands G.S. de Hoog EU035436, EU035436 

Fusicladium pomi Venturia inaequalis CBS 309.31 – – – EU035437, EU035437 

CBS 535.76 Sorbus aria Switzerland – EU035460, EU035460 

Fusicladium radiosum Venturia tremulae CBS 112625; CPC 3638 Populus tremula France – EU035438, EU035438 

Fusicladium ramoconidii CBS 462.82*; CPC 3679 Pinus sp. Netherlands G.S. de Hoog AY251086, EU035439 

Fusicladium rhodense CPC 13156* Ceratonia siliqua Greece P.W. Crous EU035440, EU035440 

Polyscytalum fecundissimum CBS 100506 Fagus sylvatica Netherlands W. Gams EU035441, EU035441 

Zeloasperisporium hyphopodioides CBS 218.95*; IMI 367520; INIFAT C94/114; MUCL 39155 Air Cuba R. F. Castañeda EU035442, EU035442 

Apiosporina collinsii CPC 12229 Amelanchier alnifolia Canada L.J. Hutchinson EU035443, EU035443 

Protoventuria alpina CBS 140.83 Arctostaphylos uva-ursi Switzerland – EU035444, EU035444 

Sympoventuria capensis CBS 120136; CPC 12838 Eucalyptus sp. South Africa P.W. Crous DQ885906, DQ885906 

CPC 12839 Eucalyptus sp. South Africa P.W. Crous DQ885905, DQ885905 

CPC 12840 Eucalyptus sp. South Africa P.W. Crous DQ885904, DQ885904 

Venturia aceris CBS 372.53 Acer pseudoplatanus Switzerland – EU035445, EU035445 

Venturia alpina CBS 373.53 Arctostaphylos alpina Switzerland – EU035446, EU035446 

Venturia anemones CBS 370.55; IMI 163998 Anemone alpina France – EU035447, EU035447 

Venturia atriseda CBS 371.55 Gentiana punctata Switzerland – EU035448, EU035448 

CBS 378.49 Gentiana lutea Switzerland J.A. von Arx EU035449, EU035449 

Venturia aucupariae CBS 365.35; IMI 163987 Sorbus aucuparia Germany – EU035450, EU035450 

Venturia cephalariae CBS 372.55 Cephalaria alpina Switzerland – EU035451, EU035451 

Venturia cerasi CBS 444.54; ATCC 12119; IMI 163988 Prunus cerasus Germany – EU035452, EU035452 

Venturia chlorospora CBS 466.61; ETH 543 Salix caesia Switzerland J. Nüesch EU035453, EU035453 

CBS 470.61 Salix daphnoides France J. Nüesch EU035454, EU035454 

Venturia crataegi CBS 368.35 Crataegus sp. Germany – EU035455, EU035455 


191

Crous et al. 



Venturia ditricha CBS 118894 Betula pubescens var. 

tortuosa 

(ITS, LSU) 

Finland – EU035456, EU035456 

Venturia fraxini CBS 374.55 Fraxinus excelsior Switzerland – EU035457, EU035457 

Venturia helvetica CBS 474.61; ETH 2571; IMI 163990 Salix helvetica Switzerland J. Nüesch EU035458, EU035458 

Venturia hystrioides CBS 117727*; ATCC 96019; CPC 5391 Prunus avium cv. Bing U.S.A. R.G. Roberts EU035459, EU035459 

Venturia lonicerae CBS 445.54; IMI 163997 Lonicera coerulea Switzerland – EU035461, EU035461 

Venturia macularis CBS 477.61; ETH 2831 Populus tremula France – EU035462, EU035462 

Venturia maculiformis CBS 377.53 Epilobium montanum France – EU035463, EU035463 

Venturia minuta CBS 478.61; ETH 523; IMI 163991 Salix nigricans Switzerland J. Nüesch EU035464, EU035464 

Venturia nashicola CBS 793.84 Pyrus serotina Japan – EU035465, EU035465 

Venturia polygoni-vivipari CBS 114207; UPSC 2754 Polygonum viviparum Norway K. & L. Holm EU035466, EU035466 

Venturia populina CBS 256.38; IMI 163996 Populus canadensis Italy – EU035467, EU035467 

Venturia pyrina CBS 120825 Pyrus communis Brazil – EU035468, EU035468 

CBS 331.65 Pyrus sp. – – EU035469, EU035469 

Venturia saliciperda CBS 214.27; IMI 163993 – – – EU035470, EU035470 

CBS 480.61; ETH 2836 Salix cordata Switzerland – EU035471, EU035471 

Venturia sp. CBS 681.74 Cedrus atlantica France W. Gams EU035472, EU035472 

Venturia tremulae var. grandidentatae CBS 695.85 Populus tremuloides Canada – EU035473, EU035473 

Venturia tremulae var. populi-albae CBS 694.85 Populus alba France – EU035474, EU035474 

Venturia tremulae var. tremulae CBS 257.38 Populus tremula Italy – EU035475, EU035475 

Venturia viennotii CBS 690.85 Populus tremula France – EU035476, EU035476 

1 ATCC: American Type Culture Collection, Virginia, U.S.A.; BBA: Biologische Bundesanstalt für Land- und Forstwirtschaft, Berlin-Dahlem, Germany; CBS: Centraalbureau voor Schimmelcultures, Utrecht, The Netherlands; CPC: Culture collection of Pedro 

Crous, housed at CBS; ETH: Eidgenössische Technische Hochschule, Institute for Special Botany, Zürich, Switzerland; FRR: Division of Food Research, CSIRO, North Ryde, N.S.W., Australia; HKUCC: The University of Hong Kong Culture Collection, Dept. 

of Ecology and Biodiversity, University of Hong Kong, Pokfulam Road, China; IMI: International Mycological Institute, CABI-Bioscience, Egham, Bakeham Lane, U.K.; INIFAT: Alexander Humboldt Institute for Basic Research in Tropical Agriculture, Ciudad 

de La Habana, Cuba; MUCL: Mycotheque de l’ Université Catholique de Louvain, Louvain-la-Neuve, Belgium; UPSC: Uppsala University Culture Collection of Fungi, Museum of Evolution, Botany Section, Evolutionary Biology Centre, Uppsala, Sweden. 

2 ITS: internal transcribed spacer regions, LSU: partial 28S rDNA sequence. 


192





0.92 0.98 Neofabraea malicorticis AY544662 

Protoventuria alpina CBS 140.83 

0.94 Clathrosporium intricatum AY616235 

Leotiomycetes, Helotiales 

Satchmopsis brasiliensis DQ195797 

1.00 

Anungitopsis speciosa CBS 181.95 Incertae sedis 

0.98 1.00 Fasciatispora petrakii AY083828 

Sordariomycetes, 

Pidoplitchkoviella terricola AF096197 

0.98 Polyscytalum fecundissimum CBS 100506 Xylariomycetidae, 

1.00 Phlogicylindrium eucalypti DQ923534 Xylariales 

1.00 

0.94 

Cladophialophora humicola AF050263 

0.84 Cladophialophora sylvestris CBS 350.83 

1.00 Cladophialophora hostae CPC 10737 

Cladophialophora scillae CBS 116461 

0.50 Cladophialophora proteae CBS 111667 

Phaeococcomyces catenatus AF050277 

1.00 Glyphium elatum AF346420 

1.00 Exophiala sp. 3 CPC 12171 




1.00 

1.00 

1.00 

0.85 

0.56 

1.00 

1.00 

1.00 




Thysanorea papuana EU041871 

Veronaea botryosa EU041874 



Ramichloridium mackenziei AF050288 

Ramichloridium anceps AF050285 






0.71 

1.00 

0.81 Exophiala pisciphila DQ823101 

0.56 Fonsecaea pedrosoi AF050276 

0.98 Exophiala sp. 1 CBS 115142 

0.60 Exophiala sp. 2 CBS 115143 

0.80 Cyphellophora laciniata CBS 190.61 


1.00 Cyphellophora hylomeconis CBS 113311 

1.00 Exophiala eucalyptorum CPC 11261 


0.77 0.68 

Cladophialophora australiensis CBS 112793 


1.00 Cladophialophora potulentorum CBS 114772 

1.00 Cladophialophora potulentorum CBS 112222 

1.00 

0.95 

1.00 

1.00 

1.00 

Cladophialophora carrionii AF050262 







Fig. 2. (Page 193–194). Consensus phylogram (50 % majority rule) of 1 500 trees resulting from a Bayesian analysis of the LSU sequence alignment using MrBayes v. 3.1.2. 

Bayesian posterior probabilities are indicated at the nodes. Ex-type sequences are printed in bold face. The tree was rooted to two sequences obtained from GenBank (Athelia 

epiphylla AY586633 and Paullicorticium ansatum AY586693). 

Chaetothyriomycetes, Chaetothyriales, Herpotrichiellaceae 

Conidiogenous cells integrated, inconspicuous, truncate, lateral 

loci 1–1.5 µm wide, or conidiogenous cells subcylindrical with 1–3 

sympodial loci (which appear as minute lateral denticles), 7–17 × 

1.5–2 µm; scars inconspicuous, neither darkened, refractive nor 

thickened. Conidia in short chains of up to 10, simple or branched, 

subcylindrical to narrowly ellipsoid, 0–1-septate, (8–)11–14(–17) 

× (1.5–)2(–2.5) µm, pale olivaceous to olivaceous-brown or pale 

brown, smooth, hila truncate, 1–1.5 µm wide, unthickened, neither 

darkened, nor refractive. 


uneven, feathery margins and dense aerial mycelium on PDA; 

pale olivaceous-grey in the middle, becoming olivaceous-grey 

in the outer zone (surface); reverse olivaceous-black, with greyolivaceous 

margins. Colonies reaching 7 mm diam after 2 wk at 25 

°C in the dark; colonies fertile. 

Specimen examined: Germany, Brandenburg, Müncheberg, from soil, Zaspel, 

Zalf & H. Nirenberg, holotype CBS H-19902, culture ex-type BBA 65570 = CBS 

117536. 

Notes: Phylogenetically Cladophialophora humicola is closely 

related to C. sylvestris Crous & de Hoog (see below). Morphologically 

the two species can be distinguished in that C. humicola lacks 

ramoconidia, and has 1-septate conidia, while those of C. sylvestris 

are 0–3-septate. 

Cladophialophora potulentorum Crous & A.D. Hocking, sp. nov. 


Etymology: Refers to its presence in fruit juices and sports drinks. 

Cladophialophorae carrionii similis, sed conidiis secundis majoribus, (6–)8–10(–13) 

× 2–3 µm. 

In vitro: Mycelium consisting of branched, septate, smooth, 

pale brown, guttulate, 1.5–2.5 µm wide hyphae. Conidiophores 

solitary, macronematous, well distinguishable under the dissecting 

microscope from aerial mycelium, pale to medium brown, 

subcylindrical, straight to somewhat curved, erect, with apical 

apparatus appearing as a tuft due to extremely long conidial 


193

Crous et al. 


Cylindrosympodium lauri CBS 240.95 

1.00 Fusicladium amoenum CBS 254.95 

1.00 Fusicladium intermedium CBS 110746 

1.00 

Fusicladium rhodense CPC 13156 

Fusicladium pini CBS 463.82 

1.00 1.00 1.00 Fusicladium ramoconidii CBS 462.82 

1.00 

Veronaeopsis simplex EU041877 



0.85 

0.95 

0.78 

0.56 

0.71 

1.00 

0.69 

0.79 

0.94 

0.56 

0.97 

0.91 

1.00 

0.97 

1.00 

1.00 

0.68 

0.55 

0.65 

0.61 

0.92 

0.94 

0.72 

0.95 




Zeloasperisporium hyphopodioides CBS 218.95 


Venturia alpina CBS 373.53 

Venturia fraxini CBS 374.55 

Venturia macularis CBS 477.61 

Venturia maculiformis CBS 377.53 

Apiosporina collinsii CPC 12229 

Fusicladium fagi CBS 621.84 

Venturia sp. CBS 681.74 

Caproventuria hanliniana AF050290 

Venturia hystrioides CBS 117727 

Venturia viennotii CBS 690.85 

Venturia inaequalis CBS 535.76 


Fusicladium radiosum CBS 112625 


Venturia tremulae var. populi-albae CBS 694.85 

Venturiatremulae var. grandidentatae CBS 695.85 

Fusicladium mandshuricum CBS 112235 

Fusicladium pomi CBS 309.31 


Venturia populina CBS 256.38 


Venturia ditricha CBS 118894 

Venturia tremulae var. tremulae CBS 257.38 

Fusicladium catenosporum CBS 447.91 

Venturia helvetica CBS 474.61 

Venturia minuta CBS 478.61 


Venturia polygoni-vivipari CBS 114207 

Venturia lonicerae CBS 445.54 


Fusicladium carpophilum CBS 497.62 

Venturia cerasi CBS 444.54 


Venturia anemones CBS 370.55 

Venturia aceris CBS 372.53 

Venturia cephalariae CBS 372.55 

Fusicladium convolvularum CBS 112706 

Venturia aucupariae CBS 365.35 

Venturia crataegi CBS 368.35 

Venturia nashicola CBS 793.84 

Venturia pyrina CBS 120825 

Venturia pyrina CBS 331.65 



Fusicladium oleagineum CBS113427 

Fusicladium phillyreae CBS 113539 

Dothideomycetes, Pleosporales, Venturiaceae 


Fig. 3. Cladophialophora australiensis (CBS 112793). A. Conidiophore. B–C. Subcylindrical ramoconidia, and ellipsoid conidia. Scale bar = 10 µm. 

194


Fig. 4. Cladophialophora chaetospira (CBS 114747). A–C. Hyphae giving rise to conidiophores with catenulate conidia. D–F. Conidia become up to 3-septate, frequently 

remaining attached in chains. Scale bars = 10 µm. 

Fig. 5. Cladophialophora hostae (CPC 10737). A–B. Conidiogenous loci (arrows). C. Hyphal coil. D–F. Branched conidial chains. G–H. Conidia. Scale bar = 10 µm. 


195

Crous et al. 

Fig. 6. Cladophialophora hostae (CPC 10737). Branched conidial chains with 

ramoconidia and conidia. Scale bar = 10 µm. 

Fig. 7. Cladophialophora humicola (CBS 117536). Conidiophore with branched 

conidial chains. Scale bar = 10 µm. 

Fig. 8. Cladophialophora humicola (CBS 117536). A. Hyphal coil. B. Conidiophore. C–F. Conidial chains with ramoconidia and conidia. Scale bar = 10 µm. 

196


Fig. 9. Cladophialophora potulentorum (CBS 115144). A. Colony on PDA. B. Conidiophore. C–D. Conidial chains. E–F. Ramoconidia and conidia. Scale bar = 10 µm. 

chains; conidiophores up to 5-septate, and 100 µm tall (excluding 

conidiogenous cells). Conidiogenous cells pale brown, smooth, 

terminal and lateral, subcylindrical, tapering towards subtruncate 

to truncate loci, 1 µm wide, somewhat darkened, thickened, but not 

refractive, loci appearing subdenticulate on lateral conidiogenous 

cells, mono- to polyblastic, proliferating sympodially, 10–35 × 1.5–2 

µm. Conidia pale brown, smooth, guttulate, occurring in branched 

chains of up to 60; hila somewhat darkened and thickened, but not 

refractive, 0.5 µm wide; ramoconidia subcylindrical, 0–1-septate, 

15–17(–20) × 2.5–3 µm; conidia ellipsoid, (6–)8–10(–13) × 2–3 

µm. 

Cultural characteristics: Colonies erumpent, spreading, with smooth 

margins and dense aerial mycelium on PDA, olivaceous-grey 

(surface), with a thin, olivaceous-black margin; reverse olivaceousblack; 

on OA olivaceous-grey (surface) with a wide olivaceousblack 

margin. Colonies reaching 25–30 mm diam after 1 mo at 25 

°C in the dark; colonies fertile, also sporulating in the agar. Not able 

to grow at 37 °C. 

Specimens examined: Australia, isolated from apple juice drink, Dec. 1986, A.D. 

Hocking, holotype CBS H-19901, culture ex-type CBS 115144 = CPC 11048; 

Australia, isolated from sports drink, Feb. 1996, A.D. Hocking, CBS 114772 = CPC 

1375 = FRR 4947; Australia, isolated from sports drink, Feb. 1996, A.D. Hocking, 

CBS 112222 = FRR 4946. 

Fig. 10. Cladophialophora potulentorum (CBS 115144). Conidiophores with chains 

of ramoconidia and conidia. Scale bar = 10 µm. 


Notes: Originally this taxon, isolated from fruit and sports drinks, 

was thought to be an undescribed species of Pseudocladosporium 

(= Fusicladium, see below). However, upon closer examination, this 

197

Crous et al. 

Fig. 11. Cladophialophora proteae (CBS 111667). A. Colony on OA. B–C. Conidiophores. D–H. Catenulate conidia. Scale bars = 10 µm. 

proved not to be the case. Conidiophores appear as distinct tufts 

under the dissecting microscope, and are readily distinguishable 

from the superficial mycelium, as is normally observed in species 

of Fusicladium, but the conidial chains are extremely long, and the 

conidia tend to be more ellipsoid than the predominantly fusiform or 

subcylindrical conidia observed in species of Fusicladium. Hyphal 

coils were also not observed in cultures of C. potulentorum, but 

are rather common in species of Fusicladium. The phylogenetic 

position of this taxon within the Herpotrichiellaceae clade also 

supports inclusion in the genus Cladophialophora. 

Cladophialophora proteae Viljoen & Crous, S. African J. Bot. 64: 

137. 1998. Fig. 11. 

≡ Pseudocladosporium proteae (Viljoen & Crous) Crous, in Crous et al., 

Cultivation and Diseases of Proteaceae: Leucadendron, Leucospermum 

and Protea: 101. 2004. 

In vitro: Mycelium consisting of branched, septate hyphae, often 

forming strands, anastomosing, smooth to finely verruculose, 

frequently constricted at septa, olivaceous, 3–4 µm wide; hyphal 

cells in older cultures becoming swollen, up to 6 µm wide. 

Conidiophores reduced to conidiogenous cells. Conidiogenous 

cells holoblastic, integrated, forming short, truncate protuberances, 

2–3 × 1.5–2 µm, concolorous with mycelium, subcylindrical. 

Conidia in vitro arranged in long acropetal chains (up to 20), simple 

or branched, subcylindrical to oblong-doliiform, (9–)13–17(–22) × 

2.5–3(–4) µm in vitro on MEA, (9–)16–22(–25) × (2.5–)3–4(–6) 

µm on SNA; 0–1(–4)-septate, pale brown to pale olivaceous, 

smooth, hila subtruncate to truncate, not thickened, but somewhat 

refractive. 


mycelium on PDA; margins irregular, feathery; greyish rose, with 

patches of pale olivaceous-grey (surface); reverse olivaceous-grey. 

Colonies reaching 10 mm diam after 2 wk at 25 °C in the dark; 


Specimen examined: South Africa, Western Cape Province, Stellenbosch, J.S. 

Marais Nature Reserve, leaves of Protea cynaroides (Proteaceae), 26 Aug. 1996, L. 

Viljoen, holotype PREM 55345, culture ex-type CBS 111667. 

Notes: Cladophialophora proteae differs from species of Fusicladium 

(= Pseudocladosporium) based on its colony colour, the slimy 

nature of colonies, as well as its conidia that have inconspicuous, 

unthickened hila (Fig. 11) (Crous et al. 2004), unlike those observed 

in species of Fusicladium. Sequence data show that this species is 

not allied to the Venturiaceae, but to the Herpotrichiellaceae. 

Cladophialophora scillae (Deighton) Crous, U. Braun & K. Schub., 


Basionym: Cladosporium scillae Deighton, N. Zealand J. Bot. 8: 

55. 1970. 

≡ Fusicladium scillae (Deighton) U. Braun & K. Schub., IMI Descriptions of 

Fungi and Bacteria 152: 1518. 2002. 

198


Fig. 12. Cladophialophora scillae (CBS 116461). A–C. Conidiophores. D–F. Catenulate conidia. Scale bar = 10 µm. 

In vivo: see Schubert & Braun (2002a) and Schubert et al. (2003). 

In vitro: Mycelium consisting of branched, septate, smooth, greenbrown 

to medium brown, guttulate hyphae, variable in width, 1.5–3 

µm diam. Conidiophores lateral or terminal on hyphae, erect, 

straight to slightly flexuous, solitary, in some cases aggregated, 

subcylindrical, curved to geniculate-sinuous, unbranched, up to 55 

µm long, 2–3 µm wide, 0–7-septate, septa in short succession, 

pale to medium brown, somewhat paler towards apices, smooth. 

Conidiogenous cells integrated, terminal or lateral as individual loci 

on hyphal cells, straight to curved, subcylindrical, up to 14(–18) µm 

long and 2 µm wide, pale to medium brown, smooth, with a single or 

few subdenticulate to denticulate loci at the apex due to sympodial 

proliferation, or reduced to individual loci, 0.8–1.5(–2) µm wide; 

scars minutely thickened and darkened, but not refractive. Conidia 

occurring in long, unbranched or loosely branched chains (–30), 

straight to slightly curved, ellipsoid to mostly narrowly subcylindrical, 

obclavate in some larger, septate conidia, (5–)10–20(–35) × 1.5–3 

µm, 0–1(–3)-septate, sometimes slightly constricted at the septa, 

subhyaline to pale brown, smooth, guttulate, tapering at ends 

to subtruncate hila, 0.8–1.5 µm wide, minutely thickened and 

darkened, but not refractive; microcyclic conidiogenesis occurring. 


smooth, even margins and dense, abundant aerial mycelium on 

PDA; grey-olivaceous (surface); reverse dark olivaceous. Colonies 

on OA olivaceous-grey, smoke-grey due to profuse sporulation, 


reverse olivaceous-grey to iron-grey, velvety, aerial mycelium 

sparse, diffuse. Colonies reaching 20 mm diam on SNA, and 40 

mm on PDA after 1 mo at 25 °C in the dark; colonies fertile. 

Specimens examined: New Zealand, Levin, on Scilla peruviana (Hyacinthaceae), 

21 Dec. 1965, G.F. Laudon, IMI 116997 holotype; Auckland, Manurewa, Auckland 

Botanic Gardens, on leaf spots of Scilla peruviana, 25 Apr. 2004, C.F. Hill, 1044, 

CBS H-19903, epitype designated here, culture ex-type CBS 116461. 

Notes: In culture Cladophialophora scillae forms a 

pseudocladosporium-like state, though the scars are somewhat 

darkened and thickened, but not refractive. Conidiophores are 

reduced to conidiogenous cells that are integrated in the mycelium, 

terminal or lateral, frequently also as an inconspicuous lateral 

denticle, with a flat-tipped scar. Conidia occur in long, branched 

chains, which are subcylindrical to narrowly ellipsoid, and are 

up to 35 µm long, 1.5–3 µm wide, thus longer and thinner than 

reported on the host, which were 0–3-septate, subcylindrical to 

ellipsoid-ovoid, 7–22 × 2.5–4 µm. Due to the fusicladioid habit of 

this species in vivo, Schubert & Braun (2002a) reallocated it to 

Fusicladium. Based on ITS sequence data, morphology and cultural 

characteristics, Cladophialophora scillae was almost identical to 

an isolate obtained from leaf spots of Hosta plantaginea in Korea. 

These isolates appeared to resemble species of Fusicladium, but 

phylogenetically they clustered in the Herpotrichiellaceae. Therefore, 

“Fusicladium” scillae was placed in the genus Cladophialophora. 

As far as we are aware, this species and C. hostae are first reports 

of phytopathogenic species within the genus Cladophialophora. 

199

Crous et al. 

Fig. 13. Cladophialophora sylvestris (CBS 350.83). A–B. Conidiophores. C. Catenulate conidia. D. Conidial mass. Scale bar = 10 µm. 

Fig. 14. Cyphellophora hylomeconis (CBS 113311). A. Colony on PDA. B–C. Hyphae with truncate conidiogenous loci. D–F. Conidia. Scale bars = 10 µm. 

Cladophialophora sylvestris Crous & de Hoog, sp. nov. 


Etymology: Refers to its host, Pinus sylvestris. 

Cladophialophorae humicolae similis, sed conidiis 0–3-septatis, (7–)10–16(–20) × 

1.5–2 µm. 

Mycelium composed of branched, smooth, pale olivaceous to 

pale brown hyphae, frequently forming hyphal coils, not to slightly 

constricted at the septa, 1–2 µm wide. Conidiophores medium 

brown, subcylindrical, flexuous, mononematous, multiseptate, up 

to 50 µm long, and 2–3 µm wide. Conidiogenous cells apical, 

sympodial, pale brown, 5–12 × 2–3 µm; scars somewhat darkened 

and thickened, not refractive. Conidia occurring in branched chains; 

ramoconidia up to 2 µm wide, giving rise apically to disarticulating 

chains of conidia; smooth, 0–3-septate, pale olivaceous, 

subcylindrical, (7–)10–16(–20) × 1.5–2 µm, with truncate ends; hila 

somewhat darkened and thickened, not refractive. 

Cultural characteristics: Colonies erumpent on PDA, with smooth, 

catenulate margins; iron-grey (surface); reverse greenish black. 



Specimen examined: Netherlands, Kootwijk, needle litter of Pinus sylvestris 

(Pinaceae), 8 Nov. 1982, G.S. de Hoog, holotype CBS H-19917, culture ex-type 


Notes: Morphologically CBS 350.83 was originally identified as 

Polyscytalum griseum Sacc., but the latter is reported to have 

conidia that are 5–5.5 × 1 µm (Saccardo 1877), which is much 

smaller than that observed for the present isolate. Furthermore, 

the type species of Polyscytalum, P. fecundissimum Riess (CBS 

100506), does not cluster within the Herpotrichiellaceae, thus 

suggesting that CBS 350.83 is best treated as a new species of 

Cladophialophora. 

Cyphellophora hylomeconis Crous, de Hoog & H.D. Shin, sp. 


Etymology: Named after its host genus, Hylomecon. 

Cyphellophorae lacinatae similis, sed conidiis longioribus et leniter angustioribus, 

(15–)25–35(–55) × (2.5–)3(–4) µm. 

Mycelium consisting of branched, greenish brown, septate, 

branched, smooth, 3–5 µm wide hyphae, constricted at septa. 

Conidiogenous cells phialidic, intercalary, appearing denticulate, 

1 µm tall, 1.5–2 µm wide, with minute collarettes (at times 

200


proliferating percurrently). Conidia sickle-shaped, smooth, medium 

brown, guttulate, (1–)3(–5)-septate, constricted at septa, widest in 

middle, or lower third of the conidium; apex subacutely rounded, 

base subtruncate, or having a slight constriction, giving rise to a 

foot cell, 1 µm long, 0.5–1 µm wide, subacutely rounded, (15–)25– 

35(–55) × (2.5–)3(–4) µm; a marginal frill is visible above the foot 

cell, suggesting this foot cell may be the onset of basal germination; 

conidia also anastomose and undergo microcyclic conidiation in 

culture. 

Cultural characteristics: Colonies slow-growing, slimy, aerial 

mycelium absent, margins smooth, catenate; surface crumpled, 

olivaceous-black to iron-grey. Colonies reaching 20 mm diam 

after 1 mo at 25 °C in the dark on PDA, 12 mm on SNA; colonies 

fertile. 

Specimen examined: Korea, Yangpyeong, on leaves of Hylomecon verlance 

(Papaveraceae), 4 Jun. 2003, H.D. Shin, holotype CBS H-19907, isotype SMK 

19550, culture ex-type CBS 113311. 

Notes: Cyphellophora hylomeconis is related to the type species 

of the genus, Cyphellophora laciniata G.A. de Vries, which also 

resides in the Herpotrichiellaceae. The genus Cyphellophora G.A. 

de Vries is phenetically distinguished from Pseudomicrodochium 

B.C. Sutton, typified by P. aciculare B.C. Sutton (1975) by melanized 

versus hyaline thalli. Phylogenetic confirmation is pending due to 

unavailability of sequence data. Decock et al. (2003) synonymised 

the hyaline genus Kumbhamaya M. Jacob & D.J. Bhat (Jacob & 

Bhat 2000) with Cyphellophora, but as no cultures of this fungus are 

available this decision seems premature. Nearly all Cyphellophora 

species accepted by Decock et al. (2003) have been found to be 

involved in cutaneous infections in humans. This also holds true 

for the species originally described as being environmental, C. 

vermispora Walz & de Hoog, which is closely related to C. suttonii 

(Ajello et al.) Decock and C. fusarioides (C.K. Campbell & B.C. 

Sutton) Decock known from proven human and animal infections. 

Decock et al. (2003) added the melanized species C. guyanensis 

Decock & Delgado, isolated as a saprobe from tropical leaf litter. 

Cyphellophora hylomeconis is the first species of the genus 

infecting a living plant host. ITS sequences are remote from those of 

the remaining Cyphellophora species, the nearest neighbour being 

C. pluriseptata G.A. de Vries, Elders & Luykx at 19.1 % distance 

(data not shown). Cyphellophora hylomeconis can be distinguished 

based on its conidial dimensions and septation. Conidia are larger 

than those of C. fusarioides (11–20 × 2–2.5 µm, 1–2-septate), and 

those of C. laciniata (11–25 × 2–5 µm, 1–3-septate) (for a key to 

the species see Decock et al. 2003). 

Exophiala sp. 1. Fig. 15. 

Mycelium consisting of smooth, branched, septate, medium brown, 

2–3 µm wide hyphae, regular in width, forming hyphal strands 

and hyphal coils, with free yeast-like cells present in culture; 

chlamydospores terminal on hyphae, frequently forming clusters 

or chains, medium brown, ellipsoid, 0–1-septate, up to 10 µm 

long and 5 µm wide. Conidiophores reduced to conidiogenous 

cells, or consisting of one supporting cell, giving rise to a single 

conidiogenous cell, subcylindrical to ellipsoid, medium brown, 

smooth, 5–12 × 3.5–4 µm, with 1(–3) phialidic loci, somewhat 

protruding, appearing subdenticulate at first glance under the 

light microscope. Conidiogenous cells integrated as lateral loci on 

hyphal cells, inconspicuous, 1–1.5 µm wide, with a slightly flaring 

collarette, (1–)1.5(–2) µm long. Conidia ellipsoid, smooth, guttulate, 

becoming brown, swollen and elongated, and at times 1-septate, 

4–5(–7) × (2.5–)3(–4) µm (description based on CBS 115142). 


Cultural characteristics: Colonies erumpent, spreading, with sparse 

to dense aerial mycelium on PDA, olivaceous-grey (surface), with a 

thin to wide, smooth, olivaceous-black margins; reverse olivaceousblack; 

on OA olivaceous-grey (surface) with wide, olivaceous-black 

margins. Colonies reaching 40–50 mm diam after 1 mo at 25 °C in 

the dark; colonies fertile, but sporulation sparse. Not able to grow 

at 37 °C. 

Specimen examined: Australia, from a fruit drink, May 2002, N.J. Charley, CBS 

115142 = CPC 11044 = FRR 5582. 

Notes: Species of Exophiala are frequently observed as agents 

of human mycoses in immunocompromised patients (de Hoog 

et al. 2000). They are found in the environment as slow-growing, 

oligotrophic colonisers of moist substrates. For example the 

thermotolerant species E. dermatitidis (Kano) de Hoog and E. 

phaeomuriformis (Matsumoto et al.) Matos et al. are common 

in public steam baths (Matos et al. 2003), while E. mesophila 

Listemann & Freiesleben can be found in showers and swimming 

pools (unpubl. data). Both species are able to cause infections 

in humans (Zeng et al. 2007). Several other species have been 

associated primarily with infections in fish and cold-blooded animals 

(Richards et al. 1978) and are occasionally found on humans 

(Madan et al. 2006). The occurrence of the present species in fruit 

drinks, therefore, is cause of concern, although it was unable to 

grow at 37 °C. This species forms part of a larger study, and will be 

treated elsewhere. 

Exophiala sp. 2. Fig. 16. 

Mycelium consisting of smooth, branched, septate, pale brown, 

1.5–3 µm wide hyphae, forming hyphal strands and hyphal coils; 

hyphae at times terminating in chains of ellipsoid chlamydospores 

that are medium brown, smooth, up to 10 µm long and 5 µm wide. 

Conidiophores subcylindrical, medium brown, smooth, consisting 

of a supporting cell and a single conidiogenous cell, or reduced 

to a conidiogenous cell, straight to curved, up to 30 µm long 

and 2–3 µm wide. Conidiogenous cells pale to medium brown, 

subcylindrical to narrowly ellipsoid or subclavate, with 1–3 apical, 

phialidic loci, 1 µm wide, 1–2 µm tall, collarette somewhat flaring, 

but mostly cylindrical, 7–20 × 2–2.5 µm; at times proliferating 

percurrently. Conidia ellipsoid, smooth, guttulate, hyaline, becoming 

pale olivaceous, apex obtuse, base subtruncate, (4–)5–7(–10) × 

2–2.5(–3) µm. 

Cultural characteristics: Colonies spreading with smooth, 

submerged margins, moderate aerial mycelium on PDA, sparse on 

OA, on PDA and OA olivaceous-grey (surface), with a wide, irongrey 

margin; reverse iron-grey. Colonies reaching 40–50 mm diam 

after 1 mo at 25 °C in the dark; colonies fertile. Not able to grow 

at 37 °C. 

Specimen examined: Australia, from bottled spring water, May 2003, N.J. Charley, 

CBS 115143 = CPC 11047 = FRR 5599. 

Notes: This strain represents another taxon occurring in bottled 

drinks destined for human consumption. As it is unable to grow at 

37 °C, it does not appear to pose any serious threat to human 

health. This species forms part of a larger study, and will be treated 

elsewhere. 

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Crous et al. 

Fig. 15. Exophiala sp. 1 (CBS 115142). A. Colony on PDA. B. Hyphal coil. C. Hyphal strand. D–H. Conidiogenous cells and loci. I–O. Conidiogenous cells and conidia. Scale 


Fig. 16. Exophiala sp. 2 (CBS 115143). A. Conidiogenous cells. B. Conidiophore with hyphal coil. C. Conidiogenous cell with hyphal strand. D. Conidia. Scale bar = 10 µm. 

202


Fig. 17. Exophiala eucalyptorum (CPC 11261). A. Colony on PDA. B–H. Hyphae, conidiogenous cells and conidia. Scale bars = 10 µm. 

Exophiala eucalyptorum Crous, sp. nov. MycoBank MB504533. 

Fig. 17. 

Etymology: Named after its occurrence on Eucalyptus leaves. 

Exophialae spiniferae similis, sed conidiis fusoidibus-ellipsoideis, (5–)6–8(–10) 

× (3–)4–5(–7) µm, et cellulis conidiogenis saepe catenatis, in catenis brevibus, 

dividentibus. 

Mycelium consisting of smooth to finely verruculose, branched, 

septate, 2–4 µm wide hyphae, at times giving rise to chains of 

dark brown, fusoid-ellipsoid chlamydospores, which can still have 

phialides, suggesting they were conidiogenous cells; hyphae 

becoming constricted at septa when fertile. Conidiophores reduced 

to conidiogenous cells. Conidiogenous cells numerous, terminal 

and lateral, mono- to polyphialidic, 5–15 × 3–5 µm; loci 1–1.5 µm 

wide and tall, with inconspicuous collarettes, at time proliferating 

percurrently; conidiogenous cells fusoid-ellipsoid, and frequently 

breaking off, appearing as short chains of conidia, but distinct in 

having conidiogenous loci. Conidia fusoid-ellipsoid, apex acutely 

rounded, base subtruncate, (5–)6–8(–10) × (3–)4–5(–7) µm; 

frequently becoming fertile, septate and brown with age. 

Cultural characteristics: Colonies erumpent, convex, smooth, slimy, 

margins feathery to crenate and smooth; aerial mycelium absent, 

growth yeast-like. Colonies on PDA, OA and SNA chestnut on 

surface and reverse. Colonies reaching 4 mm diam after 2 wk on 

PDA at 25 °C in the dark. 

Specimen examined: New Zealand, Wellington Botanical Garden, on leaf litter of 

Eucalyptus sp. (Myrtaceae), Mar. 2004, J.A. Stalpers, holotype CBS H-19905, 

culture ex-type CBS 121638 = CPC 11261. 


Notes: Exophiala eucalyptorum is rather characteristic in that, 

in culture, chains of conidiogenous cells frequently detach from 

hyphae, appearing as short, intact chains of fertile conidia. 

Its phylogenetic position is somewhat outside the core of the 

Herpotrichiellaceae containing most Capronia teleomorphs and 

the remaining opportunistic Exophiala species, but still within the 

Chaetothyriales (Figs 1–2). 

Members of Venturiaceae 

Anungitea B. Sutton and Anungitopsis R.F. Castañeda & 

W.B. Kendr. 

Sutton (1973) erected the genus Anungitea to accommodate 

species with brown, mononematous conidiophores bearing 

apically aggregated, flat-tipped, subdenticulate conidiogenous loci 

that give rise to chains of pale brown subcylindrical conidia with 

thickened, darkened hila. He compared the type species, A. fragilis 

B. Sutton with anamorph genera of the Mycosphaerellaceae, 

but did not compare it to Fusicladium, to which it is remarkably 

similar. Castañeda & Kendrick (1990b) introduced the genus 

Anungitopsis based on A. speciosa R.F. Castañeda & W.B. Kendr. 

This genus was distinguished from Anungitea by its formation of 

subdenticulate conidiogenous loci distributed along the apical 

region of the conidiophore, and by the relatively poorly defined 

appearance of these loci. No cultures are available of the extype 

species of Anungitea, but we studied strains of Anungitopsis 

203

Crous et al. 

Fig. 18. Cylindrosympodium lauri (CBS 240.95). A–C. Conidiophores with conidiogenous loci. D. Conidia. Scale bar = 10 µm. 

amoena R.F. Castañeda & Dugan (CBS 254.95, ex-type), and 

Anungitopsis intermedia Crous & W.B. Kendr. (CBS 110746, 

ex-epitype), and found them to cluster adjacent to Fusicladium 

(Venturiaceae). However, the ex-type strain of Anungitopsis 

speciosa (CBS 181.95), type species of Anungitopsis, clustered 

distantly from all other species, confirming that the genus name 

Anungitopsis is not available for any of the taxa treated here. In 

any case, A. speciosa has unusual subdenticulate conidiogenous 

loci with indistinct marginal frills, and these are obviously different 

from those of anungitea- and fusicladium-like anamorphs, including 

A. amoena and A. intermedia. The latter two species previously 

referred to as Anungitopsis belong to a sister clade of the Venturia 

(Fusicladium, incl. Pseudocladosporium) clade. Sympoventuria 

(Crous et al. 2007b), which produces a sympodiella-like anamorph 

in culture, is the only teleomorph of this clade hitherto known. The 

venturia-like habit of Sympoventuria, connected with fusicladium- 

/ pseudocladosporium-like anamorphs distributed in both clades, 

indicates a close relation between these clades, suggesting a 

placement in the Venturiaceae. Schubert et al. (2003) referred to 

the difficulty to distinguish between Anungitea and Fusicladium. 

Anungitea is undoubtedly heterogeneous. Anungitea rhabdospora 

P.M. Kirk (Kirk 1983) is, for instance, intermediate between 

Anungitea (conidiophores with a terminal denticulate conidiogenous 

cell, but conidia disarticulating in an arthroconidium-like manner) 

and Sympodiella B. Kendr. (conidiophores distinctly sympodial, 

forming arthroconidia). Other species assigned to Anungitea 

possess a distinctly swollen, lobed conidiophore base, e.g. A. 

heterospora P.M. Kirk (Kirk 1983), which is comparable with other 

morphologically similar genera, e.g., Parapleurotheciopsis P.M. 

Kirk (Kirk 1982), Rhizocladosporium Crous & U. Braun (see Crous 

et al. 2007a – this volume), and Subramaniomyces Varghese & 

V.G. Rao (Varghese & Rao 1979, Kirk 1982). The application of 

Anungitea depends, however, on the affinity of A. fragilis, the type 

species, of which sequence data are not yet available. The best 

solution for this problem is the widened application of Fusicladium 

(incl. Pseudocladosporium) to both sister clades, i.e., to the whole 

Venturiaceae. Morphologically a distinction between fusicladioid 

anamorphs of both clades is impossible. The more “fusicladium-like” 

growth is mainly characteristic for the fruiting in vivo, above all in 

biotrophic taxa, whereas the more “pseudocladosporium-like” habit 

is typical for the growth in vitro and in saprobic taxa, a phenomenon 

which is also evident in species of the morphologically similar 

genus Cladophialophora (see C. hostae and C. scillae). A potential 

placement of Anungitea fragilis within the Venturiaceae, which has 

still to be proven, would render the genus Anungitea a synonym of 

Fusicladium, but in the case of a quite distinct phylogenetic position 

a new circumscription of this genus, excluding the Venturiaceae 

anamorphs, would be necessary. Thus, a final conclusion about 

Anungitea has to be postponed, awaiting cultures and sequence 

analyses of its type species. 

The taxonomic placement of a fungus from the Canary 

Islands, isolated from leaf litter of Laurus sp. (CBS 240.95), is 

somewhat problematic. It clusters within the Venturiaceae, but 

not within Venturia s. str. itself, and it does not fit into the current 

morphological concept of Fusicladium (incl. Pseudocladosporium). 

Based on its solitary, cylindrical, hyaline conidia and pale brown 

conidiogenous structures, it resembles species accommodated in 

Cylindrosympodium W.B. Kendr. & R.F. Castañeda (Castañeda & 

Kendrick 1990a, Marvanová & Laichmanová 2007). 

Cylindrosympodium lauri Crous & R.F. Castañeda, sp. nov. 


Etymology: Named after the host genus it was collected from, 

Laurus. 

Cylindrosympodii variabilis similis, sed conidiophoris longioribus, ad 70 µm, conidiis 

subhyalinis vel dilute olivaceis. 

Mycelium consisting of brown, smooth, septate, branched hyphae, 

1.5–2.5 µm wide. Conidiophores macronematous, mononematous, 

solitary, erect, subcylindrical, straight to geniculate-sinuous, 

medium brown, smooth, 35–70 × 2.5–4 µm, 1–5-septate. 

Conidiogenous cells terminal, integrated, pale to medium brown, 

smooth, 10–35 × 2–3 µm, proliferating sympodially, with one to 

several flat-tipped loci, 1.5–2 µm wide; scars somewhat darkened, 

minutely thickened, but not refractive. Conidia solitary, subacicular 

to narrowly subcylindrical, apex subobtuse, base truncate, or 

somewhat swollen, straight or curved, smooth, subhyaline to very 

pale olivaceous, guttulate, (45–)60–70(–80) × 2.5–3(–3.5) µm, 

(4–)6–8-septate; scars are somewhat darkened, minutely thickened, 

but not refractive, 2.5–3 µm wide. 

Cultural characteristics: Colonies erumpent, convex, with smooth, 

lobed margins, and moderate, dense aerial mycelium on PDA; 

mouse-grey in the central part, and dark mouse-grey in the outer 

zone (surface); reverse dark mouse-grey. Colonies reaching 5 mm 

diam after 2 wk at 25 °C in the dark; colonies fertile. 

204


Specimen examined: Spain, Canary Islands, leaf litter of Laurus sp. (Lauraceae), 4 

Jan. 1995, R.F. Castañeda, holotype CBS H-19909, culture ex-type CBS 240.95. 

Note: The present fungus differs from Cylindrosympodium variabile 

(de Hoog) W.B. Kendr. & R.F. Castañeda (de Hoog 1985) in that the 

conidiophores are much longer, the conidia are subhyaline to very 

pale olivaceous, and the scars and hila are thin, slightly darkened, 

but not refractive. 

Venturia Sacc. and its anamorph Fusicladium 

Venturia Sacc., Syll. fung. (Abellini) 1: 586. 1882. 

= Apiosporina Höhn., Sitzungsber. Kaiserl. Akad. Wiss., Math.-Naturw. Cl., Abt. 

1, 119: 439. 1910, syn. nov. 

= Metacoleroa Petr., Ann. Mycol. 25: 332. 1927, syn. nov. 

= Caproventuria U. Braun, A Monograph of Cercosporella, Ramularia and Allied 

Genera (Phytopathogenic Hyphomycetes) 2: 396. 1998, syn. nov. 

For additional synonyms see Sivanesan, The bitunicate 

Ascomycetes and their anamorphs: 604. 1984. 

Anamorph: Fusicladium Bonord., Handb. Mykol.: 80. 1851. 

= Pseudocladosporium U. Braun, A Monograph of Cercosporella, Ramularia 

and Allied Genera (Phytopathogenic Hyphomycetes) 2: 392. 1998, syn. nov. 

For additional synonyms, see Schubert et al. (2003). 

Notes: The genus Caproventuria, based on C. hanliniana (U. Braun 

& Feiler) U. Braun, was erected to accommodate saprobic, soil-borne 

venturia-like ascomycetes with numerous ascomatal setae, and an 

anamorph quite distinct from Fusicladium (Braun 1998). The genus 

Metacoleroa is based on M. dickiei (Berk. & Broome) Petr., which 

clusters in the Venturiaceae, adjacent to Caproventuria, which has 

Pseudocladosporium anamorphs. Metacoleroa was retained by Barr 

(1987) as separate from Venturia based on its superficial ascomata 

with a thin, stromatic layer beneath the ascomata. Whether these 

criteria still justify the separation of Caproventuria and Metacoleroa 

from Venturia is debatable, and the names Venturia dickiei (Berk. & 

Broome) Ces. & de Not. and Venturia hanliniana (U. Braun & Feiler) 

Unter. are available for these organisms. The genus Apiosporina, 

which is based on Apiosporina collinsii (Schwein.) Höhn., clusters in 

the Venturiaceae, as was to be expected based on its Fusicladium 

anamorph (Schubert et al. 2003). It was distinguished from Venturia 

species by having ascospores strictly septate near the lower end 

(Sivanesan 1984). 

The anamorph genus Fusicladium has been monographed by 

Schubert et al. (2003). Morphological as well as molecular studies 

(Beck et al. 2005) demonstrated that the genus Venturia with its 

Fusicladium anamorphs is monophyletic. A separation of Venturia 

into various uniform subclades based on the previous anamorph 

genera Fusicladium, Pollaccia and Spilocaea was not evident and 

could be rejected. As in cercosporoid anamorphs of Mycosphaerella, 

features such as the arrangement of the conidiophores (solitary, 

fasciculate, sporodochial), the proliferation of conidiogenous cells 

(sympodial, percurrent) and shape, size as well as formation of 

conidia (solitary, catenate) proved to be of little taxonomic value at 

generic level. Hence, Schubert et al. (2003) proposed to maintain 

Fusicladium emend. as sole anamorph genus for Venturia. 

The genus Fusicladosporium Partridge & Morgan-Jones (type 

species: Cladosporium carpophilum Thüm.) (Partridge & Morgan- 

Jones 2003), recently erected to accommodate fusicladium-like 

species with catenate conidia, represents a further synonym of 

Fusicladium. 

Similar to their occurrence in vivo the conidiophores in 

vitro of species previously referred to the genera Spilocaea 

and Pollaccia are usually micronematous, conidia often appear 

to be directly formed on the mycelium, unilocal, determinate, 

mostly reduced to conidiogenous cells, sometimes forming a few 

percurrent proliferations, whereas the conidiophores of species 

of Fusicladium s. str. are mostly macronematous, but sometimes 

also micronematous. They are often initiated as short lateral, 

peg-like outgrowths of hyphae which proliferate sympodially, 

becoming slightly geniculate, forming a single, several or numerous 

subdenticulate to denticulate, truncate, unthickened or only slightly 

thickened, somewhat darkened-refractive conidiogenous loci. 

The genus Pseudocladosporium was described to be quite 

distinct from Fusicladium by being saprobic and connected with a 

different teleomorph, viz. Caproventuria (Braun 1998). However, 

since the type species of Caproventuria, C. hanliniana, with its 

anamorph Pseudocladosporium brevicatenatum (U. Braun & 

Feiler) U. Braun clusters together with numerous Venturia species, 

the genus Pseudocladosporium should be reduced to synonymy 

with Fusicladium. Morphologically there is no clear delimitation 

between Fusicladium and Pseudocladosporium. The typically 

pseudocladosporium-like habit, characterised by forming solitary 

conidiophores, often reduced to conidiogenous cells or even 

micronematous, and conidia formed in long chains, is mainly found 

in culture, above all in saprobic taxa. The fusicladium-like growth 

with well-developed macronematous conidiophores is usually more 

evident in vivo, above all in biotrophic taxa. There are, however, all 

kinds of transitions between these two genera. 

Fusicladium africanum Crous, sp. nov. MycoBank MB504535. 

Fig. 19. 

Etymology: Named after the continent from which it was collected, 

Africa. 

Fusicladio brevicatenato similis, sed conidiophoris brevioribus, 5–10 µm longis, 

conidiis minoribus, ad 20 × 3.5 µm, 0(–1)-septatis, locis conidiogenis et hilis 

angustioribus, 1–1.5 µm latis. 

Mycelium composed of smooth, medium brown, branched, 

septate, 1.5–2 µm wide hyphae, frequently forming hyphal coils. 

Conidiophores reduced to conidiogenous cells, solitary, pale to 

medium brown, smooth, inconspicuous, integrated in hyphae, 

varying from small, truncate lateral loci on hyphal cells, 1–1.5 µm 

wide, to micronematous conidiogenous cells, 5–10 × 2–3 µm; monoto 

polyblastic, sympodial, scars inconspicuous, 1 µm wide. Conidia 

in long, branched chains of up to 40, subcylindrical, 0(–1)-septate, 

pale brown, smooth; hila truncate, 1 µm wide, unthickened, neither 

darkened nor refractive; ramoconidia (11–)15–17(–20) × 2–3(–3.5) 

µm; conidia (8–)11–17 × 2–2.5 µm. 

Cultural characteristics: Colonies somewhat erumpent, with 

moderate aerial mycelium and smooth, lobate margins on PDA, 

ochreous to umber (surface); reverse dark umber; on OA umber; 

on SNA ochreous. Colonies reaching 9 mm diam on PDA after 2 wk 

at 25 °C in the dark; colonies fertile. 

Specimen examined: South Africa, Western Cape Province, Malmesbury, 

Eucalyptus leaf litter, Jan. 2006, P.W. Crous, holotype CBS H-19904, cultures extype 

CPC 12828 = CBS 121639, CPC 12829 = CBS 121640. 

Notes: Fusicladium africanum is a somewhat atypical member of the 

genus, as its conidial hila are quite unthickened and inconspicuous. 

Among biotrophic, leaf-spotting Fusicladium species a wider 

morphological variation was found pertaining to the structure of the 

conidiogenous loci and conidial hila, ranging from being indistinct, 

unthickened and not darkened-refractive to unthickened or almost 

so, but somewhat darkened-refractive (Schubert et al. 2003). 

Fusicladium africanum was found occurring with Sympoventuria 

capensis Crous & Seifert on Eucalyptus leaf litter in South Africa 

(Crous et al. 2007b). 


205

Crous et al. 

Fig. 19. Fusicladium africanum (CPC 12828). A. Colony on MEA. B. Hyphal coil. C. Branched conidial chain. D–F. Conidiophores with catenulate conidia. Scale bar = 10 µm. 

Fig. 20. Fusicladium amoenum (CBS 254.95). A–E. Conidiophores with conidiogenous loci. F. Conidia. Scale bar = 10 µm. 

206


Fig. 21. Fusicladium convolvularum (CBS 112706). A–B, D–I. Conidiophores with conidiogenous loci. C, J–K. Ramoconidia and conidia. Scale bars = 10 µm. 

Fusicladium amoenum (R.F. Castañeda & Dugan) Crous, K. 

Schub. & U. Braun, comb. nov. MycoBank MB504536. Fig. 20. 

Basionym: Anungitopsis amoena R.F. Castañeda & Dugan, 

Mycotaxon 72: 118. 1999. 

≡ Cladosporium amoenum R.F. Castañeda, in Untereiner et al., 1998, 

nom. nud. 

Specimen examined: Cuba, Santiago de Cuba, La Gran Piedra, fallen leaves of 

Eucalyptus sp. (Myrtaceae), 2 Nov. 1994, R.F. Castañeda, (Ho et al. 1999: 117, 

figs 2–3) iconotype, culture ex-type CBS 254.95 = ATCC 200947 = IMI 367525 = 

INIFAT C94/155 = MUCL 39143. 

Note: In culture F. amoenum has a typical pseudocladosporium-like 

morphology, though the scars are neither prominently thickened, 

nor refractive. 


207

Crous et al. 

Fig. 22. Fusicladium fagi (CBS 621.84). A. Conidiophore with truncate conidiogenous loci. B. Hypha with conidiogenous loci. C–G. Conidial chains. Scale bars = 10 µm. 

Fusicladium caruanianum Sacc., Ann. Mycol. 11: 20. 1913. 

≡ Pseudocladosporium caruanianum (Sacc.) U. Braun, Schlechtendalia 

9: 114. 2003. 

Fusicladium convolvularum Ondřej, Česká Mycol. 25: 171. 1971. 

Fig. 21. 

In vivo: Schubert et al. (2003: 37). 

In vitro on SNA: Mycelium unbranched or only sparingly branched, 

2–3 µm wide, septate, not constricted at septa, subhyaline to pale 

brown, smooth, walls unthickened or almost so. Conidiophores 

laterally arising from hyphae, erect, straight to somewhat flexuous, 

sometimes geniculate, unbranched, (6–)12–75 × (2.5–)3–4.5 

µm, aseptate or septate, pale brown or pale medium brown, 

smooth, walls somewhat thickened, sometimes only as short 

lateral conical prolongations of hyphae, occasionally irregular in 

shape. Conidiogenous cells integrated, terminal or conidiophores 

reduced to conidiogenous cells, sometimes geniculate, 6–29 µm 

long, proliferation sympodial, with several denticle-like loci, broadly 

truncate, 1.5–2(–2.5) µm wide, unthickened, somewhat refractive 

or darkened. Ramoconidia occurring, 20–28 × 5 µm, 0–1-septate, 

somewhat darker, pale medium brown, with a broadly truncate 

base, 3–4 µm wide, usually with several denticle-like apical loci. 

Conidia catenate, formed in unbranched or loosely branched 

chains, straight to sometimes curved, cells sometimes irregularly 

swollen, fusiform, subcylindrical, sometimes obpyriform, 13–35 × 

3.5–5.5(–6) µm, 0–3-septate, occasionally slightly constricted at 

the median septum, few very large conidia with up to five septa, 

up to 75 µm long, 4.5–6 µm wide, subhyaline to pale brown, 

smooth, walls slightly thickened, slightly attenuated towards apex 

and base, hila broadly truncate, 1–2 µm wide, unthickened or 

only slightly thickened, somewhat darkened-refractive; microcyclic 

conidiogenesis occurring, conidia often germinating. 

Cultural characteristics: Colonies on PDA spreading, somewhat 

erumpent, with moderate aerial mycelium and regular, but feathery 

margins; surface fuscous black, and reverse dark fuscous black. 

Colonies reaching 15 mm diam after 1 mo on PDA at 25 °C in the 

dark. 

Specimens examined: Czech Republic, Libina, okraj pole pod nadrazim (okr. 

Sumperk), on Convolvulus arvensis (Convolvulaceae), 7 Sep. 1970, Ondřej, 

holotype BRA. New Zealand, on leaves of Convolvulus arvensis, 7 Nov. 2000, 

C.F. Hill, epitype designated here CBS H-19911, culture ex-epitype CBS 112706 

= CPC 3884 = IMI 383037. 

Note: Conidiophores are somewhat longer and narrower in vitro 

than in vivo, and ramoconidia occur (Schubert & Braun 2002b, 

Schubert et al. 2003). 

208


Fig. 23. Fusicladium intermedium (CBS 110746). A–G. Conidiophores with sympodial conidiogenous loci. H. Conidia. Scale bar = 10 µm. 

Fusicladium fagi Crous & de Hoog, sp. nov. MycoBank MB504537. 

Fig. 22. 

Etymology: Named after its host, Fagus sylvatica. 

Fusicladio brevicatenato similis, sed conidiis secundis minoribus, (8–)11–17(–20) × 

3–3.5 µm, locis conidiogenis et hilis angustioribus, 1–1.5 µm latis. 

Mycelium consisting of pale to medium brown, smooth to finely 

verruculose, branched, 2–3 µm wide hyphae. Conidiophores 

integrated, terminal on hyphae, 0–1-septate, mostly reduced to 

conidiogenous cells, also lateral, visible as small, protruding, 

denticle-like loci, 10–15 × 2–3.5 µm. Conidiogenous cells 

subcylindrical, 5–15 × 2–3.5 µm, pale to medium brown, smooth 

to finely verruculose, tapering to 1–3 apical loci, 1–1.5 µm wide; 

scars inconspicuous. Conidia pale brown, smooth, guttulate, 

subcylindrical to narrowly ellipsoid, occurring in simple or branched 

chains, 0–1(–2)-septate, tapering towards subtruncate ends, 1.5– 

2.5 µm wide, aseptate conidia (8–)11–17(–20) × 3–3.5 µm, septate 

conidia up to 40 µm long and 4 µm wide; hila inconspicuous, i.e. 

neither thickened nor darkened-refractive; microcyclic conidiation 

common in older cultures. 


abundant aerial mycelium on PDA, and feathery to smooth margins; 

isabelline to patches of fuscous-black due to the absence of aerial 

mycelium, which collapses with age (surface); reverse fuscousblack. 

Colonies reaching 50 mm diam after 1 mo at 25 °C in the 

dark; colonies fertile. 

Specimen examined: Netherlands, Baarn, Maarschalksbosch, decaying leaves of 

Fagus sylvatica (Fagaceae), 1 Oct. 1984, G.S. de Hoog, holotype CBS H-10366, 

culture ex-type CBS 621.84 = ATCC 200937. 

Notes: Isolate CBS 621.84 was until recently preserved at the 

CBS as representative of Cladosporium nigrellum Ellis & Everh., a 

species known from bark of Robinia sp. in the U.S.A. Morphologically 

it is, however, quite distinct in having somewhat larger, and more 

subcylindrical to ellipsoid conidia. Conidia of C. nigrellum are 

fusiform to limoniform, 0–3-septate, 5–15 × 4–7 µm (Ellis 1976), 

possessing the typical cladosporioid scars with a central convex 

dome and a periclinal rim which characterise it as a true member 

of the genus Cladosporium Link, which has been confirmed by a 

re-examination of type material of C. nigrellum (on inner bark of 

railroad ties, U.S.A., West Virginia, Fayette Co., Nuttallburg, 20 

Oct. 1893, L.A. Nuttall, Flora of Fayette County No. 172, NY; also 

Ellis & Everh., N. Amer. Fungi 3086 and Fungi Columb. 382, BPI, 

NY, PH). 

Fusicladium intermedium (Crous & W.B. Kendr.) Crous, comb. 

nov. Mycobank MB504538. Fig. 23. 

Basionym: Anungitopsis intermedia Crous & W.B. Kendr. S. Afr. J. 

Bot. 63: 286. 1997. 

Specimens examined: South Africa, Mpumalanga, from leaf litter of Eucalyptus 

sp. (Myrtaceae), Oct. 1992, M.J. Wingfield, PREM 51438 holotype. Madagascar, 

Tamatave, Eucalyptus leaf litter, Apr. 1994, P.W. Crous, CBS H-19918, epitype 

designated here, culture ex-epitype CPC 778 = IMI 362702 = CBS 110746. 

Note: Conidiophores are dimorphic in culture, being macronematous, 

anungitopsis-like, and micronematous, more pseudocladosporiumlike. 

Fusicladium matsushimae (U. Braun & C.F. Hill) Crous, U. Braun 

& K. Schub., comb. nov. Mycobank MB504539. 

Basionym: Pseudocladosporium matsushimae U. Braun & C.F. Hill, 

Australas. Pl. Pathol. 33: 492. 2004. 


209

Crous et al. 

Fig. 24. Fusicladium pini (CBS 463.82). A–F. Conidiogenous cells with conidiogenous loci. G. Conidia. Scale bars = 10 µm. 

Fusicladium mandshuricum (M. Morelet) Ritschel & U. Braun, 

Schlechtendalia 9: 62. 2003. 

Basionym: Pollaccia mandshurica M. Morelet, Ann. Soc. Sci. Nat. 

Archéol. Toulon Var 45(3): 218. 1993. 

= Pollaccia sinensis W.P. Wu & B. Sutton, in herb. (IMI). 

Teleomorph: Venturia mandshurica M. Morelet, Ann. Soc. Sci. 

Nat. Archéol. Toulon Var 45(3): 219. 1993. 

In vivo: Schubert et al. (2003: 62). 

In vitro on OA: Mycelium loosely branched, filiform to narrowly 

cylindrical-oblong, 1–4 µm wide, later somewhat wider, up to 

7 µm, septate, sometimes slightly constricted at the septa, 

sometimes irregular in outline due to small swellings, subhyaline 

to pale brown, smooth, walls unthickened, sometimes aggregating, 

forming compact conglomerations of slightly swollen hyphal cells. 

Conidiophores usually reduced to conidiogenous cells, arising 

terminally or laterally from hyphae, subcylindrical to cylindrical, 

unbranched, 9–20 × (2.5–)4–5(–6) µm, aseptate, very rarely 1- 

septate, very pale brown, smooth, walls unthickened, monoblastic, 

unilocal, determinate, later occasionally becoming percurrent, 

enteroblastically proliferating, forming a few (up to five) annellations, 

loci broadly truncate, (2–)3–5 µm wide, unthickened, not darkened. 

Conidia solitary, straight to curved, fusiform to obclavate, distinctly 

apiculate, 24–45(–57) × (6–)7–9(–10.5) µm, (1–)2–4(–5)-septate, 

more or less constricted at septa, sometimes up to 85 µm long 

with up to 7 septa, septa often somewhat darkened, second cell 

often bulging, pale medium to medium olivaceous-brown or brown, 

smooth, walls somewhat thickened, somewhat attenuated towards 

the base, hilum broadly truncate, (2–)3–5 µm wide, unthickened, 

not darkened; microcyclic conidiogenesis not observed. 

Cultural characteristics: Colonies on OA iron-grey to olivaceousgrey 

due to aerial mycelium and sporulation (surface); reverse 

iron-grey to black, somewhat velvety; margin glabrous, olivaceous; 

aerial mycelium sparsely formed, loose, diffuse; sporulating. 

Specimens examined: China, Liaoning, on Populus simonii × P. nigra, 17 Jun. 1992, 

M. Morelet, holotype PC (PFN 1466); P. simonii, 20 Apr. 1993, epitype designated 

here CBS H-19912, culture ex-epitype CBS 112235 = CPC 3639 = MPFN 307. 

Note: Conidiophores are densely fasciculate in vivo, forming 

sporodochial conidiomata, cylindrical to ampulliform, 5–7 × 6–7.5 

µm (Schubert et al. 2003). 

Fusicladium pini Crous & de Hoog, sp. nov. MycoBank MB504540. 

Fig. 24. 

Etymology: Named after its host, Pinus. 

Fusicladio africano similis, sed conidiis minoribus, (6–)10–12(–17) × 1.5–2(–2.5) 

µm, locis conidiogenis et hilis angustioribus, 0.5–1 µm latis. 

Mycelium consisting of smooth, medium brown, branched, 

1.5–2 µm wide hyphae, giving rise to solitary, micronematous 

conidiophores. Conidiophores reduced to conidiogenous cells, 

medium to dark brown, erect, thick-walled, smooth, subcylindrical, 

widest at the base, tapering to a subtruncate apex, 5–15 × 2–3 

µm; scars flat-tipped, somewhat darkened and thickened, one to 

several in the apical region, somewhat protruding, 0.5–1 µm wide. 

Conidia in branched or unbranched chains of up to 15, medium 

brown, smooth, subcylindrical, 0–1-septate, widest in the middle, 

tapering to subtruncate ends, straight to slightly curved, (6–)10– 

12(–17) × 1.5–2(–2.5) µm; hila somewhat darkened and thickened, 

not refractive, 0.5–1 µm wide. 


mycelium and smooth margins on PDA, greyish sepia (surface); 

reverse fuscous-black; on OA patches of greyish sepia and fuscousblack 

(surface); on SNA umber (surface). Colonies reaching 15 mm 

diam on PDA after 1 mo at 25 °C in the dark; colonies fertile. 

210


Fig. 25. Fusicladium ramoconidii (CBS 462.82). A. Colony on OA. B. Hyphal coil. C–F. Conidiophores reduced to conidiogenous cells. G–H. Conidiophores. I. Conidia. Scale 


Specimen examined: Netherlands, Baarn, De Vuursche, needle of Pinus sylvestris 

(Pinaceae), 12 Apr. 1982, G.S. de Hoog, holotype CBS H-1610, culture ex-type 


Notes: This fungus was originally maintained in the CBS collection 

as Anungitea uniseptata Matsush. In culture, however, only a 

pseudocladosporium-like state was observed. Conidiophores are 

reduced to conidiogenous cells, and have several apical loci as 

in Fusicladium, but are not subdenticulate; scars are somewhat 

darkened and thickened, not refractive. Conidia of F. africanum are 

(8–)11–17(–20) × 2–3(–3.5) µm, thus similar, but somewhat larger 

than the mean conidial size range (10–12 × 1.5–2 µm) observed 

in F. pini. The conidiogenous loci and conidial hila of F. africanum 

are also somewhat larger. Although the LSU sequence of F. pini is 

identical to that of F. ramoconidii, the ITS sequence similarity is 97 

% (572/585 nucleotides). 

Fusicladium ramoconidii Crous & de Hoog, sp. nov. MycoBank 

MB504541. Figs 25–26. 

Etymology: Named after the presence of its characteristic 

ramoconidia. 

Fusicladio brevicatenato similis, sed ramoconidiis minoribus, (12–)15–17(–20) × 

2(–3) µm, locis conidiogenis et hilis minoribus, 0.5–1 µm diam. 

Mycelium consisting of branched, septate, 1.5–2 µm wide hyphae, 

pale brown, smooth, frequently with hyphal coils. Conidiophores 

integrated into hyphae, and reduced to small, lateral protruding 

conidiogenous cells, concolorous with hyphae, or macronematous, 

dark brown, erect, thick-walled, 10–40 × 3–4 µm, 0–3-septate. 

Conidiogenous cells terminal, integrated, subcylindrical, tapering to 

a rounded apex, concolorous with hyphae (as hyphal pegs), or dark 


211

Crous et al. 

Fusicladium rhodense Crous & M.J. Wingf., sp. nov. MycoBank 

MB504542. Fig. 27. 

Etymology: Named after the Greek Island, Rhodos, where it was 

collected. 

Fusicladio africano similis, sed locis conidiogenis angustioribus, 1.5–2 µm latis, et 

differt a F. pini ramoconidiis formantibus. 

Mycelium consisting of smooth to finely roughened, medium brown, 

branched, septate, 1.5–3 µm wide hyphae, frequently forming 

hyphal coils, giving rise to solitary, micronematous conidiophores. 

Conidiophores reduced to conidiogenous cells that are terminal 

or lateral on hyphae, medium brown, smooth, subcylindrical, 

subdenticulate, erect, or more distinct, up to 15 µm tall, 1.5–2 

µm wide, mono- to polyblastic; scars flat-tipped, somewhat 

darkened and thickened, but not refractive. Conidia in branched 

or unbranched chains of up to 15, pale brown in younger conidia, 

becoming medium brown, smooth, subcylindrical, 0–3-septate, 

tapering slightly towards the subtruncate ends, straight, but at times 

slightly curved, (8–)12–16(–20) × (2–)2.5–3(–4) µm; ramoconidia 

(0–)1(–3)-septate, 12–20 × 3–4 µm; conidia (0–)1-septate, 8–17 

× 2–3 µm; hila somewhat darkened and thickened, not refractive, 

1–1.5 µm wide. 

Fig. 26. Fusicladium ramoconidii (CBS 462.82). Conidiogenous cells with 

ramoconidia and conidia. Scale bar = 10 µm. 

brown on mononematous conidiophores, smooth, 3–15 × 2–3(–4) 

µm; proliferating sympodially, loci slightly thickened, darkened and 

refractive, 0.5–1 µm wide. Conidia occurring in branched chains, 

narrowly ellipsoid to subcylindrical, pale olivaceous, guttulate; 

ramoconidia (0–)1(–3)-septate, (12–)15–17(–20) × 2(–3) µm; 

conidia occurring in short chains (–15), 0–1-septate, (8–)10–12(– 

16) × 2(–3) µm; hila slightly thickened and darkened, not refractive, 

0.5–1 µm wide. 


mycelium and smooth margins on PDA, hazel to fawn (surface), 

with a thin, submerged margin; reverse brown-vinaceous; on OA 

hazel to fawn (surface) with a wide, fawn, submerged margin. 

Colonies reaching 25 mm diam on PDA after 1 mo at 25 °C in the 

dark; colonies fertile. 

Specimen examined: Netherlands, Baarn, De Vuursche, needle of Pinus sp. 

(Pinaceae), 12 Apr. 1982, G.S. de Hoog, holotype CBS H-19908, culture ex-type 


Notes: This strain has been deposited in the CBS collection as 

Pseudocladosporium hachijoense (Matsush.) U. Braun. However, 

its ramoconidia and conidia are smaller than those cited by 

Matsushima (1975) (ramoconidia up to 30 µm long, conidia 10–21 

× 2–4 µm). Although it clusters with F. pini in the LSU phylogeny, 

there are 13 bp differences in their ITS sequence data. Furthermore, 

F. ramoconidii has ramoconidia which are absent in F. pini, and 

has a faster growth rate, and hazel to fawn colonies, compared 

to the greyish sepia colonies of F. pini. The well-developed, 

septate conidiophores and ramoconidia are reminiscent of F. 

brevicatenatum, which differs, however, by its longer and wider 

ramoconidia, up to 30 × 6(–7) µm, as well as larger conidiogenous 

loci and conidial hila, 1.5–3 µm diam. 

Cultural characteristics: Colonies spreading, somewhat erumpent, 

with moderate aerial mycelium and crenate margins on PDA, 

uneven, greyish sepia (surface), margins fuscous-black; reverse 

fuscous-black; on OA smooth, spreading, with sparse aerial 

mycelium and even, regular margins, greyish sepia; on SNA 

spreading, smooth, even margins, sparse aerial mycelium, greyish 

sepia (surface). Colonies reaching 9 mm diam on PDA after 2 wk at 

25 °C in the dark; colonies fertile. 

Specimen examined: Greece, Rhodos, on branches of Ceratonia siliqua (Fabaceae), 

1 Jun. 2006, P.W. Crous & M.J. Wingfield, holotype CBS H-19910, culture ex-type 

CBS 121641 = CPC 13156. 

Note: Fusicladium rhodense has a typical pseudocladosporiumlike 

morphology in culture, with conidial scars that are somewhat 

darkened and thickened. 

Venturia hanliniana (U. Braun & Feiler) Unter., Mycologia 89: 129. 

1997. 

Basionym: Capronia hanliniana U. Braun & Feiler, Microbiol. Res. 

150: 90. 1995. 

≡ Caproventuria hanliniana (U. Braun & Feiler) U. Braun, in Braun, 

A Monograph of Cercosporella, Ramularia and Allied Genera 

(Phytopathogenic Hyphomycetes) 2: 396. 1998. 

Anamorph: Fusicladium brevicatenatum (U. Braun & Feiler) 

Crous, U. Braun & K. Schub., comb. nov. MycoBank MB504543. 

Basionym: Cladophialophora brevicatenata U. Braun & Feiler, 

Microbiol. Res. 150: 84. 1995. 

≡ Pseudocladosporium brevicatenatum (U. Braun & Feiler) U. Braun, 

in Braun, A Monograph of Cercosporella, Ramularia and Allied Genera 

(Phytopathogenic Hyphomycetes) 2: 393. 1998. 

Venturia hystrioides (Dugan, R.G. Roberts & Hanlin) Crous & U. 

Braun, comb. nov. MycoBank MB504544. Fig. 28. 

Basionym: Capronia hystrioides Dugan, R.G. Roberts & Hanlin, 

Mycologia 87: 713. 1995. 

≡ Caproventuria hystrioides (Dugan, R.G. Roberts & Hanlin) U. Braun, 

in Braun, A monograph of Cercosporella, Ramularia and allied genera 

(Phytopathogenic Hyphomycetes). Vol. 2: 396. 1998. 

Anamorph: Fusicladium sp. 

Only the anamorph was observed on OA, PDA and SNA in culture. 

212


Fig. 27. Fusicladium rhodense (CPC 13156). A. Colony on OA. B. Conidial chains and hyphal coil. C–F. Chains of ramoconidia and conidia. Scale bar = 10 µm. 

Fig. 28. Venturia hystrioides (CBS 117727). A. Conidiophores giving rise to catenulate conidia. B. Ramoconidium giving rise to conidia. C–D. Conidial chains. E. Conidia and 

conidiogenous cell with conidiogenous loci. F. Ramoconidium. G. Conidia. Scale bars = 10 µm. 


213

Crous et al. 

Excluded taxa 

Polyscytalum fecundissimum Riess, Bot. Zeitung (Berlin) 11: 

138. 1853. Fig. 29. 

Cultural characteristics: Colonies erumpent, spreading, aerial 

mycelium sparse, margins smooth; colonies sienna to umber on 

PDA, with patches of greyish sepia; reverse chestnut-brown; on 

OA whitish due to moderate aerial mycelium, with diffuse umber 

pigment in the agar; whitish on SNA. Colonies reaching 15 mm 

diam on PDA after 3 wk at 25 °C in the dark. 

Specimen examined: Netherlands, Schovenhorst, leaf litter of Fagus 

sylvatica (Fagaceae), 8 Nov. 1997, W. Gams, CBS H-6049, culture CBS 

100506. 

Fig. 29. Polyscytalum fecundissimum (CBS 100506). Conidiophores giving rise to 

catenulate conidia. Scale bar = 10 µm. 

Mycelium consisting of branched, septate, smooth, guttulate, 

1.5–2.5 µm wide hyphae, pale brown, forming hyphal strands. 

Conidiophores mostly reduced to conidiogenous cells, or if 

present, micronematous, consisting of a supporting cell, and single 

conidiogenous cell. Conidiogenous cells integrated in hyphae as 

lateral loci, or terminal, frequently disarticulating, subcylindrical, 

pale to medium brown, smooth, mono- to polyblastic, loci 1–1.5 

µm wide, 2.5 µm tall; conidiogenous cells subcylindrical, up to 

40 µm tall, and 2–2.5 µm wide. Conidia in long chains of up to 

60, branched or not, subcylindrical to narrowly ellipsoid, pale 

olivaceous to pale brown, smooth; ramoconidia 0–1(–3)-septate, 

15–20(–30) × 2–3(–3.5) µm; conidia 0(–1)-septate, 6–8(–12) × 

2–3(–3.5) µm; hila 1–1.5 µm wide, inconspicuous to somewhat 

darkened, subtruncate. 


mycelium on PDA, and smooth, even margins; olivaceous-grey 

to iron-grey (surface); reverse greenish black; on OA dark mousegrey 

(surface), with even, smooth margins. Colonies reaching 40 

mm diam after 2 wk at 25 °C in the dark; colonies fertile. 

Specimen examined: U.S.A., Washington, Wenatchee, on bing cherry fruit, Prunus 

avium cv. Bing (Rosaceae), R.G. Roberts, culture ex-type, ATCC 96019 = CBS 

117727. 

Note: Dugan et al. (1995) commented that although similar to 

“Phaeoramularia” hachijoensis, the conidia of this species were 

predominantly aseptate and somewhat shorter than those described 

by Matsushima (1975). 

Notes: Polyscytalum fecundissimum is the type species of the genus 

Polyscytalum. Several isolates of this species were investigated 

here to determine if Polyscytalum would be available for taxa that 

have a pseudocladosporium-like morphology. The clustering of 

CBS 681.74 within the Venturiaceae was surprising. However, this 

culture proved to be sterile, and therefore its identity could not be 

confirmed. 

Isolate CBS 109882 sporulated profusely. Colonies were greyolivaceous 

with olivaceous margins on PDA; conidiophores pale, 

and not dark brown as depicted for Polyscytalum in Ellis (1971); 

conidial chains were greenish yellow in mass, and pale olivaceousgreen 

under the dissecting microscope, somewhat roughened, 

polyblastic; on ITS sequence this isolate is identical to U57492, 

Cistella acuum (Alb. & Schwein.) Svrček (Helotiales), but the latter 

species should have a phialidic anamorph, so it is possible that 

this GenBank sequence is incorrect. The identity of CBS 109882 

therefore remains unresolved. 

Although isolate CBS 100506 is poorly sporulating, illustrations 

made in vitro when it was collected show this isolate to be 

authentic for the species and the genus Polyscytalum. Based on 

its LSU sequence, it is allied to Phlogicylindrium eucalypti Crous, 

Summerb. & Summerell (CBS 120080; Summerell et al. 2006), and 

is therefore unrelated to the Venturiaceae. 

Zeloasperisporium R.F. Castañeda, Mycotaxon 60: 285. 1996, 

emend. 

Hyphomycetes. Mycelium mostly superficial, hyphae septate, brown 

to olivaceous. Hyphopodia absent. Conidiophores differentiated, 

mononematous, erect, aseptate or septate, brown to olivaceous. 

Conidiogenous cells integrated, terminal, proliferation sympodial, 

polyblastic, with subdenticulate, somewhat thickened and darkened 

scars. Conidia solitary, fusiform to obclavate or cylindrical, septate, 

asperulate to verrucose, olivaceous to brown, tips always hyaline, 

thinner-walled and smooth, forming mucoid appandages, often only 

visible as a thickened frill. Synanamorph present, micronematous. 

Conidiogenous cells short cylindrical, antenna or hyphopodiumlike, 

phialidic, colarette sometimes present, aseptate, subhyaline. 

Conidia solitary, obovoid, ellipsoid, aseptate, brown to olivaceous, 

verruculose. 

Zeloasperisporium hyphopodioides R.F. Castañeda, Mycotaxon 

60: 285. 1996. Fig. 30. 

In vitro on OA: Mycelium internal to superficial, unbranched to 

sparingly branched, 1.5–3 µm wide, loosely septate, septa almost 

invisible, pale brown, smooth to asperulate, minutely verruculose, 

walls unthickened, sometimes inflated at the base of conidiophores. 

214


Fig. 30. Zeloasperisporium hyphopodioides (CBS 218.95). A–B. Conidiogenous cells. C. Conidia with apical mucoid caps. D. Conidiogenous cell with sympodial proliferation. 

E–G. Conidiogenous cells of micronematous synanamorph. H. Conidia, and microconidia of synanamorph. Scale bars = 10 µm. 

Conidiophores macronematous, arising usually laterally from 

plagiotropous hyphae, erect, straight, subcylindrical or conical, 

not geniculate, usually unbranched, rarely branched, 13–45 × 

3–4(–5) µm, slightly to distinctly attenuated towards the apex, 

tapered, aseptate, rarely with a single septum, pale brown to pale 

medium brown, smooth or minutely verruculose, walls unthickened, 

often somewhat constricted near the base. Conidiogenous cells 

integrated or conidiophores usually reduced to conidiogenous 

cells, subcylindrical to conical, proliferation sympodial, with a single 

or several subdenticulate to denticulate conidiogenous loci mostly 

crowded at or towards the apex, protuberant, truncate, 0.8–1.2 µm 

wide, thickened and darkened-refractive. Conidia solitary, straight 

to curved, ellipsoid, fusiform to obclavate, distinctly tapered towards 

the apex, apiculate, (12–)15–32 × 3.5–5.5 µm, (0–)1–2(–3)-septate, 

mainly 1-septate, usually constricted at the septa, pale brown to 

pale medium brown, asperulate to verruculose, walls unthickened or 

almost so, tips always hyaline, thinner-walled and smooth, forming 

mucoid appendages, often only visible as a thickened frill, base 

somewhat rounded or slightly bulbous, hila often situated on short 

peg-like prolongations, truncate, 0.8–1(–1.2) µm wide, thickened, 

darkened-refractive; microcyclic conidiogenesis occurring, conidia 

forming secondary conidiophores. 

Synanamorph micronematous. Conidiophores reduced 

to conidiogenous cells, numerous, occurring as short lateral 

prolongations of hyphae, antenna or telescope-like, cylindrical, 


unbranched, conidiogenesis unclear, at times appearing 

phialidic, or having one to two apical scars; up to 5 µm long, 

1–1.5 µm wide, aseptate, subhyaline, smooth. Conidia of the 

micronematous anamorph quite different from the conidia formed 

by the macronematous conidiophores, solitary, obovoid, ellipsoid 

to somewhat fusiform, 5–9 × 2.5–3 µm, aseptate, pale to pale 

medium brown, verruculose, somewhat attenuated towards the 

base, hila flat, unthickened to somewhat thickened, appearing to 

have the ability to form a slime appendage at the apex. 

Cultural characteristics: Colonies on OA iron-grey to olivaceous due 

to abundant sporulation (surface); reverse black, velvety; margin 

regular to undulate, feathery; aerial mycelium absent or sparse, 

sporulation profuse. 

Specimen examined: Cuba, isolated from air, 2 Oct. 1994, R.F. Castañeda, INIFAT 

C94/114, holotype, CBS-H 5624, H-5639, isotypes, culture ex-type CBS 218.95 = 

INIFAT C94/114 = MUCL 39155 = IMI 367520. 

Notes: Within the course of the recent phylogenetic studies 

in Herpotrichiellaceae and Venturiaceae the type culture of 

Zeloasperisporium hyphopodioides has been included since it was 

deposited at the CBS as “Fusicladium hyphopodioides”. When 

the culture was re-examined, the described short appressoriumlike, 

inflated hyphopodia with slightly warted to lobed apices 

(Castañeda et al. 1996) could be recognised as conidiogenous 

cells of a synanamorph forming a second conidial type. In addition, 

215

Crous et al. 

the conidial tips are hyaline, unthickened and smooth, and have the 

ability to form mucoid appendages that are often only visible as a 

thickened frill. These two features, viz., the synanamorph and the 

conidia with mucoid appendages, easily distinguish this genus from 

morphologically similar genera such as Fusicladium, Asperisporium 

Maubl., and Passalora Fr. Phylogenetically Zeloasperisporium 

clusters basal to the Venturiaceae. 

Discussion 

The present paper was initiated to clarify the status of 

Cladophialophora and Pseudocladosporium spp., which appear 

morphologically similar. Confusion occurs when strains with this 

morphology are identified based solely on microscopic and cultural 

characteristics. The results clarify that Cladophialophora is allied to 

the Herpotrichiellaceae and Pseudocladosporium (= Fusicladium) 

to the Pleosporales (Dothideomycetes). The plant-pathogenic 

Cladophialophora species compose a separate clade within the 

order (Fig. 1). Another, somewhat remote chaetothyrialean clade 

contains extremotolerant, rock-inhabiting species around the 

genus Coniosporium Link (Cluster 5 of Haase et al. 1999). Both 

clades are significantly distinct from the prevalently hyperparasitic 

or oligotrophic, frequently opportunistic species of the remainder of 

the order (Fig. 1). This remainder includes all Capronia teleomorphs 

sequenced to date, and is thus likely to represent the family 

Herpotrichiellaceae. The ecological trends in each of the main 

clades of Chaetothyriales are thus quite different (Braun 1998). 

Several novelties are introduced within the preponderantly plantassociated 

clade of Chaetothyriales, including two new species 

associated with leaf spots. Cladophialophora is distinguished from 

Polyscytalum, which clusters outside the Herpotrichiellaceae, and 

appears allied to Phlogicylindrium Crous, Summerb. & Summerell, 

a recently introduced genus for species occurring on Eucalyptus 

leaves (Summerell et al. 2006). Surprisingly Heteroconium 

chaetospira clusters in the Herpotrichiellaceae, and is placed 

in Cladophialophora as a distinctively pigmented member of the 

genus. Some species of Cladophialophora and Exophiala are 

newly described from a range of substrates such as fruit juices, 

drinking water and leaf litter, revealing the potential of these 

materials as ecological sources of inoculum for taxa associated 

with opportunistic human and animal infections. 

Furthermore, Pseudocladosporium belongs to the Venturiaceae, 

and is best treated as a synonym of Fusicladium, along with other 

genera as proposed by Schubert et al. (2003) and Beck et al. (2005). 

Although numerous isolates of the Venturiaceae were included for 

study, it was surprising to find relatively little variation within the 

family, suggesting that previously proposed teleomorph genera 

such as Apiosporina, Metacoleroa and Caproventuria should be 

best treated as synonyms of Venturia. The Venturiaceae is further 

extended with the inclusion of a novel sister clade of hyphomycetes 

with a pseudocladosporium-like morphology, which are also 

referred to as Fusicladium, thus widening the generic concept of 

the latter to encompass all pseudocladosporium-like anamorphs 

within the family. Some species assigned to Anungitopsis proved 

to cluster within the Venturiaceae, but the type species of the 

latter genus, A. speciosa, clustered elsewhere and possesses 

distinct conidiogenous loci, i.e., Anungitopsis cannot be reduced 

to synonymy with Fusicladium. The anamorphs of this sister clade 

of the main Venturia clade are morphologically rather close to 

taxa assigned to Anungitea. However, species of Anungitea and 

Fusicladium are morphologically barely distinguishable (Schubert 

et al. 2003), but the true affinity of Anungitea depends on its type 

species of which cultures and sequence data are not yet available. 

Several anamorph genera with divergent morphologies were 

found to cluster together, suggesting that these are either different 

synanamorphs of the same teleomorph genus, or that they may 

represent cryptic clades that will diverge further once additional 

species are added in future studies. Although the Herpotrichiellaceae 

appeared to represent quite a diverse assembledge of morphotypes, 

the Venturiaceae were again surprisingly uniform. 


Several colleagues from different countries provided material without which this work 

would not have been possible. We thank M. Vermaas for preparing the photographic 

plates, H.-J. Schroers for some gDNA and sequences used in this work, and A. 

van Iperen for maintaining the cultures. K. Schubert was financially supported by 

a Synthesys grant (No. 2559) which is gratefully acknowledged. R.C. Summerbell 

is thanked for his comments on an earlier version of the script. L. Hutchison is 

acknowledged for collecting Apiosporina collinsii for us. 

References 

Barr ME (1987). Prodomus to class Loculoascomycetes. University of Massachusetts, 

Amherst, Massachusetts. 

Beck A, Ritschel A, Schubert K, Braun U, Triebel D (2005). Phylogenetic relationships 

of the anamorphic genus Fusicladium s. lat. as inferred by ITS nrDNA data. 

Mycological Progress 4: 111–116. 


(Phytopathogenic Hyphomycetes). Vol. 2. IHW-Verlag, Eching. 



the teleomorph of Cladosporium s.str. Mycological Progress 2: 3–18. 

Braun U, Feiler U (1995). Cladophialophora and its teleomorph. Microbiological 

Research 150: 81–91. 

Cai L, McKenzie EHC, Hyde KD (2004). New species of Cordana and Spadicoides 

from decaying bamboo culms in China. Sydowia 56: 222–228. 

Castañeda RF, Fabré DE, Parra MP, Perez M, Guarro J, Cano J (1996). Some 

airborne conidial fungi from Cuba. Mycotaxon 60: 283–290. 

Castañeda RF, Kendrick B (1990a). Conidial fungi from Cuba I. University of 

Waterloo Biological Series 32: 1–53. 

Castañeda RF, Kendrick B (1990b). Conidial fungi from Cuba II. University of 

Waterloo Biological Series 33: 1–61. 

Crous PW (2002). Adhering to good cultural practice (GCP). Mycological Research 

106: 1378–1379. 



Crous PW, Denman S, Taylor JE, Swart L, Palm ME (2004). Cultivation and diseases 

of Proteaceae: Leucadendron, Leucospermum and Protea. CBS Biodiversity 

Series 2: 1–228. 

Crous PW, Groenewald JZ, Wingfield MJ (2006b). Heteroconium eucalypti. Fungal 

Planet No. 10. (www.fungalplanet.org). 

Crous PW, Mohammed C, Glen M, Verkley GJM, Groenewald JZ (2007b). 

Eucalyptus microfungi known from culture. 3. Eucasphaeria and Sympoventuria 

genera nova, and new species of Furcaspora, Harknessia, Heteroconium and 

Phacidiella. Fungal Diversity 25: 19–36. 


A, Burgess T, Barber P, Groenewald JZ (2006a). Phylogenetic lineages in the 


Decock C, Delgado-Rodriguez G, Buchet S, Seng JM (2003). A new species and 

three new combinations in Cyphellophora, with a note on the taxonomic affinities 

of the genus, and its relation to Kumbhamaya and Pseudomicrodochium. 

Antonie van Leeuwenhoek 84: 209–216. 

Dugan FM, Roberts RG, Hanlin RT (1995). New and rare fungi from cherry fruit. 

Mycologia 87: 713–718. 



Ellis MB (1976). More dematiacous hyphomycetes. Commonwealth Mycological 


216


Gams W, Verkley GJM, Crous PW (2007). CBS course of mycology, 5 th ed. 



by SSU gene analysis of members of the Herpotrichiellaceae, with special 


Ho MH-M, Castañeda RF, Dugan FM, Jong SC (1999). Cladosporium and 


72: 115–157. 

Honbo S, Padhy AA, Ajello L (1984). The relationship of Cladosporium carrionii to 

Cladophialophora ajelloi. Sabouraudia 22: 209–218. 

Hoog GS de (1985). Taxonomy of the Dactylaria complex, IV. Dactylaria, Neta, 

Subulispora and Scolecobasidium. Studies in Mycology 26: 1–60. 



Hoog GS de, Guarro, J, Gené J, Figueras MJ (2000). Atlas of clinical fungi, 2 nd 

ed. Centraalbureau voor Schimmelcultures, Utrecht and Universitat Rovira I 

Virgili, Reus. 

Hoog GS de, Nishikaku AS, Fernandez Zeppenfeldt G, Padín-González C, Burger 

E, Badali H, Gerrits van den Ende AHG (2007). Molecular analysis and 

pathogenicity of the Cladophialophora carrionii complex, with the description 

of a novel species. Studies in Mycology 58: 219–234. 

Hoog GS de, Takeo K, Göttlich E, Nishimura K, Miyaji M (1995). A human isolate of 

Exophiala (Wangiella) dermatitidis forming a catenate synanamorph that links 

the genera Exophiala and Cladophialophora. Journal of Medical and Veterinary 

Mycology 33: 355–358 

Jacob M, Bhat DJ (2000). Two new endophytic fungi from India. Cryptogamie 

Mycologie 21: 81–88. 

Kirk PM (1982). New or interesting microfungi. IV. Dematiaceous hyphomycetes 

from Devon. Transactions of the British Mycological Society 78: 55–74. 

Kirk PM (1983). New or interesting microfungi. IX. Dematiaceous hyphomycetes 

from Esher Common. Transactions of the British Mycological Society 80: 

449–467. 

Levin TP, Baty DE, Fekete T, Truant AL, Suh B (2004). Cladophialophora bantiana 

brain abscess in a solid-organ transplant recipient: case report and review of 

the literature. Journal of Clinical Microbiology 42: 4374–4378. 

Madan V, Bisset D, Harris P, Howard S, Beck MH (2006). Phaeohyphomycosis 

caused by Exophiala salmonis. British Journal of Dermatology 155: 1082– 

1084. 

Marvanová L, Laichmanová M (2007). Subilispora biappendiculata, anamorph sp. 

nov. from Borneo (Malaysia) and a review of the genus. Fungal Diversity 26: 

241–256. 

Matos T, Hoog GS de, Boer AG de, Crom I de, Haase G (2003). High prevalence of 

the neurotropic black yeast Exophiala (Wangiella) dermatitidis in steam baths. 

Mycoses 45: 373–377. 

Matsushima T (1975). Icones Microfungorum a Matsushima Lectorum. Kobe. 




Partridge EC, Morgan-Jones G (2003). Notes on hyphomycetes. XC. 

Fusicladosporium, a new genus for cladosporium-like anamorphs of Venturia. 

Mycotaxon 85: 357–370. 

Petrak F (1949). Neue Hyphomyzeten-Gattungen aus Ekuador. Sydowia 3: 259– 

266. 


Kew. 



98: 625–634. 

Richards RH, Holliman A, Helgason S (1978). Exophiala salmonis infection in 

Atlantic salmon Salmo salar L. Journal of Fish Diseases 1: 357–368. 

Saccardo PA (1877). Michelia Commentarium Mycologicum Fungos in Primis 

Italicos Illustrans. Volume 1: 1–116. Padua, Italy. 

Schubert K, Braun U (2002a). Fusicladium scillae. IMI Descriptions of Fungi and 

Bacteria 152: 1518. 

Schubert K, Braun U (2002b). Fusicladium convolvularum. IMI Descriptions of Fungi 

and Bacteria 152: 1513. 



Sivanesan A (1984). The bitunicate ascomycetes and their anamorphs. Cramer 

Verlag, Vaduz. 

Summerell BA, Groenewald JZ, Carnegie AJ, Summerbell RC, Crous PW (2006). 

Eucalyptus microfungi known from culture. 2. Alysidiella, Fusculina and 

Phlogicylindrium genera nova, with notes on some other poorly known taxa. 


Sutton BC (1973). Hyphomycetes from Manitoba and Saskatchewan, Canada. 

Mycological Papers 132: 1–143. 

Sutton BC (1975). Hyphomycetes on cupules of Castanea sativa. Transactions of 



Capronia with notes on anamorph-teleomorph connections. Mycologia 89: 

120–131. 

Untereiner WA (2000). Capronia and its anamorphs: exploring the value 

of morphological and molecular characters in the systematics of the 

Herpotrichiellaceae. Studies in Mycology 45: 141–149. 

Varghese KIM, Rao VG (1979). Forest microfungi – I. Subramaniomyces, a new 

genus of hyphomycetes. Kavaka 7: 83–85. 




White TJ, Bruns T, Taylor J (1990). Amplification and direct sequencing of fungal 

ribosomal RNA genes for phylognetics. In: A Guide to Molecular Methods and 

Applications (Innis MA, Gelfand DH, Sninsky JJ, White JW, eds). Academic 

Press, New York: 315–322. 

Zeng J-S, Sutton DA, Fothergill AW, Rinaldi MG, Harrak MJ, Hoog GS de (2007). 

Spectrum of clinically relevant Exophiala species in the U.S.A. Journal of 

Clinical Microbiology: in press. 


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doi:10.3114/sim.2007.58.08 


Molecular analysis and pathogenicity of the Cladophialophora carrionii complex, 

with the description of a novel species 

G.S. de Hoog 1,2* , A.S. Nishikaku 3 , G. Fernandez-Zeppenfeldt 4 , C. Padín-González 4 , E. Burger 3 , H. Badali 1 , N. Richard-Yegres 4 and A.H.G. 

Gerrits van den Ende 1 

1 

CBS Fungal Biodiversity Centre, Utrecht, The Netherlands; 2 Institute for Biodiversity and Ecosystem Dynamics, University of Amsterdam, Amsterdam, The Netherlands; 

3 

Laboratory of Immunopathology of Mycosis, Department of Immunology, São Paulo University, São Paulo, Brazil; 4 National Experimental University “Francisco de Miranda” 

(UNEFM), Coro, Venezuela 

*Correspondence: Sybren de Hoog, s.hoog@cbs.knaw.nl 

Abstract: Cladophialophora carrionii is one of the four major etiologic agents of human chromoblastomycosis in semi-arid climates. This species was studied using sequence 

data of the internal transcribed spacer region of rDNA, the partial β-tubulin gene and an intron in the translation elongation factor 1-alpha gene, in addition to morphology. 

With all genes a clear bipartition was observed, which corresponded with minute differences in conidiophore morphology. A new species, C. yegresii, was introduced, which 

appeared to be, in contrast to C. carrionii, associated with living cactus plants. All strains from humans, and a few isolates from dead cactus debris, belonged to C. carrionii, for 

which a lectotype was designated. Artificial inoculation of cactus plants grown from seeds in the greenhouse showed that both fungi are able to persist in cactus tissue. When 

reaching the spines they produce cells that morphologically resemble the muriform cells known as the “invasive form” in chromoblastomycosis. The tested clinical strain of C. 

carrionii proved to be more virulent in cactus than the environmental strain of C. yegresii that originated from the same species of cactus, Stenocereus griseus. The muriform 

cell expressed in cactus spines can be regarded as the extremotolerant survival phase, and is likely to play an essential role in the natural life cycle of these organisms. 

Taxonomic novelty: Cladophialophora yegresii de Hoog, sp. nov. 

Key words: Cactus, chromoblastomycosis, Cladophialophora, endophyte, extremotolerance, phylogeny, taxonomy. 

INTRODUCTION 

Cladophialophora carrionii (Trejos) de Hoog, Kwon-Chung & 

McGinnis is one of the most frequent etiologic agents of human 

chromoblastomycosis, a chronic cutaneous disease characterised 

by verrucose skin lesions eventually leading to emerging, cauliflowerlike 

eruptions. The species is particularly observed in arid and semiarid 

climates of e.g. South and Central America (Lavelle 1980) and 

Australia (Trejos 1954, Riddley 1957). The current hypothesis is that 

patients suffering from chromoblastomycosis are rural workers who 

acquire the infection after being pricked by cactus thorns or splinters 

(Rubin et al. 1991, Fernández-Zeppenfeldt et al. 1994). A classical 

case reported by O’Daly (1943) concerns traumatic inoculation with 

thorns of “guazábara” (Opuntia caribaea), a common xerophilic 

plant in semi-arid Venezuela. This hypothesised traumatic route of 

infection was later supported by Richard-Yegres & Yegres (Richard- 

Yegres & Yegres 1987; strain SR3 = CBS 863.96) and Fernández- 

Zeppenfeldt et al. (1994), who isolated strains from Prosopis 

juliflora litter. Cladophialophora Borelli has also been detected in 

association with spines of the common xerophyte Aloe vera and of 

the Cactaceae: Opuntia caribaeae, O. caracasana, Stenocereus 

griseus and Cereus lanuginosus (Borelli 1972, Yegres et al. 1996). 

Thorny American cacti are important components of the xerophyte 

flora of the arid climate of our study area in Falcon State, Venezuela 

(Richard-Yegres et al. 1992). Muriform cells are produced in 

human skin and represent the supposed pathogenic invasive form 

of fungi causing chromoblastomycosis (Mendoza et al. 1993). 

Early experiments involving the inoculation of several species of 

cold-blooded animals have shown the abundant production of the 

characteristic muriform cells in vivo (Trejos 1953). 

A similar plant origin of chromoblastomycosis has been 

supposed for a related agent of chromoblastomycosis, Fonsecaea 

pedrosoi (Brumpt) Negroni. Marques et al. (2006) isolated this 

species from the shells of Babassu coconuts (Orbignya phalerata). 

The habit of local people to sit on these shells might explain the 

frequent occurrence of lesions on the buttocks (Silva et al. 1995). 

Salgado et al. (2004) found the species on the thorns of a Mimosa 

pudica plant which a patient could identify as the source of traumatic 

onset of his chromoblastomycosis. 

Recently, with the development of molecular tools for species 

identification, doubt has arisen about the correctness of this 

supposed route of infection. The question whether environmental 

and clinical strains represent exactly the same species needs to be 

re-determined. In order to establish this for C. carrionii-associated 

chromoblastomycosis, reference strains from the CBS culture 

collection, supplemented with a large set of strains from semiarid 

Venezuela, have been verified using molecular tools that are 

currently routinely employed to answer taxonomic questions in 

black yeasts and their filamentous relatives (de Hoog et al. 2003), 

particularly the internal transcribed spacer (ITS) region of rDNA, 

the partial β-tubulin gene (BT2), and an intron in the translation 

elongation factor 1-alpha (EF1). In addition, a series of three 

experiments has been conducted concerning inoculation into 

and superficial application onto germlings of Stenocereus griseus 

obtained by cultivation in vitro, mature plants of S. griseus from the 

wild, and in spines of S. griseus collected in the semi-arid area of 

study. Our aim is to reveal the role of the cactus S. griseus in the 

life cycle of its associated Cladophialophora spp., and to determine 

whether a link could be made to C. carrionii for obtaining a better 

understanding of human chromoblastomycosis. 


Fungal strains and morphology 

Strains studied are listed in Table 1. This list comprises strains which 

have morphologically been identified as C. carrionii. Reference 

strains from the CBS culture collection, as well as fresh isolates 

from patients and the environment have been included. Strains 

were lyophilised and stored in liquid nitrogen soon after deposit at 

CBS. Stock cultures for transient working collections were grown 

on slants of 2 % malt extract agar (MEA) and oatmeal agar (OA) at 

24 °C. For morphological observation, slide cultures were made of 

strains grown on potato-dextrose agar (PDA) (de Hoog et al. 2000) 

and mounted in lactophenol cotton blue. 

219

De Hoog et al. 

Table 1. Isolation data of Cladophialophora strains examined. 

Name CBS nr. Other reference(s) 1 GenBank mtDNA* 

(Kawasaki 

ITS, BT2, EF1 et al. 1993) 

Source [human: duration, localization, sex, age 

(Pérez-Blanco et al. 2003)]. 

Geography 

A / I / 1: C. carrionii 

117904 UNEFM 0004-02 = dH 14480 EU137281, –, – Chromoblastomycosis; 14 y; hip, thigh, leg; male 38 Falcon State, Venezuela 

117891 UNEFM 0002-00 = dH 14475 EU137278, –, EU137222 Chromoblastomycosis; 1 y; male 62 Falcon State, Venezuela 

117906 UNEFM 0014-96 = dH 14504 EU137288, EU137171, EU137231 Chromoblastomycosis; 0.5 y; hand; male 45 Falcon State, Venezuela 

117897 UNEFM 0011-03 = dH 14497 EU137314, –, EU137254 Chromoblastomycosis; 0.5 y; hand; male 42 Falcon State, Venezuela 

859,96 UNEFM 9617 = dH 10703 EU137295, EU137178, EU137237 Dry plant debris, arid zone Falcon State, Venezuela 

117898 UNEFM 0010-98 = dH 14496 EU137308, –, EU137246 Chromoblastomycosis; 20 y; hand; female 59 Falcon State, Venezuela 

117889 UNEFM 0003-04 = dH 14478 –, EU137190, – Chromoblastomycosis; 20 y; thigh, leg; female 78 Falcon State, Venezuela 

114392 UNEFM 82267 = dH 13261 EU137267, EU137150, EU137211 Chromoblastomycosis; leg; female Falcon State, Venezuela 

114394 UNEFM 9803 = dH 13263 EU137307, –, EU137245 Chromoblastomycosis; hand; male 22 Falcon State, Venezuela 

114396 UNEFM 2001/1 = dH 13265 EU137269, EU137152, EU137213 Chromoblastomycosis; arm; male 35 Falcon State, Venezuela 

114399 UNEFM 2003/2 = dH 13268 EU137272, EU137155, EU137216 Chromoblastomycosis; arm; female 64 Falcon State, Venezuela 

114401 UNEFM 9901 = dH 13270 EU137274, EU137157, EU137218 Chromoblastomycosis; arm; female 40 Falcon State, Venezuela 

114402** UNEFM 9902 = dH 13271 EU137275, EU137158, EU137219 Chromoblastomycosis; arm; female 40 Falcon State, Venezuela 

114403 UNEFM 95195 = dH 13272 EU137276, EU137159, EU137220 Chromoblastomycosis; arm; male Falcon State, Venezuela 

117899 UNEFM 0010-04 = dH 14495 EU137301, EU137183, EU137241 Chromoblastomycosis; 2 y; hand; male 57 Falcon State, Venezuela 

117901 UNEFM 0009-03 = dH 14492 EU137312, EU137197, EU137252 Chromoblastomycosis; 8 y; arm; female 41 Falcon State, Venezuela 

114393 UNEFM 9801 = dH 13262 EU137268, EU137151, EU137212 Chromoblastomycosis; hand; male 72 Falcon State, Venezuela 

108.97** UNEFM 9501 = dH 10704 EU137306, EU137188, EU137265 Chromoblastomycosis; skin Falcon State, Venezuela 

114397 UNEFM 84020 = dH 13266 EU137270, EU137153, EU137214 Chromoblastomycosis; hand, arm; male 54 Falcon State, Venezuela 

114404 UNEFM 95656 = dH 13273 EU137311, EU137196, EU137251 Chromoblastomycosis; arm; male Falcon State, Venezuela 

117902 UNEFM 0008-03 = dH 14489 EU137283, EU137166, EU137226 Chromoblastomycosis; 3 y; arm; male 42 Falcon State, Venezuela 

117893 UNEFM 0001-00 = dH 14470 EU137316, EU137200, – Chromoblastomycosis; 2 y; knee; male 19 Falcon State, Venezuela 

117892 UNEFM 0001-02 = dH 14471 EU137277, EU137160, EU137221 Chromoblastomycosis; 8 y; knee; male 52 Falcon State, Venezuela 

117908 UNEFM 0013-04 = dH 14502 –, EU137191, – Chromoblastomycosis; 6 y; back; male 13 Falcon State, Venezuela 

109.97** UNEFM 9503 = dH 10706 –, –, – Chromoblastomycosis; skin Falcon State, Venezuela 

857.96 UNEFM 9408 = dH 10707 EU137294, EU137177, EU137236 Chromoblastomycosis; skin Falcon State, Venezuela 

114398 UNEFM 2003/1 = dH 13267 EU137271, EU137154, EU137215 Chromoblastomycosis; arm; female 67 Falcon State, Venezuela 

114400 UNEFM 2003/3 = dH 13269 EU137273, EU137156, EU137217 Chromoblastomycosis; arm; male 50 Falcon State, Venezuela 

117909 UNEFM 0013-00 = dH 14501 EU137287, EU137170, EU137230 Chromoblastomycosis; arm; male Falcon State, Venezuela 

114395 UNEFM 9802 = dH 13264 EU137299, EU137182, EU137240 Chromoblastomycosis; leg; female 22 Falcon State, Venezuela 

166.54 MUCL 10088 EU137290, EU137173, – Skin lesion in human Falcon State, Venezuela 

862.96 UNEFM 9603 = dH 10700 EU137315, EU137199, EU137255 Dry plant debris, semi-arid zone Falcon State, Venezuela 

863.96** IFM 41444 = UNEFM SR3 = dH 10699 AB109169 / EU137296, EU137179, EU137238 Dry spine (Opuntia caribaea) on soil, semi-arid zone Falcon State, Venezuela 

861.96 UNEFM 9607 = dH 10701 EU137309, EU137194, EU137249 Dry plant debris, semi-arid zone Falcon State, Venezuela 

117896 dH 14498 EU137285, –, EU137228 Hand lesion Falcon State, Venezuela 

114397 UNEFM 84020 = dH 13266 EU137270, EU137153, EU137214 Chromoblastomycosis, hand and arm Falcon State, Venezuela 

220

Cladophialophora carrionii complex 

117905 dH 14505 EU137300, –, – Chromoblastomycosis, hand, male Falcon State, Venezuela 

117900 dH 14493 EU137284, –, EU137227 Chromoblastomycosis, hand, male Falcon State, Venezuela 

114392 UNEFM 82267 = dH 13261 EU137267, EU137150, EU137211 Chromoblastomycosis, leg, female Falcon State, Venezuela 

– FMC 248 AF397181, –, – Chromoblastomycosis Venezuela 

– IFM 41807 AB109175, –, – Group mt-I – Venezuela 

– IFM 4812 AB109168, –, – Group mt-I – Venezuela 

– IMTSP 690 AF397180, –, – Chromoblastomycosis Brazil 

410.96 UAMH 4392 = NCMH 1010 = DUKE 2403 EU137310, EU137195, EU137250 Chromoblastomycosis – 

163.54 EU137304, EU137186, EU137243 Chromoblastomycosis Australia 

117903 dH 14482 EU137282, –, EU137225 Chromoblastomycosis, forearm, male – 

362.70 M.J. Campos 4555 = dH 15806 EU137302, EU137184, EU137242 Human Mozambique 

260.83 CDC B-1352 = FMC 282 = ATCC 44535 (ex-T of C. ajelloi) EU137292, EU137175, EU137234 Group mt-I Skin lesion in human Uganda 

986.96 UAMH 5717 EU137297, EU137180, – Clinical material – 

– IFM 4805 AB087204, –, – – – 

– IFM 4811 AB109178, –, – – – 

– IFM 41814 AB109176, –, – – – 

B / II / 2: C. carrionii 

160.54 ATCC 16264 = CDC A-835 = MUCL 40053 = IFM 4808 (ex-LT of C. carrionii) AB109177 / EU137266, EU137201, EU137210 Group mt-II Chromoblastomycosis, human Australia 

– Todd Pryce 200867 = dH 13218 Human Australia 

406.96 MRL 1114 = UAMH 4366 = dH 15847 EU137317, EU137202, EU137256 Human Queensland, Australia 

100434 ATCC 32279 = dH 10745 = IP 518 = RV 16499 EU137289, EU137172, EU137232 Human Madagascar 

– IFM 4810 AB109170, –, – – – 

– IFM 41446 = DCU 606 AB109171, –, – – – 

C: C. carrionii 

– IFM 41651 AB109174, –, – Group mt-I – China 



– IFM 4985 AB109179, –, – – – 

– IFM 4986 AB109180, –, – – – 

D / III / 3: C. yegresii 

114406 UNEFM SgSR1 = dH 13275 EU137323, EU137208, EU137263 Stenocereus griseus asymptomatic plant Falcon State, Venezuela 

114407 UNEFM SgSR2 = dH 13276 EU137324, –, EU137264 Stenocereus griseus asymptomatic plant Falcon State, Venezuela 

114405** UNEFM SgSR3 = dH 13274 (ex-T of C. yegresii) EU137322, EU137209, EU137262 Stenocereus griseus asymptomatic plant Falcon State, Venezuela 

1 Abbreviations: ATCC = American Type Culture Collection, Manassas, U.S.A.; CBS = Centraalbureau voor Schimmelcultures, Utrecht, The Netherlands; CDC = Centers for Disease Control and Prevention, Atlanta, U.S.A.; DCU = Department of 

Dermatology, School of Medicine, Chiba, Japan; dH = G.S. de Hoog working collection; FMC = Faculdade de Medicina, Caracas, Venezuela; ITMSP = Instituto de Medicina Tropical de São Paulo, São Paulo, Brazil; IFM = Research Center for 

Pathogenic Fungi and Microbial Toxicoses, Chiba University, Chiba, Japan; IP = Institut Pasteur, Paris, France; MUCL = Mycotheque de l’Université de Louvain, Louvain-la-Neuve, Belgium; RV = Prince Leopold Institute of Tropical Medicine, Antwerp, 

Belgium; UAMH = The University of Alberta Microfungus Collection and Herbarium, Edmonton, Canada; UNEFM = Universidade Nacional Experimental Francisco de Miranda, Coro, Falcon, Venezuela. 

Ex-T = Type strain; ex-LT = Lectotype strain. 

*Type I and II groups based on mitochondrial DNA restriction fragment length polymorphism: H-1 and H-2 = restriction patterns 1 and 2, respectively, with HaeIII enzyme; M-1 and M-2 = restriction patterns 1 and 2, respectively, with MspI enzyme; S-1 

and S-2 = restriction patterns 1 and 2, respectively, with Sau3AI enzyme. 

** Used in plant and mouse inoculation experiments. 


221


Table 2. Results from MrAic using corrected Akaike Information Criterion (AICc). 

Fragment/Gene Model df* lnL* AICc* wAICc* 

rRNA ITS TrNG 89 -21.840.556 4575.9992” 0.3080 

EF-1α HKYG 88 -19.392.355 41.147.171 0.5242 

ß-Tubulin SYMIG 90 -28.547.745 59.261.933 0.1913 

*df = degrees of freedom; lnL = log likelihood; AICc = corrected AIC; wAICc = weighted corrected AIC. 

DNA extraction 

Approximately 1 cm 2 mycelium of 30-d-old cultures was transferred 

to a 2 mL Eppendorf tube containing 300 µL TES-buffer (Tris 1.2 % 

w/v, Na-EDTA 0.38% w/v, SDS 2 % w/v, pH 8.0) and about 80 mg 

of a silica mixture (Silica gel H, Merck 7736, Darmstadt, Germany 

/ Kieselguhr Celite 545, Machery, Düren, Germany, 2 : 1, w/w). 

Cells were disrupted mechanically in a tight-fitting sterile pestle for 

approximately 1 min. Subsequently 200 µL TES-buffer was added, 

the mixture was vortexed, 10 µL proteinase K was added and 

incubated for 10 min at 65 °C. After addition of 140 µL of 5 M NaCl 

and 1/10 vol CTAB 10 % (cetyltrimethylammoniumbromide) buffer, 

the material was incubated for 30 min at 65 °C. Subsequently 

700 µL SEVAG (24 : 1, chloroform : isoamylalcohol) was mixed to 

solution, incubated during 30 min on ice water and centrifuged for 

10 min at 14 000 rpm. The supernatant was transferred to a new 

tube with 225 µL 5 M NH 4 

-acetate, incubated on ice water and 

centrifuged again for 10 min at 14 000 rpm. The supernatant was 

transferred to another Eppendorf tube with 0.55 vol isopropanol and 

spun for 5 min at 14 000 rpm. Subsequently, the pellet was washed 

with ice cold 70 % ethanol. After drying at room temperature it was 

re-suspended in 48.5 µL TE buffer (Tris 0.12 % w/v, Na-EDTA 0.04 

% w/v) plus 1.5 µL RNAse 20 U/mL and incubated for 15–30 min 

at 37 °C. 

Sequencing and phylogenetic reconstruction 

Three loci, namely the internal transcribed spacers (ITS), ß- 

tubulin (BT2) and translation elongation factor 1-α (EF1), were 

sequenced. For ITS sequencing, amplification was performed 

with V9G (5’-TTACGTCCCTGCCCTTTGTA-3’) and LS266 (5’- 

GCATTCCCAAACAACTCGACTC-3’). Sequencing reactions 

were conducted with ITS1 and ITS4 primers (White et al. 

1990). For BT2 amplification and sequencing, primers Bt2a 

(5’-GGTAACCAAATCGGTGCTGCTTTC-3’) and Bt2b (5’- 

ACCCTCAGTGTAGTGACCCTTGGC-3’) were used (Glass & 

Donaldson 1995) and for EF1 amplification and sequencing, primers 

EF1-728F (5’CATCGAGAAGTTCGAGAAGG-3’) and EF1-986R 

(5’-TACTTGAAGGAACCCTTACC-3’) (Carbone & Kohn, 1999). 

Sequences were aligned in BioNumerics v. 4.5 (Applied Maths, 

Kortrijk, Belgium), exported and converted into Phylip interleaved 

format (Felsenstein 1993). 

Calculation of ILD (incongruence length difference) was 

performed in PAUP v. 4.0b10 (Swofford 2003). A combined data set 

of ITS, EF1 and BT2 sequences was created. Optimality criterion 

was set to parsimony. The total number of characters was 1 263 

with equal weight, while 677 characters were constant, and 396 

parsimony-informative. Gaps were treated as missing, and treebisection-reconnection 

(TBR) was used as branch-swapping 

algorithm. Maximum number of trees was set to 100 and left 

unchanged. 

Substitution model testing 

The program MrAic (www.abc.se/~nylander/; Nylander 2004) 

was used to select a substitution model. MrAic is a Perl script for 

calculating the Akaike Information Criterion (AIC), corrected Akaike 

Information Criterion (AICc), Bayesian Information Criterion (BIC), 

and Akaike weights for nucleotide substitution models and model 

uncertainty. Using an ML algorithm, likelihood scores under different 

models were estimated using Phyml (http://atgc.lirmm.fr/phyml/). All 

56 models implemented in Modeltest (Posada & Crandall 1998) 

were evaluated. These models were also combined with proportion 

of invariable sites (I) and/or gamma distribution shape parameter 

(G). A difference between Modeltest and MrAic is that the latter 

does not evaluate all models on the same, approximate topology 

as in PAUP (Swofford 1981). Instead, Phyml was used to try to find 

the maximum of the likelihood function under all models. This is 

necessary for finding AIC, AICc, or BIC for the models. The AICc 

calculation (Table 2) was used to select the right model for the ratio 

of parameters to characters (Nchar/Nparameters < 40; Burnham & 

Anderson 2002) for all loci. The substitution matrix of the models 

is printed next to the trees. Another advantage of using MrAic in 

combination with Phyml was the obtained accuracy of tree topology 

and the greater calculation speed (Guindon & Gascuel 2003). 

Population genetic analyses 

In order to confirm the intraspecific diversity shown in the MP trees, 

the number of populations in the C. carrionii complex was inferred 

with Structure v. 2.2 (Pritchard et al. 2000) using genotype data 

of the ITS regions of rRNA gene and of the partial EF1 and BT2 

genes. Genotypes of these three loci of 43 isolates were sorted 

on the basis of sequence similarity. Structure is a model-based 

clustering method for using multilocus genotype data to infer 

population structure and assign individuals to populations. The 

parameters were as follows: the length of burn-in period was 

set to 10 6 , number of MCMC repeats after burn-in 30 000; the 

ancestry model: admixture (individuals have mixed ancestry and 

is recommended as starting point for most analyses). Uniform prior 

for ALPHA was set to 1.0 (default) and all allele frequencies were 

taken as independent among populations with λ set to 1.0 (default). 

Probability of the data (for estimating K) was also computed 

(Falush et al. 2003). The burn-in period length and number of 

MCMC repetitions after burn-in were set as 10 000 and 100 000, 

and admixture model and allele frequencies correlated model were 

chosen for analysis. The number of populations (K) was assumed 

from two to four. 

Association of multilocus genotypes was screened with the 

multilocus option in BioNumerics. To test for reproductive mode in 

each population, index of association (I A 

, a measure of multilocus 

linkage disequilibrium) was calculated with Multilocus v. 1.2.2 (www. 

bio.ic.ac.uk/evolve/software/multilocus). The null hypothesis for this 

analysis was complete panmixia. The values of I A 

were compared 

between observed and randomised data sets. The hypothesis would 

be rejected when p < 0.05. Population differentiation (index: theta, 

θ) was also detected using the same software and a null hypothesis 

for this analysis is no population differentiation. When observed θ is 

statistically significantly different from those of random datasets (p 

< 0.05), population differentiation should be considered. 

222


A reticulogram was reconstructed using T-rex (Makarenkov 

2001, Makarenkov & Legendre 2004) (www.labunix.uqam.ca/ 

~makarenv/trex.html) on C. carrionii / Cladophialophora sp. The 

program first computed a classical additive tree using one of the 

five available tree reconstruction algorithms. Subsequently, at each 

step of the procedure, a reticulation (a new edge) was chosen that 

minimised the least-squares or the weighted least-squares loss 

function; it was added to the growing reticulogram. Two statistical 

criteria (Q1 and Q2) were proposed to measure the gain in fit when 

reticulations were added. The minimum of each of these criteria 

may suggest a stopping rule for addition of reticulations. With 

HGT (horizontal gene transfer) reticulogram reconstruction option 

(Makarenkov 2001) the program mapped the gene tree into the 

species tree using the least-squares method. Horizontal transfers 

of the considered gene were then shown in the species tree. The 

reticulate network was created in the ITS tree, which served as 

a species tree and compared with a gene tree, EF1. Degrees of 

recombination or horizontal gene transfer were also visualised 

using SplitsTree v. 4.8 software (Huson & Bryant 2006). Split 

decomposition (Bandelt & Dress 1992) was applied on three loci of 

the entire C. carrionii complex. Calculation was done with default 

settings of characters transformation using uncorrected P-values, 

equal angles and optimise box iterations set to 1. Star- or brushlike 

trees indicate clonal development, while reticulation indicates 

genetic exchange. 

Isolation of fungi for inoculation experiments 

Nine plants of Stenocereus griseus, located within 50 m radius 

of the house of a patient with chromoblastomycosis due to strain 

UNEFM 9902 = CBS 114402 (C. carrionii) in Sabaneta (Miranda, 

Falcón State, Venezuela), were analysed. Four fragments of 

approx. 2 × 3 × 1 cm were excised from each plant at brownish 

superficial lesions in upper branches. Sampled fragments were 

soaked in mineral oil for 15 min at 23 °C under agitation at 150 

rpm (Fernández-Zeppenfeldt et al. 1994). Subsequently four 

cultivations were made per sample on agar slants. Strains with 

cultural characteristics and morphology similar to C. carrionii (de 

Hoog et al. 2000) were selected. Final identification was made by 

sequencing of the ITS region, by determining the ability of strains 

to grow at 35, 37, 38 and 40 °C, and whether they could break 

down 20 % gelatin (Richard-Yegres & Yegres 1987, Fernández- 

Zeppenfeldt et al. 1994). Environmental strain UNEFM-SgSR3 = 

CBS 114405 (Cladophialophora sp.) and clinical strain UNEFM 

9902 = CBS 114402 (C. carrionii) were selected for the inoculation 

experiments. 

Inoculum preparation 

Approximately 1 cm 2 of a culture on Sabouraud’s glucose agar 

(SGA) was transferred to 50 mL YPG medium (yeast extract 0.5 

%, peptone 0.5 %, glucose 2 %) (de Hoog et al. 2000), shaken at 

150 rpm and incubated for 3 d at 23 °C (Yegres et al. 1991). Five 

mL aliquots of the starter culture were transferred serially every 4 

d to 500 mL flasks containing 100 mL synthetic medium (d-glucose 

2 %, KH 2 

PO 4 

0.2 %, NH 4 

SO 4 

0.1 %, urea 0.03 %, MgSO 4 

0.03 

%, CaCl 2 

0.003 %; pH 6.2) shaken at 150 rpm at 23 °C. After 4 d 

the suspensions, which were predominantly conidia, were filtered 

through sterile gauze, ground in 50 mL 0.85 % saline, centrifuged 

at 2 000 rpm, and repeatedly washed with saline until a clear 

supernatant was obtained. The suspensions were adjusted to 5 × 

10 6 cells/mL (Yegres et al. 1991, Cermeño & Torres 1998). Inocula 

of 2 mL were checked for viability in lactritmel medium (de Hoog et 

al. 2000). 


Experimental cactus germlings 

Young cactus plants (Stenocereus griseus) were obtained in the 

laboratory (Clausnitzer 1978) by cultivation from seeds of a single 

cardon fruit collected near the house of the patient infected with CBS 

114402 in the endemic area for chromoblastomycosis in Falcon 

State, Venezuela. The seeds were rinsed with sterile distilled water, 

the contents washed by agitation for 10 min at 120 rpm in 250 mL 

sodium hypochlorite 4 % (v/v), and subsequently with sterile distilled 

water at 120 rpm for 5 min. The supernatant was decanted, 250 mL 

HCl 20 % was added, the seeds were incubated for 3 h, decanted 

and washed repeatedly with sterile distilled water. Seeds were 

then dried for 24 h on filter paper at 37 °C. Onset of germination 

was obtained by incubation of the seeds in a moist chamber on 

filter paper for 15 d under alternately 8 h of continuous white light 

(26 W) and 16 h of darkness; bud emergence was observed daily. 

Germlings of 1 cm in length, with green colour and having two 

leaves were transplanted to 128-container germinators until roots 

developed. The sterile substrate contained 5 parts Sogemix® and 

1 part river soil from the region where the fruit was collected. The 

daily light regime was as above; plants were watered every 10 d 

with 5 mL sterile tap water for 1 yr. 

Inoculation of S. griseus germlings 

Fungal suspensions (0.1 mL) were either injected using a syringe 

(13 × 0.4 mm) at a depth of approximately 5 mm into cortical 

tissue (Fig. 1A), or superficially applied onto (Fig. 1B) 96 randomly 

selected 1-yr-old plants: 50 % using clinical strain CBS 114402 

(C. carrionii) and 50 % using environmental strain CBS 114405 

(Cladophialophora sp.). The controls were 64 plants which were 

treated similarly, but using sterile saline (0.85 %). The growth 

chambers with inoculated plants stayed in the laboratory under the 

conditions specified above. From day 15 post inoculation onwards 

every 15 th day, six plants of each treatment were sectioned 

longitudinally from the apex and transversely by means of a handheld 

microtome, examined directly in glycerin water (25 %), and 

cultured in lactritmel medium (Fernández-Zeppenfeldt et al. 1994). 

Experimental cactus plants 

A total of 150 whole S. griseus plants ≤ 15 cm tall and without 

macroscopically visible lesions, were dug from an area within a 

50 m radius of the house of the patient with chromoblastomycosis 

as specified above. Plants were transported to the laboratory and 

transplanted individually into polyethylene bags with a capacity of 

1 kg, using as substrate river soil from the same area. Plants were 

maintained outside, directly adjacent to the laboratory to adjust 

at average temperatures of 32 °C and with natural daylight. They 

were watered with tap water every 15 th d for a period of 6 mo. 

Scar formation in mature S. griseus plants 

For inoculation purposes, 150 sharp, wooden toothpicks 4 × 0.3 × 

0.2 cm were washed and boiled for 3 min in tap water to eliminate 

resins (Yegres & Richard-Yegres 2002). This procedure was 

repeated three times. Three batches of 50 toothpicks each were 

kept separate in Petri dishes. Plates were incubated for 15 d at 

23 °C after inoculating each batch with 1 mL fungal suspension (5 

× 10 6 cells/mL) of either strain CBS 114405 or CBS 114402, with 

sterile water as control. A total of 50 randomly selected plants were 

inoculated (Fig. 1C) halfway up the shaft with a toothpick colonised 

with CBS 114402 (C. carrionii), CBS 114405 (Cladophialophora 

sp.) or the control (Yegres & Richard-Yegres 2002). 

Starting from 2 wk post inoculation, 10 plants were randomly 

chosen every 15 d, and tissue samples taken at the point of 

223


+ + + - - 

- 

+ 

+ 

A B C D 

Clinical C. carrionii 

> Environmental C. yegresii 

Fig. 1. Diagram of inoculation experiments with results. A. Inoculation of young cactus; B. Superficial application of young cactus; C. Traumatic application of mature cactus, 

with brown resulting scar; D. Superficial application of mature spines. Indications +/- refer to positive resp. negative results of re-isolated strains. Lower line: circles represent 

production of muriform cells, filaments represent hyphal growth. 

Fig. 2. Split decomposition of the C. carrionii complex using SplitsTree with uncorrected (P-value) distances. Nodes are shown only with different genotypes; hence EF1 shows 

the largest number of nodes. Extensive reticulation is noted in all loci. 

224


0.1 

99 

CBS102227 


92 

94 

85 



100 
















dH16363 























dH14614 

K=5 K=4 

Venezuela 

Venezuela 

Venezuela 

Venezuela 

Venezuela 

Venezuela 

Venezuela 

Venezuela 

unknown 

Venezuela 

Venezuela 

Venezuela 

Venezuela 

Venezuela 

Venezuela 

unknown 

Venezuela 

Venezuela 

Venezuela 

Venezuela 

Venezuela 

Venezuela 

Venezuela 

Venezuela 

Venezuela 

Venezuela 

Venezuela 

Australia 

Venezuela 

Venezuela 

Mozambique 

Uganda 

Australia 

Madagascar 

Australia 

Venezuela 

Venezuela 

Venezuela 

I 

II 

III 

Fig. 3. Phylogenetic tree (Neighbour-joining) of the C. carrionii complex based on EF1 (with grouping I–III) using the same strains of Fig. 1, generated using the HKY+G model. 

The model was calculated using ML in MrAic software. Bootstrap cut-off = 80 %. CBS 834.96 was taken as outgroup. Columns were generated with Structure software 

hypothesising K = 4 and K = 5, and alleles independent. Geographical origins in black refer to isolates from humans (chromoblastomycosis); origins in green refer to isolates 

from plant material. 

inoculation, and from the thorns directly adjacent to this area. 

Samples were rinsed with 4 % sodium hypochlorite for 3 min, and 

subsequently washed in sterile distilled water for re-isolations, and 

for histological study by means of light microscopy (Fernández- 

Zeppenfeldt et al. 1994). 

Experiments with spines of S. griseus 

Ninety cactus spines of 2.5 cm av. length were collected from 

a single S. griseus plant located near the home of the patient 

infected with CBS 114402, at approx. 2.5 m height, superficially 

sterilised, and divided into three groups, of which 30 spines were 

inoculated with CBS 114402 (C. carrionii), 30 with CBS 114405 

(Cladophialophora sp.) and 30 to be used as control, inoculated 

with a saline solution (Fig. 1D). A similar series composed of 90 

spines of 1.5 cm average length was collected at approx. 1 m 

height. All spines were incubated in sterile Petri dishes with filter 

paper (Whatman #1) with 2 mL saline solution; subsequently 0.1 

mL fungal suspension was applied. Twenty spines were analysed 

weekly by means of longitudinal sectioning with a hand-held 

microtome, cultured as above and observed microscopically until 

day 75 post incubation. 


225


ROOT 















5 







dH16363 














1 7 

4 6 

3 2 9 

10 

8 


A 

B 




A 

Unknown 

Australia 

Australia 

Australia 

Madagascar 

Mozambique 

D 

Uganda 

C. carrionii 

C. yegresii 

Fig. 4. Reticulogram of South American strains of Cladophialophora species and strains from other continents (mentioned) constructed with T-rex software. ITS rDNA (with 

grouping A, B, D) was used as species tree and compared with the ß-tubulin gene tree. First 10 reticulations are shown with numbers. 

Statistics 

Survival of the cactus seedlings and collected plants following 

inoculation were evaluated using the X 2 -test (P = 0.05 was 

considered significant). Stem lesions resulting from inoculations 

were analysed with Student’s T-test (P = 0.01 was considered 

significant). 

RESULTS 

The rDNA ITS region was sequenced for 43 strains identified as 

C. carrionii based on morphology. Sequences of 16 additional 

strains were downloaded from GenBank. Five distantly related, 

unidentified cladophialophora-like species were added, with CBS 

834.96 as outgroup. In C. carrionii, 203 positions were compared 

in ITS1, 158 in the 5.8S rRNA gene and 182 in ITS2 (Table 3). 

The sequences could be aligned with confidence over their entire 

lengths. Over the data set, 35 positions were polymorphic, of which 

33 were phylogenetically informative (Table 3), the two remaining 

being variable T-repeats near the ends of ITS1 and 2. 

For ITS sequences the AICc selected the TrN+G model (TrNG; 

Tamura & Nei 1993). The base frequency of ITS: T = 0.2467, C 

= 0.2897, A = 0.2247, G = 0.2390, TC = 0.5364, AG = 0.4636. 

The EF1 tree was built with substitution model HKY+G; the base 

frequency of EF1: T = 0.2990, C = 0.2665, A = 0.2123, G = 0.2221, 

TC = 0.5655, AG = 0.4345. The best model for BT2 sequences 

was the SYM+I+G (symmetrical model). The base frequency for 

BT2: T = 0.2255, C = 0.2953, A = 0.2463, G = 0.2328, TC = 0.5208, 

AG = 0.4792. Bootstrap values of the EF1 tree were calculated 

with PAUP using parsimony and with maxtrees set to 500 and 500 

replicates (data not shown). Total number of characters was 191 of 

which 101 were parsimony-informative. Tree length was 365 and 

had the following indices: Consistency Index = 0.685, Retention 

Index = 0.542 and Homoplasy Index = 0.315. 

The original tree length, L o 

was 1 055, the tree length of the 

combined data, L c 

was 1 062. The resulting incongruence length 

difference L = (L c 

–L o 

) was 7 (P = 0.24). The observed ILD was not 

significantly greater than expected by chance and it was concluded 

that the sequences were congruent and could be used together in 

a combined analyses. 

Split decomposition based on the same alignment generated 

extensive recombination. The structure found with three loci was 

robust, with the exception of separation of CBS 834.96 and CBS 

102227 with EF1 (Fig. 2). 

The core of the network, comprising the strains listed in Table 1, 

was analysed in more detail. With ITS, four groups were recognised 

(A–D; Table 1). (A) was the main group with 36 strains / sequences; 

FMC 248 differed only by a small T-repeat and was regarded as a 

member of (A). The remaining groups were smaller, differing from 

group (A) maximally by two consistent positions (Table 3). Group 

(C) mainly comprised sequences from GenBank and all originated 

from Abliz et al. (2004). One of the strains of group (C), IFM 4808, 

concerned a subculture of CBS 160.54, which is an original isolate 

of Trejos (1954) representing C. carrionii. Re-sequencing indicated 

that it was a member of group (B). Analysis of our electropherograms 

of this isolate was not suggestive of heterothallism. None of the 

positions characterising groups (A)–(C) were also found to differ in 

group (D), which deviated in 16 mutations in ITS1 and 8 in ITS2; 

17 of the mutations were transitions, 7 were transversions and 7 

indels. Group (D) was clearly distinct from the complex of (A)–(C), 

226


Table 3. Nucleotide variability of ITS1-2 ribosomal DNA regions of 

Cladophialophora carrionii (A – C) and C. yegresii (D). 

rDNA domains (length), with variable nucleotide positions. 

ITS1 (201-203) A B C D 

16 C C C T 

17 T C T T 

19 T T T C 

51 A A A G 

57 A A A T 

90-92 TG- TG- TG- CGT 

101 T T T C 

103 C C C T 

104 G A G G 

106 A A A G 

114 T T T C 

122 T T T C 

132 C C C T 

137 A A A C 

141 C C C T 

145 - - - A 

163-170 6-10T 6-10T 6-10T TTGTATCT 

180 - - - A 

183 G G G A 

190 T/A A A A 

5.8S (158) Monomorphic 

ITS2 (178-182) A B C D 

36 C T T T 

48 T T G T 

49 T T T C 

51 - - - C 

114 C C C G 

140 A A A G 

155 - - - T 

178-179 -- -- -- CT 

with a total of 27 mutations. 

For multilocus analysis with ITS, EF1 and BT2 a smaller set 

of strains was compared. Sequences of the 205 bp long element 

of EF1 contained 32 phylogenetically informative mutations. Three 

entities were distinguished (I–III; Fig. 3). With BT2, three groups 

with the same composition were recognised. Strains of ITS group 

(C) were not available for study. 

On the basis of multilocus screening in BioNumerics, concordant 

groups (A)–(D) were tested with the Structure programme. When 

K was set at 4 or 5, consistent groupings were noted, indicated as 

I, II and III (Fig. 3), corresponding with ITS groups (A), (B) and (D), 

respectively in Table 1. 

The possibility that group (D) / (III) included a member of another, 

morphologically similar but phylogenetically unrelated group of fungi 

was excluded by SSU sequencing. Genera morphologically similar 

to Cladophialophora, such as Cladosporium Link, Devriesia Seifert 


& N.L. Nick., Phaeoramularia Munt.-Cvetk., Pseudocladosporium 

U. Braun and Stenella Syd. proved to be remote (data not shown). 

With T-rex, interaction between groups (B) and (D) was noted, 

rather than between groups (B) and (A), despite the high sequence 

similarity of (A) and (B) (Fig. 4). 

Morphological observation revealed that representatives of 

ITS groups (A)–(C) generally had conidiophores that arise at right 

angles from creeping hyphae (Fig. 5), while those of (D) tend to 

be ascending, hyphae gradually becoming conidiophore-like. Since 

slight correspondence was found in independent markers and 

phenetic criteria, we considered group (D) to represent a separate 

species, which is described as follows. 

Cladophialophora yegresii de Hoog, sp. nov. MycoBank 

MB500208. Figs 6, 7D–F. 

Etymology: Named after Francisco Yegres, Venezuelean 

mycologist. 

Coloniae in agaro PDA dicto 22 °C planae, olivaceo-virides, pulverulentae vel 

velutinae, margine integra; reversum olivaceo-atrum. Hyphae fertiles dilute olivaceovirides, 

ascendentes, paulatim in catenas conidiorum concolorium vertentes. 

Conidiorum catenae ramosae, conidia dilute olivaceo-viridia, levia et tenuitunicata, 

4.5–6 × 2.5 µm, faciliter liberata, cicatricibus modice pigmentatis. Chlamydosporae 

et cellulae zymosae absentes. Synanamorphe phialidica non visa. Teleomorphe 

ignota. 

Holotypus cultura sicca CBS H-18464 in herbarium CBS praeservatur. 

Colonies on PDA at 22 °C evenly olivaceous green, powdery to 

velvety, with entire margin; reverse olivaceous black. Fertile hyphae 

pale olivaceous green, ascending, gradually changing over into 

concolorous chains of conidia. Conidial system profusely branched. 

Conidia pale olivaceous green, smooth- and thin-walled, 4.5–6 

× 2.5 µm, detached rather easily, with slightly pigmented scars. 

Chlamydospores and yeast cells absent. Phialidic synanamorph 

not observed. Teleomorph unknown. 

Specimen examined: Venezuela, Falcon state, from asymptomatic Stenocereus 

griseus cactus, G. Fernández-Zeppenfeldt, CBS H-18464 holotype, culture ex-type 

CBS 114405 = UNEFM SgSR3. 

Notes: Of the 48 dematiaceous isolates obtained from 36 fragments 

of the cactus Stenocereus griseus, four strains originating from 

four different plants of S. griseus presented morphological and 

physiological key characteristics of Cladophialophora carrionii or 

C. yegresii (de Hoog et al. 2000, 2006). Gelatin liquefaction was 

negative in all strains and the maximum growth temperature was 

37 °C. After identification to species level using sequence data (de 

Hoog et al. 2006), both C. carrionii and C. yegresii appeared to be 

among the strains isolated. 

A total of 256 plants obtained at the end of 1 yr from germlings, 

had ribs, spines, and an average height of 15 cm. The 96 

germlings inoculated with fungal suspensions of the test strains 

CBS 114402 (C. carrionii, clinical) and CBS 114405 (C. yegresii, 

environmental) remained without visible external lesions during the 

year of experimentation. Histological sections of the 96 inoculated 

plants consistently revealed internal growth of the fungi in their 

filamentous form. Muriform cells were not observed, neither on 

the epidermis, nor in the internal tissue, spines or roots. The reisolated 

cultures demonstrated the viability of the fungi during the 

entire experimental process: CBS 114402 (C. carrionii) was grown 

from 26 (54.16 %) of the plants and CBS 114405 (C. yegresii) in 

23 (47.90 %) of the plants. The X 2 test did not reveal significant 

differences between the isolates (X 2 c = 0.0729 < X 2 t = 3.84). The 

32 control plants remained without external lesions, and in the 

227


Fig. 5. Microscopic morphology of C. carrionii, strain CBS 160.54. Conidiophore erect, i.e. mostly arising at 90° from creeping hypha (sketch). Scale bar = 10 µm. 

histological sections no internal or external fungal elements were 

observed. None of the fungi isolated from the control plants proved 

to be a species of Cladophialophora. 

With 96 plants with superficial application of spore suspension 

(48 plants for each isolate, either clinical or environmental) neither 

internal nor external lesions were observed. Histological sectioning 

did not reveal fungal elements in or on plant tissue. Short hyphal 

elements and meristematic cells were occasionally seen around 

and inside the outer layers of the spines. The re-isolated strains 

proved that the fungi survived during the entire experimental 

procedure: CBS 114402 (C. carrionii) was isolated from 32 (66.67 

%) plants and CBS 114405 (C. yegresii) from 33 (68.75 %). The X 2 

test did not detect significant differences in survival rates among 

the isolates (X 2 c = 0.4375 < X 2 t = 3.84). 

Mature plants inoculated using colonised toothpicks showed 

average scarring of 1.88 cm diam with C. carrionii and 1.33 cm 

diam with C. yegresii, around the point of inoculation. In histological 

sections of 100 plants, dark, septate hyphae with inflated elements 

were observed at the points of inoculation. Muriform cells were not 

observed. Re-isolated strains were evidence of isolate viability: 

228


Fig. 6. Microscopic morphology of C. yegresii, strain CBS 114405. Conidiophore ascending, i.e. mostly emerging from hyphal end that is gradually growing upwards to become 

a conidiophore (sketch). Scale bar = 10 µm. 

CBS 114402 (C. carrionii) was grown from 36 (72 %) plants and CBS 

114405 (C. yegresii) from 30 (60 %). The fungi could not be isolated 

from spines. The 50 plants used as controls showed scarring of 

1.06 cm diam on average around the point of inoculation. No fungal 

elements were seen in direct examinations and histological sections 

of these plants. The retro-cultures were negative. The scarring 

responses of the plants to the clinical strain, environmental strain 

and control proved to be highly significant: 

Clinical CBS 114402 vs. environmental CBS 114405: þ = 0.000832, 

P = 0.01; 

Clinical CBS 114402 vs. control: þ = 0.00003128, P = 0.01; 

Environmental CBS 114405 vs. control: þ = 0.005343, P = 0.01. 

Spines 2.5 cm av. in length, seeded with suspensions of CBS 

114402 (C. carrionii) and CBS 114405 (C. yegresii), developed 

toruloid hyphal elements with some dark, swollen cells similar 

to muriform cells known in human tissue. The re-isolated strains 

proved the species to survive during the experimental procedure 

(< 75 d). Similar results were obtained with the spines 1.5 cm av. 

in length. No cladophialophora-like fungi were isolated from the 

controls. 


229


Fig. 7. Conidial morphology in selected branches of (upper row: A–C) C. carrionii, strain CBS 260.83; (lower row: D–F) C. yegresii, strain CBS 114405. In this respect the two 

species are identical. Scale bar = 10 µm. 

DISCUSSION 

Taxonomy of Cladophialophora 

Judging from SSU rDNA phylogeny data, all Cladophialophora 

species that are consistently associated with pathology to humans 

belong to the Herpotrichiellaceae in the order Chaetothyriales 

(Haase et al. 1999). Within this order, the genus Cladophialophora 

is polyphyletic. Conidia of all species are produced in branched 

chains on poorly differentiated hyphae. This very simply structured 

conidial system may lead to confusion with morphologically 

similar but unrelated fungi that are encountered as contaminants 

in the hospital environment. The genus Cladosporium comprises 

ubiquitous airborne fungi which mostly have erect, more or less 

differentiated conidiophores, and dark conidial scars. They are 

associated with Davidiella Crous & U. Braun teleomorphs and 

belong to the Dothideomycetes, family Davidiellaceae (Braun et al. 

2003, Schoch et al. 2006). Pseudocladosporium was introduced by 

Braun (1998) with three species differing from Cladophialophora 

mainly by intercalary hyphal cells with lateral extensions that bear 

conidial chains, having Caproventuria U. Braun teleomorphs (see 

Crous et al. 2007 – this volume). The group is classified in the 

Venturiaceae and Mycosphaerellaceae in the Dothideales (Braun et 

al. 2003). The anamorph genus Devriesia comprises thermophilic 

saprobes with a cladophialophora-like appearance and producing 

230


dark, multi-celled chlamydospores alongside the hyphae. 

Phylogenetically this genus is related to the Mycosphaerellaceae, 

in the Dothideomycetes (Seifert et al. 2004). 

Cladophialophora carrionii was originally introduced by Trejos 

(1954) on the basis of 46 strains from Venezuela, Australia and 

South Africa. He did not indicate a holotype. For this reason isolate 

Trejos 27 = Emmons 8619 = CBS 160.54, the first strain mentioned 

by Trejos (1954), is selected here as representative for C. carrionii. 

A dried specimen of this strain has been deposited as lectotype 

in the Herbarium of the Centraalbureau voor Schimmelcultures as 

CBS H-18465. 

The ex-type strain of Cladophialophora ajelloi Borelli, CBS 

260.83, proved to be indistinguishable from C. carrionii, which was 

also known to be able to produce phialides in addition to catenate 

conidia (Honbo et al. 1984). Remarkably, a strain identified as C. 

ajelloi from Samoa (CBS 259.83; Goh et al. 1982) proved to be 

related to but consistently different from all strains of the C. carrionii 

complex. The 43-year-old male patient in otherwise good health 

carrying this fungus had a 5 × 3 cm erythematous, scaling lesion 

on his arm. Muriform cells were present in superficial dermis and 

stratum corneum. This clearly represents yet another agent of 

human chromoblastomycosis. The name C. ajelloi is not available 

for this taxon, as this is a synonym of C. carrionii. The taxon will be 

formally described in a forthcoming paper. 

Members of ITS groups (A)–(D) were shown to be close to 

each other in SSU phylogeny (data not shown) underlining that all 

analysed species were correctly assigned to Cladophialophora. 

This genus was defined by melanised acropetal chains of conidia, 

near absence of conidiophores, and phylogenetic affinity to the 

order Chaetothyriales. Strains (A)–(D) clustered in a clade which 

contained a mixture of species of Cladophialophora, Fonsecaea 

Negroni and Phialophora Medlar. From a point of view of human 

disease, the species of the clade were known as agents of brain 

infection [C. bantiana (Sacc.) de Hoog et al., F. monophora (M. Moore 

& F.P. Almeida) de Hoog et al.], disseminated disease [C. devriesii 

(A.A. Padhye & Ajello) de Hoog et al.], cutaneous disease [C. boppii 

(Borelli) de Hoog et al.] and particularly chromoblastomycosis (C. 

carrionii, Fonsecaea, Phialophora). 

Diversity of Cladophialophora carrionii / C. yegresii 

Infraspecific variability was observed within C. carrionii. The groups 

(A)–(C) were separated on the basis of five mutations in the ITS 

region, which were supported by mutations in EF1 and BT2, as 

confirmed by analysis in Structure, where the same separation 

(K = 5) of entities was observed. Furthermore, K = 4 unites groups 

(B/II) and (D/III), despite the fact that the sequence of (B) is more 

close to those of (A). With T-rex software a similar relationship 

between [(B), C. carrionii] and [(D), C. yegresii] was noted, 

suggesting horizontal gene flow between these entities. This is 

remarkable, since (C) strains predominantly inhabit remote deserts 

in Madagascar and Australia, while (D) is found in equally remote 

localities in Venezuela. Extensive reticulation was observed in all 

genes with SplitsTree. With ITS and BT2, CBS 834.96 and CBS 

102227 cluster closely together, while in the more variable EF1 

data these are all widely apart, suggesting that in Cladophialophora 

other mechanisms than recombination may occur. 

Group (C) contained ITS sequences taken from the public 

domain, originating from a single study (Abliz et al. 2004). 

Remarkably, strain IFM 4808 found in group (C) on the basis of data 

from Abliz et al. (2004), was the same isolate as CBS 160.54, which 

was found repeatedly in group (B) in our data set (Table 1). A similar 


phenomenon was observed with strain IFM 41444 = CBS 863.96, 

of which GenBank deposition AB109169 consistently deviated from 

our data in a frequently observed mutation. A possible explanation 

of these consistent sequence conflicts is heterozygosity. Although 

most chaetothyrialean fungi are supposed to be haploid (Szaniszlo 

2002; Zeng et al. 2007), some strains have a double DNA content 

in yeast cells (Ohkusu et al. 1999). Teleomorphs are not known 

in Cladophialophora and related black yeasts, but many species 

are known to form profuse hyphal anastomoses (de Hoog et al. 

2006), allowing parasexual processes and mitotic recombination. 

However, all electropherograms including those from the study of 

Abliz et al. (2004), which were kindly sent by K. Fukushima (Chiba, 

Japan), were unambiguous, without double peaks. This matched 

with the observation of preponderant clonality despite frequent 

anastomoses in Exophiala J.W. Carmich. (Zeng et al. 2007). An 

alternative explanation might be the occurrence of paralogous ITS 

repeats, as reported earlier in Fusarium Link (O’Donnell & Cigelnik 

1997). 

The remaining diversity within C. carrionii as confirmed by 

Structure shows some geographical structuring of populations, 

in that group (A) does not occur in Asia, group (B) is limited to 

Australia and Africa, and group (C) has thus far only been reported 

from Asia. The wide distribution of most genotypes suggests, 

however, that worldwide occurrence is likely to become apparent 

when more strains have been analysed. All climate zones where 

C. carrionii was isolated were semi-arid to arid, desert-like. 

Genotypes were not limited to the endemic semi-arid areas, and 

thus a relatively rapid vector of dispersal has to be hypothesised 

enabling the fungus to cross climate zones where the saprobic 

phase is unable to survive. Kawasaki et al. (1993) analysed three 

further loci in mtDNA using RFLP. Only some of their strains were 

available for sequencing. These had all identical mtDNA profiles, 

with the exception of IFM 4808 = CBS 160.54, that differed in two 

markers (Table 1). If we assume that there is no real separation of 

ITS groups (A) and (C) (see above), the conclusion is warranted 

that mtDNA allows distinction of polymorphism at the same level of 

diversity as detected in this study with ITS, EF1 and BT2. 

South America harbours group (D) which represents a second 

species, C. yegresii. This species thus far has not been found 

on humans, and seems to be restricted to living Stenocereus 

cactus plants. Nishimura et al. (1989) published a strain from 

chromoblastomycosis in China which matched the morphology of 

strains now classified as C. yegresii, but as far as we are aware this 

strain has not been sequenced. 

Ecology and virulence of Cladophialophora carrionii / C. 

yegresii 

Cladophialophora carrionii was preponderantly found as an agent 

of human infection and only occasionally on dead plant debris, 

mainly seceded cactus needles. The only three strains available 

of C. yegresii were isolated from living, asymptomatic Stenocereus 

plants surrounding the cabin of a symptomatic patient from whom C. 

carrionii, CBS 114402 was isolated. Although in some publications 

convincing evidence was presented that infections originate from 

puncture by plant material (e.g., Salgado et al. 2004), it now 

becomes clear that the environmental look-alikes of clinical strains 

do not necessarily belong to the same species (Crous et al. 2006, 

Mostert et al. 2006), but may be members of other, related taxa with 

slightly different ecology; an unambiguous connection between a 

clinical and an environmental strain still has to be proven. 

The endemic area of the two species, C. carrionii and C. 

231


yegresii, has a semi-arid climate, with average yearly temperatures 

of 24 ºC, scarce rainfall (up to 600 annual mL) and is located at 

moderate altitude (up to 500 m) (Borelli 1979, Richard-Yegres & 

Yegres 1987). The landscape is dominated by large cacti and other 

xerophytes. Stenocereus griseus is a columnar American cactus 

with a very strong, protective external epidermis that allows the 

accumulation of water in the shaft and enables tolerance of extreme 

drought. The species produces ovoidal, thorny fruits of about 5 cm 

diam, which are commonly eaten by the local population. It has 

therefore been suggested that patients with chromoblastomycosis 

acquire their infection by traumatic inoculation with cactus spines, 

similar to the supposed infection process of Madurella mycetomatis 

(Laveran) Brumpt in the arid climate of Africa (Ahmed et al. 2002). 

The frequent occurrence of 16 / 1 000 for chromoblastomycosis in 

areas endemic for Cladophialophora in Venezuela (Yegres et al. 

1985; Yegűez-Rodriguez et al. 1992) indicates a marked invasive 

potential for C. carrionii. Local goat-keepers are particularly at risk: 

in 1984, 14 of 18 patients investigated had these occupational 

characteristics (Yegres et al. 1985). Nevertheless, virulence of C. 

carrionii is low when inoculated into the footpads of mice (Yegres 

et al. 1998); also an environmental strain of C. carrionii failed to 

produce lesions in mice and in a volunteer (Richard-Yegres & 

Yegres 1987). 

We performed inoculation experiments with C. carrionii and 

C. yegresii using freshly grown, healthy cacti in the greenhouse. 

The plants were followed over a 1-yr period; during all this time 

the control plants remained without lesions. Both Cladophialophora 

strains were able to produce infection when syringe-inoculated 

deep into young cactus tissue. Histopathology showed septate 

hyphae between host cells, and the shaft was maintained over 

prolonged periods without causing visible damage. This absence 

of appreciable destruction would categorise them as endophytes. 

Cactus tissue is rich in carbohydrates, vitamins and minerals (Vélez 

& Chávez 1980) which may promote endophyte growth. 

In contrast, suspensions applied superficially lead to growth 

on and in spines only. The absence of infection after superficial 

application indicates that the fungi are unable to invade healthy 

plant tissue from the surface and thus they cannot be characterised 

as obligatory phytopathogens. 

The two species differed in the degree of scarring after traumatic 

inoculation into mature plants: the clinical strain C. carrionii was 

consistently more virulent than C. yegresii that originated from 

the same host plant. Both species showed the same viability in 

re-isolated cultures. In nature, the fungi are likely to invade only 

when the integrity of the epidermis is broken, as happens e.g. 

by goat feeding or transmission by sap-sucking birds or piercing 

insects. They also show the same transformation to meristematic 

morphology (González et al. 1990) when entering hard spine 

tissue. A possible trigger for this conversion is the dominance of 

lignin in the spines. Survival on and in spines is enhanced by their 

capturing of atmospheric water formed after nightly condensation. 

The fact that superficial application leads to colonisation around and 

inside the spines suggests that the spines play a role in mechanic 

dispersion of the fungi. 

A possibly coincidental mechanism of dispersal might be 

traumatic inoculation into living tissue of humans or animals, where 

the same muriform cells are formed, defining the skin disease 

chromoblastomycosis. It may be questioned whether animal/ 

human inoculation plays a role in the evolution of the fungus. ITS 

differences between the two species are observed in 23 positions, 

with a ratio of transitions : transversions of 2 : 1 (Table 3). Thus 

no saturation of mutations has taken place and the diversification 

can be regarded as an example of recent sympatric speciation. 

Cladophialophora carrionii is widely distributed, and shows a higher 

degree of diversity than C. yegresii. This would be suggestive for 

a longer evolutionary time span of existence and C. carrionii then 

should be regarded as ancestral to C. yegresii, with the latter 

showing a founder effect due to the absence of polymorphisms. 

However, such an order of event (a host jump from humans to 

cactus) is difficult to imagine. It is more likely that C. yegresii is 

the original cactus endophyte exhibiting extremotolerance via its 

muriform cells. T-rex data suggested a more direct connection of 

C. yegresii with African and Australian rather than Venezuelean 

strains of C. carrionii. We suppose that the low degree of observed 

variation in C. yegresii is not a founder effect, but rather a sampling 

effect, as living cacti have thus far not been studied outside the 

framework of our study on the patient with Cladophialophora 

chromoblastomycosis. The difference in virulence may be simply 

explained by C. carrionii, which lives as a saprobe on dead cactus 

debris for part of its life cycle, and is less adapted to an endophytic 

life style. 

Cladophialophora cf. carrionii is known to occur on lignified 

materials, such as wood chips of Eucalyptus crebra and wooden 

remains of Prosopis juliflora and Stenocereus griseus (Riddley 

1957, Yegres et al. 1985, Fernández-Zeppenfeldt et al. 1994). This 

does not exclude a certain degree of pathogenicity to humans, as 

also pathogens like Cryptococcus neoformans (Sanfelice) Vuill. 

are known to have an essential part of their life cycle in hollows 

of Eucalyptus trees. Cryptococcus neoformans produces diphenol 

oxidase to degrade lignin, an aromatic polymer in the cell wall of 

plants and a component of wood (Cabral 1999). Similar degradation 

pathways are present in Cladophialophora carrionii (Prenafeta- 

Boldú et al. 2006). 

The natural occurrence of C. carrionii and C. yegresii in 

association with xerophytes has been proven, but their environmental 

route of dispersal is still unknown. As transformation to meristematic 

cells takes place when the hyphae reach the spines and on dead 

spines, the muriform cell apparently is the extremotolerant phase 

of the species. The conidial anamorph can be found sporulating on 

rotten spines directly after rainfall (Richard-Yegres & Yegres 1987), 

but as the fungus has thus far never been isolated from outside air, 

it is still unclear how a new host plant is reached. 

The behaviour of C. carrionii on humans, provoking the very 

characteristic disease, chromoblastomycosis, of which the agents 

are limited to the ascomycete family Herpotrichiellaceae (de Hoog 

et al. 2000) is puzzling. In humans, the extremophilic muriform 

anamorph is expressed rather than hyphae, and thus humans do 

not seem a natural reservoir of the fungus. Nevertheless some 

acquired cellular immunity seems to be involved. Albornoz et al. 

(1982) demonstrated that a significant share of the local population 

of goat keepers (Yegres et al. 1985) is asymptomatically infected 

with C. carrionii; Iwatsu et al. (1982) detected cutaneous delayed 

hypersensitivity in rats experimentally-infected with agents of 

chromoblastomycosis. With murine experimental infection of 

the related fungus Fonsecaea pedrosoi, Ahrens et al. (1989) 

found enlargement and metastasis of lesions in athymic but not 

in normal mice, or in mice with defective macrophage function. 

Several authors (Kurita 1979, Nishimura & Miyaji 1981, Polak 

1984) observed a significant role of acquired cellular immunity in F. 

pedrosoi, while Cardona-Castro & Agudelo-Flórez (1999) obtained 

chronic infection in immunocompetent mice when inoculated 

intraperitoneally. Garcia Pires et al. (2002) noted an unbalance 

between protective Th1 and less efficient Th2 responses. The 

possible host response leads to different clinical types, referred 

232


to as tuberculoid and suppurative granuloma, respectively. 

The existence of genetic constitutional factors in susceptibility is 

underlined by a marked frequency of family relationships among 

symptomatic individuals (Yegűez-Rodriguez et al. 1992). The 

disease is not observed in local animals such as goats, possibly 

due to their high body temperature (≈ 39 °C). Nevertheless, hyphal 

fragments artificially inoculated into goats led to transformation into 

muriform cells, but the lesions disappeared within 60 d (Martínez 

et al. 2005). Further animal experiments using strains identified 

according to new taxonomy will be necessary to answer questions 

on the role of the fungus on warm-blooded animals. 


The authors are indebted to F. Yegres, N. Richard-Yegres and V. Maigualida Pérez- 

Blanco for providing a large set of strains for study and information on strain ecology, 

and to R.C. Summerbell for helpful suggestions on the manuscript. P. Abliz and K. 

Fukushima are acknowledged for making sequence data available for study. K.F. 

Luijsterburg is thanked for technical assistance. The study was supported in part 

by the Fondo Nacional de Investigaciones Cientificas y Tecnológicas (Fonacit), 

Venezuela. 

REFERENCES 

Abliz P, Fukushima K, Takizawa K, Nishimura K (2004). Specific oligonucleotide 

primers for identification of Cladophialophora carrionii, a causative agent of 

chromoblastomycosis. Journal of Clinical Microbiology 42: 404–407. 

Ahmed AOA, Adelmann D, Fahal A, Verbrugh H, Belkum A van, Hoog GS de (2002). 

Environmental occurrence of Madurella mycetomatis, the major agent of human 

eumycetoma in Sudan. Journal of Clinical Microbiology 40: 1031–1036. 

Ahrens J, Graybill JR, Abishawl A, Tio FO, Rinaldi MG (1989). Experimental murine 

chromomycosis mimicking chronic progressive human disease. American 

Journal of Tropical Medicine and Hygiene 40: 651–658. 

Albornoz MB, Marin C de, Iwatsu T (1982). Estudio epidemiologico de un area 

endemica para cromomicosis en el Estado Falcon. Investigaciόn Clínica 23: 

219–228. 

Bandelt HJ, Dress AW (1992). Split decomposition: a new and useful approach to 

phylogenetic analysis of distance data. Molecular Phylogenetics and Evolution 

1: 242–252. 

Borelli D (1972). Significado del dimorfismo de ciertos hongos parásitos. 

Mycopathologia et Mycologia Applicata 46: 237–239. 

Borelli D (1979). Reservarea de algunos agentes de micosis. Medicina Cutánea 

4: 367–370. 


(phytopathogenic hyphomycetes), Vol. 2. IHW-Verlag, Eching. 


taxonomy of Cladosporium-like hyphomycetes, including Davidiella gen. nov., 


Burnham KP, Anderson DR (2002). Model selection and multimodel inference, a 

practical information-theoretic approach. 2nd ed, Springer, New York. 

Cabral L (1999). Wood, animals and human beings as reservoirs for human 

Cryptococcus neoformans infection. Revista Iberoamericana de Micologia 16: 

77–81. 

Carbone I, Kohn LM (1999). A method for designing primer sets for speciation 

studies in filamentous ascomycetes. Mycologia 91: 553–556. 

Cardona-Castro N, Agudelo-Flórez P (1999). Development of a chronic 

chromoblastomycosis model in immunocompetent mice. Medical Mycology 37: 

81–83. 

Cermeño J, Torres J (1998). Método espectrofotométrico en la preparación del 

inóculo de hongos dematiaceos. Revista Iberoamericana de Micologia 15: 

155–157. 

Clausnitzer I (1978). Germinación de las semillas del dato o frutos del cardón, 

Lemaireocereus griseus (Haw) Britt & Rose. Revista de la Facultad de 

Agronomia de la Universidad del Zulia 5: 351–365. 





Crous PW, Slippers B, Wingfield MJ, Rheeder J, Marasas WFO, Philips AJL, Alves 

A, Burgess T, Barber P, Groenewald JZ (2006). Phylogenetic lineages in the 


Falush D, Stephens M, Pritchard JK (2003). Inference of population structure: 

Extensions to linked loci and correlated allele frequencies. Genetics 164: 

1567–1587. 

Felsenstein J (1993). PHYLIP (phylogeny inference package), version 3.6a2. 

Department of Genetics, Univ. Washington, Seattle. 

Fernández-Zeppenfeldt G, Richard-Yegres N, Yegres F, Hernández R (1994). 

Cladosporium carrionii: hongo dimórfico en cactáceas de la zona endémica 

para la cromomicosis en Venezuela. Revista Iberoamericana de Micologia 11: 

61–63. 

Garcia Pires d’Ávila SC, Pagliari C, Seixas Duarte MI (2002). The cell-mediated 

immune reaction in the cutaneous lesion of chromoblastomycosis and their 

correlation with different clinical forms of the disease. Mycopathologia 156: 

51–60. 

Glass NL, Donaldson G (1995). Development of primer sets designed for use with 

PCR to amplify conserved genes from filamentous ascomycetes. Applied and 

Environmental Microbiology 61: 1323–1330. 

Goh KS, Padhye AA, Ajello L (1982). A Samoan case of chromoblastomycosis 

caused by Cladophialophora ajelloi. Sabouraudia 20: 1–5. 

González R, Caleiras E, Guanipa O, Garcia-Tamayo J, Siva L, Colina C, Fernández- 

Zeppenfeldt G, Pacheco I (1990). Chromomycosis: ultrastructural characteristics 

of the suppurative granuloma in Cladosporium carrionii skin lesions. Jornada 

Microscopica Electrónica 4: 133–134. 

Guindon S, Gascuel O (2003). A simple, fast, and accurate algorithm to estimate 

phyogenies by maximum likelihood. Systematic Biology 52: 696–704. 

Haase G, Sonntag L, Melzer-Krick B, Hoog GS de (1999). Phylogenetic interference 



Honbo S, Padhye AA, Ajello L (1984). The relationship of Cladosporium carrionii to 

Cladophialophora ajelloi. Sabouraudia 22: 209–218. 

Hoog GS de, Guarro J, Gené J, Figueras MJ (2000). Atlas of Clinical Fungi, 2 nd 

ed. Centraalbureau voor Schimmelcultures, Utrecht and Universitat Rovira i 

Virgili, Reus. 

Hoog GS de, Vicente VA, Caligiorne RB, Kantarcioglu AS, Tintelnot K, Gerrits 

van den Ende AHG, Haase G (2003). Species diversity and polymorphism in 

the Exophiala spinifera clade containing opportunistic black yeast-like fungi. 

Journal of Clinical Microbiology 41: 4767–4778. 

Hoog GS de, Zeng JS, Harrak MJ, Sutton DA (2006). Exophiala xenobiotica sp. nov., 

an opportunistic black yeast inhabiting environments rich in hydrocarbons. 

Antonie van Leeuwenhoek 90: 257–268. 

Huson DH, Bryant D (2006). Application of phylogenetic networks in evolutionary 

studies. Molecular Biology and Evolution 23: 254–267. 

Iwatsu T, Miyaji M, Taguchi H, Okamoto S (1982). Evaluation of skin test for 

chromoblastomycosis using antigens prepared from culture filtrates of 

Fonsecaea pedrosoi, Phialophora verrucosa, Wangiella dermatitidis and 

Exophiala jeanselmei. Mycopathologia 77: 59–64. 

Kawasaki M, Ishizaki H, Miyaji M, Nishimura K, Matsumoto T, Honbo S, Muir D 

(1993). Molecular epidemiology of Cladosporium carrionii. Mycopathologia 

124: 149–152. 

Kurita N (1979). Cell-mediated immune responses in mice infected with Fonsecaea 

pedrosoi. Mycopathologia 68: 9–15. 

Lavelle P (1980). Chromoblastomycosis in Mexico. Pan American Health 

Organization Scientific Publications 396: 235–247. 

Makarenkov V (2001). T-Rex: reconstructing and visualizing phylogenetic trees and 

reticulation networks. Bioinformatics 17: 664–668. 

Makarenkov V, Legendre P (2004). From a phylogenetic tree to a reticulated 

network. Journal of Computational Biology 11: 195–212. 

Marques SG, Pedroso Silva CM, Saldanha PC, Rezende MA, Vicente VA, Queiros- 

Telles F, Lopes Costa JM (2006). Isolation of Fonsecaea pedrosoi from the shell 

of the babassu coconut (Orbignya phalerata Martius) in the Amazon region of 

Maranhão Brazil. Japanese Journal of Medical Mycology 47: 305–311. 

Martínez E, Rey Valieron C, Yergres F, Reyes R (2005). El caprino: aproximación 

a un modelo animal en la cromomicosis humana. Investigaciόn Clínica 46: 

131–138. 

Mendoza L, Karuppayil SM, Szaniszlo PJ (1993). Calcium regulates in vitro 

dimorphism in chromoblastomycotic fungi. Mycoses 36: 157–164. 

Mostert L, Groenewald JZ, Summerbell RC, Gams W, Crous PW (2006). Taxonomy 

and pathology of Togninia (Diaporthales) and its Phaeoacremonium anamorphs. 


Nishimura K, Miyaji M (1981). Defense mechanisms of mice against Fonsecaea 

pedrosoi infection. Mycopathologia 76: 155–166. 

Nishimura K, Miyaji M, Taguchi H, Wang DL, Li RY, Meng ZH (1989). An ecological 

study on pathogenic dematiaceous fungi from China. In: Current problems of 

opportunistic fungal infections. Proceedings of the 4 th International Symposium 

of the Research Center for Pathogenic Fungi and Microbial Toxicoses, Chiba 

University, Chiba: 17–20. 

Nylander JAA (2004). MrAic.pl. Programme distributed by the author. Evolutionary 


233


Biology Centre, Uppsala University. 

O’Daly JA (1943). La cromoblastomicosis en Venezuela. Memorias Primera Jornada 

Venezolana de Venereología. Caracas. 

O’Donnell K, Cigelnik E (1997). Two divergent intragenomic rDNA ITS2 types within 

a monophyletic lineage of the fungus Fusarium are nonorthologous. Molecular 

and Phylogenetic Evolution 7: 103–116. 

Ohkusu M, Yamaguchi M, Hata K, Yoshida S, Tanaka S, Nishimura K, Hoog GS de, 

Takeo K (1999). Cellular and nuclear characteristics of Exophiala dermatitidis. 


Polak A (1984). Experimental infection of mice by Fonsecaea pedrosoi and Wangiella 

dermatitidis. Sabouraudia 22: 167–169. 

Posada D, Crandall KA (1998). Modeltest: testing the model of DNA substitution. 

Bioinformatics 14: 817–818. 

Prenafeta-Boldú FX, Summerbell R, Hoog GS de (2006). Fungi growing on aromatic 

hydrocarbons: biotechnology’s unexpected encounter with biohazard? FEMS 

Microbiology Reviews 30: 109–130. 

Pritchard JK, Stephens M, Donnelly P (2000). Inference of population structure 

using multilocus genotype data. Genetics 155: 945–959. 

Richard-Yegres N, Yegres F (1987). Cladosporium carrionii en vegetacion xerofila: 

aislamiento en una zona endemica para la cromomicosis en Venezuela. 

Dermatologia Venezuelana 25: 15–18. 

Richard-Yegres N, Yegres, Zeppenfeldt G (1992). Cromomicosis: endemia rural, 

laboral y familiar en Venezuela. Revista Iberoamericana Micologia 9: 38–41. 

Riddley M (1957). The natural habitat of Cladosporium carrionii a cause of 

chromoblastomycosis. Australian Journal of Dermatology 4: 23–27. 

Rubin HA, Bruce S, Rosen T, McBride ME (1991). Evidence for percutaneous 

inoculation as the mode of transmission for chromoblastomycosis. Journal of 

the American Academy of Dermatology 25: 951–954. 

Salgado L, Pereira da Silva J, Picanço Diniz JÁ, Batista da Silva M, Fagundes da 

Costa P, Teixeira C, Salgado UI (2004). Isolation of Fonsecaea pedrosoi from 

thorns of Mimosa pudica, a probable natural source of chromoblastomycosis. 

Revista Instituto de Medicina Tropical, Sao Paulo 46: 33–36. 

Schoch CL, Shoemaker RA, Seifert KA, Hambleton A, Spatafora JW, Crous PW 


Mycologia 98: 1041–1052. 




Silva JP, Rocha RM da, Moreno JS (1995). The coconut babaçu (Orbignya phalerata) 

as a probable risk of human infection by the agent of chromoblastomycosis 

in the State of Maranhão, Brazil. Revista do Sociedad Brasiliera de Medicina 

Tropical, Sao Paulo 28: 49–52. 

Swofford DL (1981). Utility of the distance-Wagner procedure. In: Advances in 

cladistics, vol. 1 (Funk VA and Brooks DR, eds.). Annals of the New York 

Botanical Gardens, New York: 25–44. 

Swofford DL (2003). PAUP*. Phylogenetic Analysis Using Parsimony (*and Other 


Szaniszlo PJ (2002). Molecular genetic studies of the model dematiaceous 

pathogen Wangiella dermatitidis. International Journal of Medical Microbiology 

292: 381–390. 

Tamura K, Nei M (1993). Estimation of the number of nucleotide substitutuions in the 

control region of mitochondrial DNA in humans and chimpanzees. Molecular 

Biolology and Evolution 10: 512–526. 

Trejos A (1953). Cromoblastomycosis experimental en Bufo marinus. Revista de 

Biologia Tropical 1: 39–53. 

Trejos A (1954). Cladosporium carrionii n. sp. and the problem of cladosporia 

isolated from chromoblastomycosis. Revista de Biologia Tropical 2: 75–112. 

Vélez F, Chávez J (1980). Los Cactus de Venezuela. Colecciones Inagro, 

Venezuela. 





Yegres F, Niel F, Gantier JC, Richard-Yegres N (1998). Murine humoral immune 

response against Cladophialophora carrionii and Fonsecaea pedrosoi infection. 

Journal de Mycologie Médical 8: 179–182. 

Yegres F, Richard-Yegres N (2002). Cladophialophora carrionii: Aportes al 

conocimiento de la endemia en Venezuela durante el siglo XX. Revista de la 

Sociedad Venezuelana de Microbiologίa 2: 153–157. 

Yegres F, Richard-Yegres N, Medina-Ruiz E, González-Vivas R (1985). 

Cromomicosis por Cladosporium carrionii en criadores de caprinos del estado 

Falcon. Investigaciόn Clínica 26: 235–246. 

Yegres F, Richard-Yegres N, Nishimura K, Miyaji M (1991). Virulence and 

pathogenicity of human and environmental isolates of Cladosporium carrionii in 

new born ddY mice. Mycopathologia 114: 71–76. 

Yegres F, Richard-Yegres N, Perez-Blanco M (1996). Cromomicosis. In: Temas 

de Micologia Médica. (Bastardo de Albornoz M, ed.). Caracas, Venezuela: 

87–102. 

Yegüez-Rodriguez J, Richard-Yegres N, Yegres F, Rodríguez Larralde A (1992). 

Cromomicosis: susceptibilidad genética en grupos familiares de la zona 

endémica en Venezuela. Acta Cientίfica Venezuelana 43: 98–102. 

Zeng J-S, Sutton DA, Fothergill AW, Rinaldi MG, Harrak MJ, Hoog GS de (2007). 

Spectrum of clinically relevant Exophiala species in the U.S.A. Journal of 

Clinical Microbiology: in press. 

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doi:10.3114/sim.2007.58.09 


Taxonomy, nomenclature and phylogeny of three cladosporium-like hyphomycetes, 

Sorocybe resinae, Seifertia azaleae and the Hormoconis anamorph of Amorphotheca 

resinae 

K.A. Seifert, S.J. Hughes, H. Boulay and G. Louis-Seize 

Biodiversity (Mycology & Botany), Eastern Cereal and Oilseed Research Centre, Agriculture & Agri-Food Canada, Ottawa, Ontario K1A 0C6 Canada 

*Correspondence: Keith A. Seifert, seifertk@agr.gc.ca 

Abstract: Using morphological characters, cultural characters, large subunit and internal transcribed spacer rDNA (ITS) sequences, and provisions of the International Code 

of Botanical Nomenclature, this paper attempts to resolve the taxonomic and nomenclatural confusion surrounding three species of cladosporium-like hyphomycetes. The type 

specimen of Hormodendrum resinae, the basis for the use of the epithet resinae for the creosote fungus {either as Hormoconis resinae or Cladosporium resinae) represents 

the mononematous synanamorph of the synnematous, resinicolous fungus Sorocybe resinae. The phylogenetic relationships of the creosote fungus, which is the anamorph of 

Amorphotheca resinae, are with the family Myxotrichaceae, whereas S. resinae is related to Capronia (Chaetothyriales, Herpotrichiellaceae). Our data support the segregation 

of Pycnostysanus azaleae, the cause of bud blast of rhododendrons, in the recently described anamorph genus Seifertia, distinct from Sorocybe; this species is related to the 

Dothideomycetes but its exact phylogenetic placement is uncertain. To formally stabilize the name of the anamorph of the creosote fungus, conservation of Hormodendrum 

resinae with a new holotype should be considered. The paraphyly of the family Myxotrichaceae with the Amorphothecaceae suggested by ITS sequences should be confirmed 

with additional genes. 

Key words: Amorphothecaceae, Cladosporium resinae, creosote fungus, Hormoconis resinae, jet fuel fungus, kerosene fungus, Myxotrichaceae, Pycnostysanus, resinicolous 

fungi. 

INTRODUCTION 

The ascomycete Amorphotheca resinae Parbery (1969) grows in 

hydrocarbon-rich substrates such as jet fuel, cosmetics and wood 

preserved with creosote or coal tar. This fungus is widely known 

by the anamorph name Hormoconis resinae (Lindau) Arx & G.A. 

de Vries or its obligate synonym Cladosporium resinae (Lindau) 

G.A. de Vries. It produces lightly pigmented, warty conidiophores, 

and branched, acropetally developing chains of lightly pigmented 

ameroconidia lacking conspicuous scars (Fig. 1B–E). This species 

is known colloquially as the “creosote fungus”, the “kerosene 

fungus” or the “jet fuel fungus”; to avoid confusion caused by the 

many heterotypic names with the epithet “resinae”, in this paper 

we generally will use the oldest of these informal names, “creosote 

fungus”, when referring to A. resinae or its anamorph. This fungus 

grows in jet fuel contaminated with small amounts of water, and the 

mycelium clogs fuel lines and corrodes metal parts. Consequently, 

fuel tanks in airports are monitored for this fungus by private 

companies using various physiological or biochemical tests. 

Sorocybe resinae (Fr.) Fr. produces dark black colonies on conifer 

resin, comprising dark synnemata and an effuse mononematous 

synanamorph, both with cladosporium-like conidiogenous cells 

and conidia. Unlike the anamorph of the creosote fungus, the 

conidia of Sorocybe resinae are dark brown and the lateral walls 

are conspicuously thicker than the poles (Fig. 2D–G). Colonies 

with only the mononematous anamorph sometimes occur, and 

the mononematous anamorph can be sparse on colonies bearing 

synnemata. However, the conidia of the mononematous anamorph 

have identical pigmentation and lateral wall thickening to that of 

the synnematous anamorph. The mononematous anamorph rarely 

has been referred to by its own binomial name although, as we will 

show, there is a species epithet available. For the same reasons 

given above for Amorphotheca Parbery, generally we will refer to 

Sorocybe resinae herein as “the resin fungus”. 

Despite the micromorphological differences noted above, there 

is disagreement about whether the creosote fungus is conspecific 

with the mononematous synanamorph of the resin fungus (Parberry 

1969). The name for the anamorph of the creosote fungus is based 

on Hormodendrum resinae Lindau (1906). Christensen et al. 

(1942) presented a study of a cladosporium-like fungus commonly 

isolated from wood impregnated with creosote and coal tar and 

applied Lindau’s name without examining its type. A later ecological 

study by Marsden (1954) employed the same name for the same 

fungus. An extra dimension was added to the confusion when de 

Vries (1952, using the name Cladosporium avellaneum G.A. de 

Vries) described four formae for the creosote fungus (differing in 

the colours of their conidia, the production of setae, or the total 

absence of conidia), each based on single conidium isolates made 

from one parent culture. De Vries (1955) and Parberry (1969) 

examined the holotype of Hormodendrum resinae and concluded 

that it represented the creosote fungus. Hughes (1958), prior to 

the description of Amorphotheca or Hormoconis Arx & G.A. de 

Vries, examined the same specimen and considered it to be the 

mononematous synanamorph of the resin fungus. If Hughes (1958) 

is correct, then neither the species Hormodendrum resinae, nor 

the genus that it typifies, Hormoconis, can represent the creosote 

fungus, as intended by Parberry (1969) or von Arx and de Vries (in 

von Arx 1973). 

In this paper, we present micromorphological, cultural and 

molecular evidence that the resin fungus is a different species from 

the creosote fungus. Combined with re-examination of the holotype 

of Hormodendrum resinae, this information is used to provide a 

revised taxonomy and nomenclature for these two species. A third 

cladosporium-like fungus, Seifertia azaleae, is also considered in 

our discussion of generic concepts. 

Historical review 

The history of the fungus now known as Sorocybe resinae began 

with Fries (1815), who described Racodium resinae Fr. as follows: 

“310. Racodium resinae, expansum molliusculum dense contextum nigrum, filis 

inaequalibus. 

In resina Pini Abietis in silvis Suecia passim. 

Habitu et loco natali distinctum. Fila divaricato-ramosa; alia rigidula apice 

capituli sera, sub miscrosc. Coremio Link similia, Demat. villosum Schleich. huic 

simile; sed sub microsc. fila maxime differunt.” 

235

Seifert et al. 

Fig. 1. Amorphotheca resinae, colony characters and anamorph micromorphology. A. 10-d-old colony on PDA. B, D–E. Micromorphology of conidiophores, showing acropetal 

conidial chains, ramoconidia, and conidia. C. Conidia. DAOM 170427; for C, E see scale bar in D. 

The comparison with Coremium Link indicates the probability of a 

synnematous fungus, and an authentic specimen of Fries’ fungus, 

which as the only known authentic material we interpret as the 

holotype, is preserved in Link’s herbarium (see below). It represents 

the synnematous form of the resin fungus 1 . 

1 Persoon (1822) described a form of R. resinae “β piceum”. Hughes (1968) examined 

the holotype of this form, and it represents the mycelium of the ascomycete 

Strigopodia resinae (Sacc. & Bres.) S.J. Hughes. This taxon is thus not relevant to 

the three species that are the focus of this paper. 

Fries (1832) later transferred his species to Sporocybe 

Fr. (1825), a genus then used for relatively conspicuous dark 

hyphomycetes with dry spores (Mason & Ellis 1953). The 1832 

description explicitly stated... “capitulo rotundato inaequali, sporidiis 

seriatis, stipite aequali simplici.” The use of “capitulo” and “stipite” 

imply what would now be recognised as a synnematous fungus. 

Fries (1832) further characterised the habit of the fungus as “habitu 

stipitum Calicii,” a further comparison to a group of black, stipitate 

lichenized fungi classified in Calicium Pers., which under a hand 

lens look similar to a dark synnematous fungus. 

Fries (1849) next described the genus Sorocybe Fr. for this 

fungus, as follows: 

236

Hormoconis resinae and morphologically similar taxa 

Fig. 2. Sorocybe resinae, synnematous form. A. Colony on bark of living, standing conifer. B. Synnemata. C. Four-month-old colony on DG18. D–G. Acropetally developing 

chains of conidia. Note that the lateral walls are conspicuously thickened; compare with Fig. 3. A, C. DAOM 239134. B, D–G. DAOM 11381. 

Sorocybe Fr. 

Habitus prioris. sed mycelium floccosum densum, stroma corneo-carbonaceum, 

sporis moniliformi-concatenatis basi excipulum incompletum praebens. 

1. S. resinae. Fr. 1–4. at raro fructif. Klotzsch. exs. C. 2. 

Because this description explicitly referred to the Systema, Fries 

presumably was segregating the fungus, originally described 

as Racodium resinae, into a new monotypic genus (McNeill et 

al. 2007; Art. 33.3) and this interpretation of R. resinae as the 

basionym generally has been followed in subsequent treatments 

of Sorocybe resinae. 

As noted in Table 1, Fries’ Racodium resinae was placed in 

several other hyphomycete genera by eighteenth century authors. 

These diversions need not be reviewed in detail here because the 

modern status of these other genera, and their lack of similarity with 

Sorocybe, is clear. 

Bonorden (1851) described Hormodendrum Bonord., with four 

species originally placed in Penicillium Link by Corda (1839); H. 

olivaceum (Corda) Bonord. (≡ Penicillium olivaceum Corda 1839) 

was designated as lectotype by Clements & Shear (1931). This 

genus was frequently, but incorrectly, spelled “Hormodendron”. 

Bonorden’s descriptions and illustrations are of variable quality by 

modern standards, and his herbarium is unknown (Stafleu et al. 

1995). Consequently the actual identities of the species Bonorden 

placed in Hormodendrum are unknown and Corda’s Cladosporium 

olivaceum (Corda) Bonord. was dismissed in Penicillium monographs 

because the drawing shows branched conidial chains 

(Thom 1930), although the specimen has apparently not been 

re-examined. The generic name was used as a segregate for 

Cladosporium Link by some authors (e.g. Kendrick 1961), in 

particular for species with ameroconidia (de Vries 1952). Although it 

sometimes has been considered a synonym of Cladosporium, it will 

remain a nomen dubium until the type species is properly typified. 

Unaware of the resinicolous fungus described by Fries, Lindau 

described two species growing on conifer resin, Pycnostysanus 

resinae Lindau (1904), the type of this anamorph generic name, 

and Hormodendrum resinae Lindau (1906). The former was 

clearly illustrated and described as a synnematous species. The 

protologue of the latter concludes with, “Mit Pycnostysanus resinae 

hat die Art nichts zu tun.” Clearly, Lindau observed no synnemata 

on the specimen of the mononematous fungus and he believed 

it was a different fungus, rather than what would now be called a 

synanamorph of the synnematous fungus that he had described 

previously. Lindau (1910) reproduced the 1904 illustration of 

Pycnostysanus resinae as Stysanus resinae (Fr.) Sacc. (1906), 

thus accepting its identity with the species originally described 

as Racodium resinae Fr. Lindau (1910) made no mention of 

Hormodendrum resinae, indicating he still made no association 

between the synnematous and mononematous fungi on resin. 

De Vries (1952) described a new species, Cladosporium 

avellaneum G.A. de Vries, isolated from cosmetics. Later, he noted 

the similarities between his C. avellaneum and the creosote fungus, 

and suggested that they were the same species (de Vries 1955), 

replacing the name of one of his previously described formae, 

i.e. viride, with the forma name resinae. He examined Lindau’s 

type of Hormodendrum resinae and decided that it provided an 

earlier epithet for C. avellaneum. He transferred the species into 

Cladosporium as C. resinae (Lindau) G.A. de Vries, and this name 

was widely used for the creosote fungus until 1973. This binomial 

is still commonly employed in non-taxonomic literature, especially 

commercial publications dealing with the creosote fungus. 


237


Table 1. Nomenclature and synonymies for the creosote fungus and the resin fungus, showing the use of the same basionym for the two fungi. The “false” names and 

synonymies for the anamorph of the resin fungus are indicated by blue text. The second nomenclatural solution described in the text would have the effect of switching the 

blue text to black for the creosote fungus, and to simultaneously switch the equivalent black text to blue for the mononematous synanamorph of the resin fungus. Holotypes 

we have examined, and the herbarium where they are deposited, are marked with exclamation points, and details of these specimens are noted in Materials and Methods. 

Creosote fungus 

Teleomorph: Amorphotheca resinae Parberry, Australian J. Bot. 17: 340. 1969. 

Anamorph 

Hormodendrum resinae Lindau, in Rabenh. Krypt.-Fl., 2, 1 (Pilze) 8: 699. 1906 (B!). 

≡ Cladosporium resinae (Lindau) G.A. de Vries, Antonie van Leeuwenhoek 21: 167. 1955. 

≡ Hormoconis resinae (Lindau) von Arx & G.A. de Vries, in von Arx, Verh. K. Ned. Akad. Wet., Afd. Natuurk. 61: 62. 1973. 

= Cladosporium avellaneum G.A. de Vries, Contribution to the knowledge of the genus Cladosporium, Uitg. Druk. Hollandia, p. 56, 1952. 

Resin fungus 

Mononematous synanamorph: 

Hormodendrum resinae Lindau, in Rabenh. Krypt.-Fl., 2, 1 (Pilze) 8: 699. 1906 (B!) 

≡ Cladosporium resinae (Lindau) G.A. de Vries, Antonie van Leeuwenhoek 21: 167. 1955. 

≡ Hormoconis resinae (Lindau) von Arx & G.A. de Vries, in von Arx, Verh. K. Ned. Akad. Wet., Afd. Natuurk. 61: 62. 1973. 

Synnematous anamorph: 

Sorocybe resinae (Fr.) Fr., Summa Veg. Scan. 2: 468. 1849. 

≡ Racodium resinae Fr., Obs. Mycol. 1: 216. 1815 (basionym) (B!). 

≡ Sporocybe resinae (Fr.) Fr., Syst. Mycol. 3: 341. 1832. 

≡ Dendryphion resinae (Fr.) Corda, Icon. Fung. 6: 11. 1854. 

≡ Stysanopsis resinae (Fr.) Ferr., Flora Ital. Crypt., 1 (Fungi, Hyphales), p. 187. 1910. 

? = Dematium nigrum Link, Mag. ges. naturf. Fr. 3: 21. 1809 (B!). 

≡ Sporotrichum nigrum (Link) Link, Mag. Ges. naturf. Fr. Berlin 7: 35. 1815. 

= Pycnostysanus resinae Lindau, Verh. Bot. Ver. Brandenb. 45 : 160. 1904 (B!). 

≡ Stysanus resinae (Lindau) Sacc., Syll. Fung. 18: 651. 1906. 

In his study of type collections of classical hyphomycetes, 

Hughes (1958) included Pycnostysanus resinae Lindau and 

Hormodendrum resinae Lindau as facultative synonyms of 

Sorocybe resinae (Fr.) Fr., with Racodium resinae Fr. and several 

other nomenclatural variants as obligate synonyms (Table 1). The 

synnematous Pycnostysanus resinae was cited as “Pycnostysanus 

state [i.e. synanamorph] of Sorocybe resinae”. Hormodendrum 

resinae thus remained to represent the mononematous 

synanamorph of what was interpreted as a single species. 

Parberry (1969) described a cleistothecial ascomycete, 

Amorphotheca resinae, for the teleomorph of the creosote fungus. 

He also examined the holotype of Hormodendrum resinae and 

agreed with the conclusions of de Vries (1955). He used the 

epithet resinae for the teleomorph to correspond with that of the 

anamorph. He discounted the possibility that the synnematous 

Sorocybe resinae could be the same fungus as Hormodendrum 

resinae because synnemata never developed in his cultures of the 

creosote fungus. 

Von Arx and de Vries (in von Arx 1973) described the genus 

Hormoconis, typified by Hormodendrum resinae, with the new 

combination Hormoconis resinae (Lindau) Arx & G.A. de Vries. 

Their intention was to erect an anamorph genus for the anamorph 

of the creosote fungus, which they suggested was improperly 

classified in Cladosporium because it lacked darkened, thickened 

secession scars on the conidia. 

A third cladosporium-like fungus is relevant to this story. 

Seifertia azaleae (Peck) Partridge & Morgan-Jones [until recently 

known as Pycnostysanus azaleae (Peck) E.W. Mason] is a 

cosmopolitan fungus causing bud blast and twig blight of azaleas 

and rhododendrons. This species is morphologically similar to 

Sorocybe resinae, but the conidia are paler and lack laterally 

thickened walls. Sorocybe and Pycnostysanus have often been 

considered taxonomic synonyms (Ellis 1976, Carmichael et al. 

1980); as shown above, both are based on the synnematous form 

of the resin fungus. Partridge and Morgan-Jones (2002) argued that 

Sorocybe resinae and “Pycnostysanus azaleae” are not congeneric, 

and described the new genus Seifertia Part. & Morgan-Jones for 

the Rhododendron fungus. They observed that the connection 

between conidia in Seifertia azaleae is much narrower than in 

Sorocybe resinae, and that minute denticles are visible on the 

conidiogenous cells of the former fungus. The broader connections 

between conidia of Sorocybe resinae result in broadly protuberant 

conidiogenous loci on the conidiogenous cells, and more truncate 

detached conidia. 


Herbarium material and fungal strains 

Full details of herbarium material examined are listed below. 

Cultures and dried herbarium specimens were studied in 90 % 

lactic acid without stains; preparations of some exsiccate and types 

were mounted in glycerin jelly. Cultures were grown on potatodextrose 

agar (PDA, Difco), oatmeal agar (OA, Samson et al. 

2004), Blakeslee’s malt extract agar (MEA, Samson et al. 2004) and 

dichloran-18 % glycerol agar (DG-18, Samson et al. 2004). Colony 

characters were taken from cultures grown at 25 °C in darkness. 

Cultures are maintained in the Canadian Collection of Fungal 

Cultures (DAOM), Agriculture & Agri-Food Canada, Ottawa. 

238


Exsiccati and types 

Dematium nigrum [scr. Link]. E. Hbr. Link (23) = Sporocybe 

resinae III. 341 [scr. ? ] (herb. Link, B). 

Hormodendrum resinae Lindau, n. sp. Fl. v. Hamburg 206, auf 

Harz an Picea excelsa, Sachsenwald, leg. O. Jaap, 29-4-1906. 

[scr. Lindau]. (DAOM 41888, slide prepared from the holotype 

preserved in B.) 

Pycnostysanus resinae Lindau nov. gen. et nov. spec., Kabát et 

Bubák: Fungi imperfecti exsiccati no. 99. Auf erhärteten Fichtenharz 

an Brockenweg, am Dreieckigen Pfahl in Harz, Deutschland, leg. 

G. Lindau, 13.VIII. 1903 (holotype, B). 

Racodium resinae Fries. E. Hbr. Link, Fries legi, Smol. [scr. 

Fries]. (DAOM 41890, slide prepared from herb. Link, B). This 

is the presumed holotype of R. resinae, the basionym for the 

resin fungus, Sorocybe resinae. The specimen includes dark, 

decapitated synnemata, brown conidia with laterally thickened 

walls, and acropetal conidial chains, allowing it to be recognised 

as the fungus we now know as S. resinae. Fries perhaps sent this 

fungus to Link to see if it could be differentiated from Coremium 

Link. The minimal details, that the fungus was collected by Fries, 

presumably in Småland (a province of Sweden), match the details 

in the protologue of this species. 

Sorocybe resinae. “Fungi Rhenani Fasc. II, 1863, L. Fuckel, no. 

129, ad Abietis resinam, raro Hieme, in sylva Hostrichiensi” (as 

Myxotrichum resinae Fr., DAOM 55543 ex FH). “Flora Suecica, 

2956, Ad resinam piceae, Småland: Femsjö, Prostgaidsshogen, 

6 Aug. 1929, leg. J.A. Nannfeldt, s.n.” (as Stysanus resinae (Fr.) 

Sacc., DAOM 41891 ex UPS). “Flora Suecica, 4709, Ad resinam 

abietinum, Uppland: Bondkysko sin Valsätra, 9 May 1932, leg. J.A. 

Nannfeldt” (as Hormodendrum resinae Lindau, DAOM 41889 ex 

UPS). “[ on wood scr. Berkeley] J.E. Vize, Hereford 1877” (as Torula 

pinophila Fr., DAOM 113425 ex K). “Sydow, Mycotheca germanica, 

350. Auf Fichtenharz… am Brockenweg 30.9.1904, leg. P. Sydow” 

(DAOM 41893). 

Other material examined 

Sorocybe resinae. Canada, British Columbia: Burnaby, Central Park, on resin of 

Tsuga heterophylla, leg. S. & L. Hughes, 17 Aug. 2000 (DAOM 228572a, 228573a); 

Cameron Lake, Cathedral Grove, on Pseudotsuga menziesii, leg. isol. S.J. Hughes, 

21 Aug. 1957 (DAOM 56088a). Ladysmith, Ivy Green Park, on resinous exudates, 

leg. R.J. Bandoni no. BC-978, 18 Apr. 1960, det. S.J. Hughes (DAOM 70462). 

North Vancouver, Lynn Valley Conservation Area, leg. det. S.J. Hughes, 1 Jul. 1975 

(DAOM 139385); North Vancouver, Lynn Valley Conservation Area, on bark of living 

conifer (probably Pseudotsuga menziesii), leg. isol. K.A. Seifert no. 1574, 26 May 

2002 (single conidium isolate, culture and specimen DAOM 239134; ITS GenBank 

EU030275, LSU GenBank EU030277); Terrace, near Kalum, on Tsuga heterophylla, 

leg. W.G. Ziller no. V-6549, 10 July 1950, det. S.J. Hughes (DAOM 59657); Queen 

Charlotte Islands, east coast of Moresby Island, north side of Gray Bay, 53°08’ N, 

131°47’ W, on Picea sitchensis, leg. I. Brodo, M.J. Schepanek, W.B. Schofield, 

28 Sep. 1973, det. S.J. Hughes (DAOM 144757); Queen Charlotte Islands, 

Graham Island, Tow Hill area, on resin of Picea sitchensis, leg. S.A. Redhead no. 

4440, 20 Sep. 1982, det. G.P. White (DAOM 184025); Revelstoke, Wigwam, on 

Tsuga heterophylla, leg. W. Ziller V-6567 det. S.J. Hughes, 6 Jun. 1950 (DAOM 

59710); Vancouver Island, Cathedral Grove, Cameron Lake, on Pseudotsuga 

menziesii, leg. det. S.J. Hughes, 21 Aug. 1957 (DAOM 56088a); Vancouver Island, 

Caycuse, on resin of Pseudotsuga menziesii, leg. det. S.J. Hughes, 17 Jul. 1972 

(DAOM 139355); Vancouver Island, Lake Cowichan, Honeymoon Bay, on resin 

of Pseudotsuga menziesii, leg. J Ginns, det. S.J. Hughes, 29 Oct. 1971 (DAOM 

134968); Vancouver Island, Lake Cowichan, Mesachie Lake Forest Experimental 

Station, leg. det. S.J. Hughes, 5 Jul. 1972 (DAOM 139277a, DAOM 139278) and 

6 Jul. 1072 (DAOM 139281). Czechoslovakia, Ještěd near Liberec, leg. det. S.J. 

Hughes, on resin of Larix europaea, 10 May 1955 (DAOM 51723). United States, 

Oregon: Andrews’ Experimental Forest, Forest Service Rd. no 1553, on resin of 

Tsuga heterophylla, leg. det. S.J. Hughes, 10 May 1969 (DAOM 134565); Andrews’ 

Experimental Forest, Blue River, on resin of conifer, cut wood, leg. det. K.A. Seifert 

no. 69, 10 Jul. 1981 (DAOM 228203); Oregon, del Norte Co., J. Smith’s State Park, 

on Tsuga heterophylla, leg. det. S.J. Hughes, 11 May 1069 (DAOM 134614); Devil’s 

Elbow State Park, Cape Perpetus, on Picea sitchensis, leg. det. S.J. Hughes, 6 May 


1969 (DAOM 134615); Linn Co., near Cascadia, on Pseudotsuga menziesii, leg. R. 

Fogel, det. S.J. Hughes, 14 May 1969 (DAOM 127885); U.S. Forest Service Rd. no. 

126, North fork Cape Creek, on resin of Abies grandis, leg. det. S.J. Hughes, 7 May 

1969 (DAOM 134852,134563); Willamette National Forest, McKenzie Bridge Camp 

Grounds, leg. det. S.J. Hughes, 10 May 1969 (DAOM 134564). Washington: Kittitas 

Co., Wanatchee National Forest, Rocky Run, on Abies nobilis, leg. Field Mycology 

Class 1955, 22 Jul. 1955, det. S.J. Hughes, (mononematous synanamorph only, 

DAOM 118934 ex WSP 45210, as Helminthosporium sp.); Jefferson Co., Olympic 

National Forest, 10 mi Camp, Sec. 17, T26N, R3W, on Pseudotsuga mucronata, 

leg. Field Mycology Class, 22 Jul. 1955 (DAOM 113801 ex WSP 45212, as 

Helminthosporium); Grays Harbor Co., Twin Harbors Beach State Park, resin of 

Picea sitchensis, leg. W.B. & V.G. Cooke, 24 Jul. 1951, det. S.J. Hughes (DAOM 

118970 ex WSP 28432). 

Amorphotheca resinae. Isolated from jet fuel by P. Emonds (culture, DAOM 

170427 = ATCC 22711, ITS GenBank EU030278, LSU GenBank EU030280). 

Canada, British Columbia, source unknown, isol. “Mrs. Volkoff”, Jul. 1969 (culture, 

DAOM 194228, ITS GenBank EU030279). 

Seifertia azaleae. All on flower buds of Rhododendron spp. Canada, British 

Columbia: Burnaby, Central Park, leg. S.J. Hughes, 17 Aug. 2000 (DAOM 228571); 

Vancouver, Stanley Park, leg. K.A. Seifert no. 1571, 11 May 2002 (culture and 

specimen, DAOM 239136, LSU GenBank EU030276). Ireland, Munter, Kerry, 

near Glenbeigh (ca. N 52° 03’ W 9° 54’), leg. K.A. Seifert no. 3197, 26 Sep. 2006 

(culture and specimen, DAOM 239135, ITS GenBank EU030273). Netherlands, 

Gelderland, Kröller-Müller Museum, leg. K.A. Seifert no. 1235, 12 May 2000 (DAOM 

227136). United Kingdom, Wales, Hafod Estate (ca. N 52° 22’ W 3° 51’), leg. K.A. 

Seifert no. 3198, 1 Oct. 2006 (culture and specimen, DAOM 239137, ITS GenBank 

EU030274). 

DNA extraction, amplification and sequencing 

DNA was isolated using a FastDNA Kit and the FastPrep 

FP120 (BIO 101 Inc.) or an UltraClean Microbial DNA Isolation 

Kit (Mo Bio Laboratories, Inc., Solana Beach, CA, U.S.A.) using 

mycelium removed from agar cultures. PCR and cycle sequencing 

reactions were performed on a Techne Genius thermocycler 

(Techne Cambridge Ltd.). PCR reactions were performed using 

Ready-To-Go Beads (Amersham Canada Ltd.) in 25 µL volumes, 

each containing 20–100 ng of genomic DNA, 2.5 units pure Taq 

DNA Polymerase, 10 mM Tris-HCl (pH 9.0), 50 mM KCl, 1.5 mM 

MgCl 2 

, 200 µM of each dNTP, 0.2 µL of each primer (50 µM), and 

stabilizers including bovine serum albumin. The reaction profile 

included an initial denaturation for 4 min at 94 °C, followed by 30 

cycles of 1.5 min denaturation at 95 °C, 1 min annealing at 56 °C, 

2 min extension at 72 °C, with a final extension of 10 min at 72 °C. 

Amplicons were purified by ethanol/sodium acetate precipitation 

and resuspended as recommended for processing on an ABI 

PRISM 3100 DNA Analyzer or an ABI 373 Stretch DNA Sequencer 

(Applied Biosystems, Foster, CA). Amplification products were 

sequenced using the BigDye v. 2.0 Terminator Cycle Sequencing 

Ready Reaction Kit (ABI Prism/Applied Biosystems) following the 

manufacturer’s directions. An approximately 1 000 bp portion of the 

large subunit (LSU) ribosomal DNA was amplified and sequenced 

using primers LR0R and LR6, and cycle-sequenced using primers 

LR0R, LR3R, LR16 and LR6 (Vilgalys & Hester 1990, Rehner & 

Samuels 1995; www.biology.duke.edu/fungi/mycolab/primers. 

htm). The complete ITS and 5.8S rRNA genes were amplified 

and sequenced using the primers ITS5 and ITS4, with ITS2 and 

ITS3 primers used for cycle sequencing when necessary (White 

et al. 1990). Some sequences were derived from single PCR 

amplifications of the ITS5–LR6 region. 

Data matrices were subjected to parsimony analysis using 

heuristic searches in PAUP* v. 4.0b10 (Swofford 2002) with 

simple stepwise addition of taxa, and tree bisection-reconnection 

(TBR) branch swapping. Uninformative characters were removed 

for all analyses. Strict consensus trees were calculated, and the 

robustness of the phylogenies was tested using full bootstrap 

analyses (1 000 replications). For all analyses, GenBank accession 

numbers are given on the tree figures, and the sequences generated 

in this study are indicated in bold. 

239


Fig. 3. Hormodendrum resinae, A–B. Conidiophores and acropetally developing chains of conidia. C. Conidia. Note that the lateral walls are conspicuously thickened compared 

to the walls at the poles. From a slide (DAOM 41888) prepared from the holotype (B). 

Byssoascus striatisporus U17912 

99 

96 

10 changes 

88 88 

99 “Mycosphaerella mycopappi” U43480 

* Seifertia azaleae DAOM 239136 EU030276 

Pleomassaria siparia AY004341 

* 

Pleospora herbarum AF382386 

* Cochliobolus heliconiae AF163978 

Setosphaeria monoceras AY016368 

Trematosphaeria heterospora AY016369 

Iodosphaeria aquatica AF452044 

Westerdykella cylindrica AY004343 

Letendraea helminthicola AY016362 

Lojkania enalia AY016363 

Botryosphaeria ribis AY004336 

Capnodium citri AY004337 

Discosphaerina fagi AY016359 



80 Phialophora americana AF050279 

99 79 Capronia mansonii AY004338 

Glyphium elatum AF346420 

97 Terfezia gigantea AF499449 

Sorocybe resinae DAOM 239134 EU030277 

Ascosphaera apis AY004344 

Saccharomyces cerevisiae J01355 

Phaeotrichum benjaminii AY004340 

Calicium viride AF356670 

Pertusaria mammosa AY212831 

73 Pseudocyphellaria perpetua AF401954 

Xylographa vitiligo AY212849 

Neofabraea alba AY064705 

Melanelia exasperatula AJ421436 

* Lophodermium pinastri AY004334 

99 Cudonia lutea AF433138 

Spathularia flavida AF433141 

Ascocoryne cylichnium AF353580 

Bisporella citrina AF335454 

Hormoconis resinae DAOM 170427 EU030280 

Rhytisma acerinum AF356696 

Pleuroascus nicholsonii AF096196 

Golovinomyces cichoracearum AB022360 

Fig. 4. Parsimony analysis of large subunit sequences, demonstrating the phylogenetic positions of Amorphotheca resinae, Sorocybe resinae and Seifertia azaleae (all shown 

in bold) in the Ascomycota. One of 12 equally parsimonious trees (1 888 steps, CI = 0.390, RI = 0.554, RC = 0.216, HI = 0.610) with Golovinomyces cichoracearum as the outgroup. 

Bootstrap values above 70 % are shown at the relevant nodes, with an asterisk representing 100 % bootstrap support; branches with thick lines occurred in all equally 

parsimonious trees. 

Leotiomycetes Dothideomycetes 

240


The large subunit matrix was assembled from the closest 

BLAST matches using our sequences for the three fungi of 

interest, S. resinae, A. resinae and S. azaleae; Golovinomyces 

cichoracearum was added as an out-group to root the tree. 

Although these sequences were put into a single matrix, there 

is no implication that this data set represents the diversity of the 

Ascomycota. The alignment was calculated using MAFFT (Katoh et 

al. 2002) and adjusted using Se-Al (Sequence Alignment Program 

v. 1.d1; http://evolve.zoo.ox.ac.uk/software/Se-Al/main.html) to 

maximise homology. 

The internal transcribed spacers alignment including Sorocybe 

resinae was derived from an alignment of Capronia and related 

anamorphs used by Davey & Currah (2007), originally produced 

using MAFFT. This data set was modified considerably using Se- 

Al to maximise homology, but still included several areas where 

the homology of aligned sequences was difficult to evaluate. ITS 

sequences of Amorphotheca resinae were used to retrieve closely 

related sequences using a BLAST search of GenBank, and these 

relevant sequences were added to an alignment of Oidiodendron 

Robak sequences from the study of Hambleton et al. (1998), and 

then adjusted using Se-Al. 

We attempted direct PCR from two specimens containing only 

the putative mononematous synanamorph of Sorocybe resinae 

(DAOM 228772a, 228573a), to allow comparison of sequences 

obtained from cultures of the synnematous synanamorph. 

These attempts, using the same methods outlined above, were 

unsuccessful. 

RESULTS 

Cultural characters and micromorphology 

Most micromorphological characters of the resin fungus Sorocybe 

resinae (Partridge & Morgan-Jones 2002), the creosote fungus 

Amorphotheca resinae (Parbery 1969, de Vries 1952, 1955, Ho 

et al. 1999) and the rhododendron fungus Seifertia azaleae (Ellis 

1976, Partridge & Morgan-Jones 2002, Glawe & Hummel 2006) are 

well-described in the literature and will not be repeated here. 

The three species are readily distinguished based on growth 

rates and overall cultural phenotypes. Agar colonies of Sorocybe 

resinae are coal-black, wrinkled, and restricted in growth, no matter 

what agar medium is employed; even after 3 mo, the colonies are 

rarely more than 2 cm diam (Fig. 2C). Synnemata did not form in 

our cultures; in vivo, the synnemata produce branched, acropetal 

chains of conidia with laterally thickened walls (Figs 2D–G). No 

thickened, refractive or darkened secession scars were evident 

on individual conidia or ramoconidia. Conidial masses were 

removed from the mononematous and synnematous parts of a 

freshly collected specimen (DAOM 56088a) and grown on PDA 

and sterilised conifer wood. There were no discernable differences 

between colonies derived from the two types of conidiophores, in 

all cases yielding restricted black colonies, or in their microscopic 

characters. Therefore, we conclude that these two types of 

conidiophores represent synanamorphs of one fungus. An identical 

conclusion was reached by Partridge & Morgan-Jones (2002). We 

documented the occurrence of this fungus in California, Oregon, 

and Washington State, U.S.A. and British Columbia, Canada, on 

resinous exudates on Abies nobilis, Picea sitchensis, Pseudotsuga 

menziesii and Tsuga heterophylla. 

Microscopic features from the holotype specimen of 

Hormodendrum resinae Lindau are shown in Fig. 3. Dark, thickwalled 

conidiophore stipes give rise to branched, acropetally 


developing conidial chains. The conidia are relatively darkly 

pigmented, and the lateral walls are more conspicuously thickened 

and darkened than the polar walls. There are no obvious thickened, 

refractive or darkened secession scars on any of the cells. Apart 

from the production of synnemata, the characters of the conidia 

and conidium ontogeny are identical in Lindau’s specimen and the 

synnematous specimens of Sorocybe resinae examined. 

In contrast, both the resin fungus and the rhododendron fungus 

have spreading rather than restricted agar colonies. Cultures of 

the resin fungus are sandy brown (Kornerup & Wanscher 1989), 

planar and powdery, growing 4–4.5 cm diam in 10 d on PDA (Fig. 

1A). Cultures of the rhododendron fungus are slower, growing 

2.5–3.5 cm diam after 21 d on MEA (not shown). They are planar 

and greyish brown, with an orange-brown reverse. No synnemata 

were observed in our cultures of the rhododendron fungus on MEA, 

OA or PDA, but cladosporium-like conidiation occurred in the aerial 

mycelium. 

Phylogeny 

The large subunit analysis (LSU) was used to demonstrate the 

general phylogenetic relationships of the resin fungus Sorocybe 

resinae (DAOM 239134), the creosote fungus Amorphotheca resinae 

(DAOM 170427, 194228) and the rhododendron fungus Seifertia 

azaleae (DAOM 239136), and subsequent analyses of the internal 

transcribed spacers were used to estimate more precise affinities. 

Fig. 4 shows the LSU analysis and demonstrates that Sorocybe 

resinae appears to be a member of the Herpotrichiellaceae, 

Chaetothyriales, A. resinae is related to the inoperculate 

discomycetes (Leotiomycetes) and Seifertia azaleae is most closely 

related to a sequence labelled Mycosphaerella mycopappi A. Funk 

& Dorworth, which is unrelated to Mycosphaerella s. str. 

For the ITS alignment of Sorocybe resinae, two preliminary 

parsimony analyses were conducted, one with informative 

characters from the full alignment, the second with a subset with 

179 characters excluded from seven ambiguously aligned regions. 

The consistency indices (full 0.301, partial 0.324), tree topologies, 

and bootstrap supports for the two analyses were relatively similar. 

Therefore, the complete alignment was used for the tree presented 

here (Fig. 5). The data matrix included 57 taxa, with 352 of 752 

characters phylogenetically informative. Sorocybe resinae clearly 

is related to Capronia and allied anamorph genera, as suggested 

by the LSU analysis. In the ITS analysis (Fig. 6) it forms a wellsupported 

clade with C. villosa Samuels, that is a well-supported 

sister group to species now in three different anamorph genera, 

Phaeococcomyces nigricans (M.A. Rich & A.M. Stern) de Hoog, 

Ramichloridium cerophilum, and an undescribed species of 

Heteroconium Petr. 

The ITS matrix for A. resinae included 42 taxa, with 171 

phylogenetically informative characters in the 530 base alignment. 

The phylogenetic analysis confirmed the relationship of this 

species with the Leotiomycetes, and provided a more precise 

hypothesis of its family-level relationships (Fig. 6). Amorphotheca 

resinae DAOM 170427 and 194228 had identical ITS sequences 

to another strain of the same species reported in GenBank 

(AY251067, from Braun et al. 2003), and one bp substitution from 

a second strain (AF393726 based on the isotype ATCC 200942 

= CBS 406.68). These four sequences formed a sister group 

to two sequences of “Cladosporium” breviramosum Morgan- 

Jones (AF393683, AF393684). The well-supported clade of A. 

resinae and C. breviramosum, which represent the proposed 

family Amorphothecaceae, was previously noted by Braun et 

al. (2003). The nesting of this clade within two well-supported 

241


100 

76 

99 

99 

100 

AY064704 Neofabraea alba 

AF281377 Neofabraea alba 

AF281397 Neofabraea perennans 

AF281388 Neofabraea malicorticis 

AF281365 Neofabraea krawtzewii 

AY129289 Pseudeurotium ovale var. ovale 

AY129290 Pseudeurotium ovale var. milkoi 

AY129287 Pseudeurotium bakeri 

AY129286 Pseudeurotium zonatum 

AY129288 Pseudeurotium desertorum 

AF307760 Geomyces pannorum 

AJ390390 Geomyces asperulatus 

AY789290 Heyderia abietis 

Z81441 Piceomphale bulgarioides 

AY781230 Leptodontidium elatius 

AY348594 Calycina herbarum 

AF062802 Oidiodendron periconioides 

99 

100 

79 

99 

88 

100 

100 

89 

88 

100 

98 

100 

AF062787 Oidiodendron pilicola 

AF062798 Oidiodendron maius 

AF062790 Oidiodendron citrinum 

AF062789 Oidiodendron chlamydosporicum 

AF062797 Oidiodendron griseum 

AF307774 Oidiodendron tenuissimum 

AF062792 Oidiodendron flavum 

AF062810 Myxotrichum arcticum 

AF062809 Oidiodendron truncatum 

AF062808 Oidiodendron tenuissimum 

AF062805 Oidiodendron setiferum 

AF062788 Oidiodendron cerealis 

AJ635314 Oidiodendron myxotrichoides 

AF062811 Myxotrichum cancellatum 

AF062817 Byssoascus striatisporus 

AF062803 Oidiodendron rhodogenum 

AF062814 Myxotrichum deflexum 

88 

A. resinae DAOM 170427, 194228, EU030278-9 

AF393726 Amorphotheca resinae isotype 

AY251067 Amorphotheca resinae 

AF393684 Cladosporium breviramosum 

AF393683 Cladosporium breviramosum 

AF062813 Myxotrichum chartarum 

AF062812 Myxotrichum carminoparum 

AF062816 Myxotrichum stipitatum 

5 changes 

Fig. 5. Parsimony analysis of internal transcribed spacers sequences, demonstrating the position of Amorphotheca resinae (shown in bold) in the ascomycete family 

Myxotrichaceae. One of 44 equally parsimonious trees (645 steps, CI = 0.460, RI = 0.758, RC = 0.349, HI = 0.540) with mid-point rooting. Bootstrap values above 70 % are 

shown at the relevant nodes; branches with thick lines occurred in all equally parsimonious trees. 

Myxotrichaceae Pseudeurotiaceae 

clades of Myxotrichum spp. and the associated anamorph genus 

Oidiodendron, which comprise the family Myxotrichaceae, has not 

been documented previously. 

The ITS sequences of two strains of Seifertia azaleae were 

474 bp and differed by one bp. BLAST searches with these 

sequences revealed significant homologies only with unidentified 

fungi, and lower probability matches with various members of 

the Dothideomycetes. Therefore, no taxonomically meaningful 

phylogenetic analysis can be presented with these ITS sequences. 

The species does seem to have affinities with the Dothideomycetes, 

but the putative relationship with Mycosphaerella, suggested by the 

LSU analysis, could not be confirmed with the ITS analysis. 

DISCUSSION 

Micromorphological comparisons, differences in culture characters, 

and phylogenetic analysis all support the conclusion that the 

mononematous synanamorph of Sorocybe resinae, the resin 

fungus, is different from the anamorph of Amorphotheca resinae, 

the creosote fungus. Based on ribosomal DNA sequences, the 

creosote fungus is related to the family Myxotrichaceae, the genus 

Myxotrichum and its Oidiodendron anamorphs (Fig. 5). In this 

gene tree, Myxotrichum and the Myxotrichaceae are paraphyletic, 

with Amorphotheca and the Amorphothecaceae nested within 

them. Sorocybe appears to be an additional anamorph genus 

phylogenetically associated with Capronia (Herpotrichiellaceae, 

Chaetothyriales, Fig. 6). The genetic connection between the 

synnematous and mononematous morphs of S. resinae was 

verified by morphological comparison of polyspore isolates derived 

from the two synanamorphs. However, the living cultures are no 

longer available and the connection was not confirmed with single 

conidium isolations. The type specimen of Hormodendrum resinae 

(Fig. 3) is the basis for the application of the most frequently 

used anamorph epithet for the creosote fungus. This specimen 

represents the mononematous synanamorph of Sorocybe resinae, 

not the anamorph of Amorphotheca resinae. 

It is difficult to understand how these two fungi were confused 

when their micromorphologies are so different. The conidia are of 

the same general size and shape, but in both morphs of Sorocybe 

resinae (Figs 2D–G, 3C), the lateral walls are conspicuously 

thickened, a condition not present in the creosote fungus (Fig. 1C), 

and the conidia are much darker. In his monograph of Cladosporium, 

de Vries (1952) noted that single conidium isolates of C. avellaneum 

gave rise to four different colony types. In 1955, he extended these 

observations and decided that the much darker resin fungus was 

242

Fig. 6 


87 

89 

74 

100 

100 

83 

93 

94 

100 

93 

Capronia epimyces AY156968 

Capronia epimyces AF050245 

Capronia mansonii AF050247 


Exophiala dermatitidis AY857526 

Exophiala heteromorpha AY857524 

Exophiala heteromorpha AF083207 

88 

97 

94 

Rhinocladiella similis AY857529 

Melanchlenus oligospermus AY163555 

Exophiala oligosperma AY857534 

Exophiala nishimurae AY163560 

Rhinocladiella basitona AY163561 

94 

Exophiala spinifera AM176734 

Ramichloridium mackenziei AY857540 

Rhinocladiella atrovirens AY618683 

Melanchlenus eumetabolus AY163554 

Capronia dactylotricha AF050243 

Ramichloridium anceps DQ826740 

Exophiala lecaniicorni AY857528 

Exophiala mesophila AF542377 

Exophiala attenuata AF549446 

Exophiala pisciphila DQ826739 

Exophiala crusticola AM048755 

Capronia nigerrima AF050251 

Capronia parasitica AF050253 


Capronia pilosella DQ826737 



Cladophialophora carrionii AY857520 

Cladophialophora minourae AY251087 

Phialophora verrucosa AF050283 

Capronia semiimmersa AF050260 

Cladophialophora emmonsii AY857518 

Cladophialophora emmonsii AB109184 

Cladophialophora bantiana AY857517 

Cladophialophora sp. CBS 110551 AY857510 

Cladophialophora devriesii AB091212 

Cladophialophora devriesii AM114417 

Cladophialophora sp. CBS 102230 AY857508 

Cladophialophora sp. CBS 114326 AY 

Cladophialophora arxii AY857509 

Cladophialophora arxii AB109181 

Cladophialophora minourae AB091213 

Cladophialophora sp. AY781217 

Cladophialophora boppii AB109182 

Capronia fungicola AF050246 

83 

90 

94 

99 

98 

99 

80 

100 

100 

100 

Sorocybe resinae DAOM 239134, EU030275 

Capronia villosa AF050261 

Phaeococcomyces nigricans AY843154 


Heteroconium sp. AJ748260 

Phaeococcomyces catenatus AY843041 

Phaeococcomyces chersonesos AJ507323 




10 changes 

Fig. 6. Parsimony analysis of internal transcribed spacers sequences, demonstrating the position of Sorocybe resinae (shown in bold) among species of Capronia 

(Herpotrichiellaceae, Chaetothyriales) and its associated anamorph genera. One of 34 equally parsimonious trees (2 607 steps, CI = 0.301, RI = 0.506, RC = 0.153, HI = 0.699), 

with mid-point rooting. Bootstrap values above 70 % are shown at the relevant nodes; branches with thick lines occurred in all equally parsimonious trees. 

the same as one of his mutant forms of the creosote fungus, 

despite never having isolated such a dark spored form from any of 

his cultures. Parbery (1969) implied that the demonstrated ability 

of the creosote fungus to grow on a diversity of hydrocarbon-rich 

substrates favoured the thought that it would be able to grow on 

conifer resin. If cultures of the true Sorocybe resinae had been 

available, it is unlikely that this confusion would have persisted for 

so long. In vitro, the creosote fungus and the resin fungus are so 

different (Figs 1A, 2C) that it would difficult to defend the idea that 

they were mutants of the same fungus. These differences in the 

cultures are reflected by the disparate phylogenetic affinities of 

what now are clearly demonstrated to be two different species. 

Unfortunately, the name Hormodendrum resinae has been 

misapplied to the creosote fungus, a species of economic 

importance. Also unfortunately, this species is the type of 


Hormoconis, a generic name that the community concerned with 

this fungus has been slow to adapt to in the 30 years since its 

introduction. There are several possible solutions to this problem. 

The conventional solution would be to apply names based strictly 

on the type specimens and accept Hormoconis as a synonym of 

Sorocybe, or to use it as a generic name for the mononematous 

synanamorph of the resin fungus. A new anamorph genus would 

then be described for the creosote fungus, making Cladosporium 

avellaneum G.A. de Vries the basionym for its type. However, the 

resulting binomial would be unfamiliar to those concerned with the 

creosote fungus, and the earlier literature citing H. resinae would 

be misleading. 

A more parsimonious solution is possible. Article 14.9 of the 

International Code of Botanical Nomenclature (McNeil et al. 2006) 

allows for conservation of a name with a different type from that 

243


designated by the authors. The name Hormodendrum resinae is 

not otherwise needed because the mononematous synanamorph 

of the resin fungus is rarely referred to by a Latin binomial, and 

because Sorocybe resinae is based on a different type. Therefore, 

a new type specimen could be proposed and conserved for 

Hormodendrum resinae Lindau, preferably the holotype of A. 

resinae (MELU 7130). This would make the anamorph-teleomorph 

connection unequivocal, maintain current species epithets and 

taxonomic authorities, and ensure that most of the historical 

literature can be interpreted easily without the need to consult 

complicated nomenclators (Table 1). However, by perpetuating the 

use of the epithet “resinae”, this change would also perpetuate the 

misunderstanding that resin is a possible substrate for the creosote 

fungus. In any case, the use of this epithet for the teleomorph of 

the creosote fungus, Amorphotheca resinae, is legitimate and valid, 

and unlikely ever to be changed. 

A third option would be an intermediate one. The application 

of the name Cladosporium avellaneum G.A. de Vries has never 

been in doubt, and it would be possible to conserve this species as 

the type of Hormoconis. This has the advantage of maintaining the 

familiar generic name Hormoconis, in combination with a species 

epithet that has been consistently applied. Furthermore, this solution 

would allow the confusion about the application and correct author 

citation around the epithet “resinae” for the anamorph of creosote 

fungus to recede. 

The second and third solutions require formal taxonomic 

proposals to be published in Taxon. We will argue the merits of 

these possible solutions at more length in that venue. 

The phylogenetic position of A. resinae raises additional 

taxonomic problems. This fungus typifies the monotypic family 

Amorphothecaceae, which has been considered incertae 

sedis since its description by Parbery (1969). Our phylogenetic 

analysis suggests that this family sits within the Myxotrichaceae. 

Amorphothecaceae (1969) is the older name, but Myxotrichaceae 

(1985) is well-entrenched in the mycological literature. As a 

consequence, the Myxotrichaceae are paraphyletic with respect 

to the Amorphothecaceae. The peridium of A. resinae, the only 

species presently placed in this family, lacks the thick-walled 

appendages that characterise most species of the Myxotrichaceae. 

Furthermore, the acropetal-blastic features of the anamorph of 

A. resinae differ from the thallic-arthric conidiogenesis of the 

other anamorphs associated with the Myxotrichaceae, principally 

Oidiodendron. These morphological differences explain why the 

affinity of A. resinae with the Myxotrichaceae was not noted before. 

A formal proposal to conserve Myxotrichaceae as the name for this 

family might be prudent eventually, but this should await analysis of 

additional genes to confirm the phylogenetic relationship. 

Whether Cladosporium breviramosum, originally isolated from 

discoloured wallpaper, is actually a distinct species from A. resinae 

requires further study. It is clear that this species, if it is distinct, 

would be a member of Hormoconis rather than Cladosporium. Apart 

from the study of additional specimens, it might be fruitful to attempt 

to induce an Amorphotheca-like teleomorph in the two available 

cultures of C. breviramosum, and to compare the morphology with 

that of A. resinae. According to Parbery (1969), A. resinae includes 

both homothallic and heterothallic strains. 

Unfortunately, the phylogenetic affinities of Seifertia azaleae 

were not established with certainty in this study. Its closest relative 

in the LSU analysis is a sequence identified as Mycosphaerella 

mycopappi Funk & Dorworth (U43480, based on the apparent 

type culture ATCC 64711), but this sequence does not cluster 

with others representing the family Mycosphaerellaceae (data not 

shown). Similarly, the ITS sequences of the rhododendron fungus 

did not cluster with the many ITS sequences of Mycosphaerella 

available. Presently, it seems that Seifertia azaleae fungus is allied 

with the Dothideomycetes, but its precise affinities are uncertain. It 

is clear that this fungus should not be classified in Pycnostysanus 

(a taxonomic synonym of Sorocybe), and continued recognition of 

the monotypic genus Seifertia seems justified. 


We are grateful to Sarah Hambleton and Marie Davey for providing ITS alignments 

that served as the basis for analyses in this paper. Walter Gams and Scott Redhead 

provided expert nomenclatural advice and helpful reviews of this manuscript. 

We thank Uwe Braun for proposing the third option for resolving the name of the 

creosote fungus, outlined in the text. The Canadian Collection of Fungal Cultures 

(DAOM) provided several of the strains for this study. The curators of DAOM and B 

kindly provided access to critical type specimens. 

REFERENCES 

Arx JA von (1973). Centraalbureau voor Schimmelcultures Baarn and Delft. Progress 

Reports 1972. Verhandelingen der Koninklijke Nederlandsche Akademie van 

Wetenschappen, Afdeeling Natuurkunde 61: 59–81. 

Bonorden HF (1851). Handbuch der allgemeinen Mykologie. Stuttgart. 




Carmichael JW, Kendrick WB, Connors IL, Sigler L (1980). Genera of hyphomycetes. 

University of Alberta Press, Edmonton. 

Christensen CM, Kaufert FH, Schmitz H, Allison JL (1942). Hormodendrum resinae 

(Lindau), an inhabitant of wood impregnated with creosote and coal tar. 

American Journal of Botany 29: 552–558. 

Clements FC, Shear CL (1931). The genera of Fungi. H. W. Wilson, New York. 

Corda, ACJ (1839). Icones fungorum hucusque cognitorum, vol. 3. Prague. 

Davey ML, Currah RS (2007). A new species of Cladophialophora (hyphomycetes) 

from boreal and montane bryophytes. Mycological Research 111: 106–116. 



Fries EM (1815). Observationes mycologicae, vol. 1, p. 216. 

Fries EM (1832). Systema Mycologicum, vol. 3(2). E. Moritz, Greifswald. 

Fries EM (1849). Summa vegatabilium Scandinaviae,vol. 2: 468. 

Glawe DA, Hummell RL (2006). New North American host records for Seifertia 

azaleae. Pacific Northwest Fungi 1(5): 1–6. 

Hambleton S, Egger KN, Currah RS (1998). The genus Oidiodendron: species 

delimitation and phylogenetic relationships based on nuclear ribosomal DNA 

analysis. Mycologia 90: 854–869. 



72: 115–157. 



Hughes SJ (1968). Strigopodia. Canadian Journal of Botany 46: 1099–1107. 

Katoh K, Misawa K, Kuma K, Miyata T (2002). MAFFT: a novel method for rapid 

multiple sequence alignment based on fast Fourier transform. Nucleic Acids 

Research 30: 3059–3066. 

Kendrick WB (1961). Hyphomycetes of conifer leaf litter. Hormodendrum 

staurophorum sp. nov. Canadian Journal of Botany 39: 833–835. 

Kornerup A, Wanscher JH (1989). Methuen Handbook of Colour, 3 rd edn. Methuen, 

London. 

Lindau G (1904). Beiträge zur Pilzflora des Harzes. Verhandlungen des Botanischen 

Vereins der Provinz Brandenburg 45: 148–161 (‘1903’). 

Lindau G (1906). Dr. L. Rabenhorst’s Kryptogamen-Flora von Deutschland, 

Oesterreich und der Schweiz. Zweite Auflage. Erster Band: Die Pilze 

Deutschlands, Österreichs und der Schweiz. VIII. Abteilung: Fungi imperfecti: 

Hyphomycetes (erste Hälfte), Lief. 102: 641–704. 

Lindau G (1910). Dr. L. Rabenhorst’s Kryptogamen-Flora von Deutschland, 

Oesterreich und der Schweiz. Zweite Auflage. Erster Band: Die Pilze 

Deutschlands, Österreichs und der Schweiz. IX. Abteilung: Fungi imperfecti: 

Hyphomycetes (zweite Hälfte), Dematiaceae (Phaeophragmiae bis 

Phaeostaurosporae), Stilbaceae, Tuberculariaceae, sowie Nachträge, 

Nährpflanzenverzeichnis und Register. Leipzig. 

244


Marsden DH (1954). Studies of the creosote fungus, Hormodendrum resinae. 

Mycologia 46: 161–183. 

Mason EW, Ellis MB (1953). British species of Periconia. Mycological Papers 56: 

1–127. 

McNeill J, Barrie FR, Burdet HM, Demoulin V, Hawksworth DL, Marhold K, 

Nicolson DH, Prado J, Silva PC, Skog JE, Wiersema JH, Turland NJ (eds) 

(2007). International Code of Botanical Nomenclature (Vienna Code). Regnum 

Vegetabile 146: 1–568. 

Parbery DG (1969). Amorphotheca resinae gen. nov., sp. nov.: The perfect state of 

Cladosporium resinae. Australian Journal of Botany 17: 331–357. 




Persoon CH (1822). Mycologia europaea. Sectio Prima. Erlangae. 

Rehner SA, Samuels GJ (1995). Molecular systematics of the Hypocreales: a 

teleomorph gene phylogeny and the status of their anamorphs. Canadian 

Journal of Botany 73 (Suppl 1): S816–S823. 

Samson RA, Hoekstra ES, Frisvad JC (2004). Introduction to food- and airborne 

fungi. 7 th ed. Centraalbureau voor Schimmelcultures, Utrecht. 

Stafleu FA, Frans A, Mennega EA (1995). Taxonomic Literature. A selective guide 

to botanical publications and collections with dates, commentaries and types. 

Suppl. III (Br - Ca). Regnum Vegetabile 132: 1–550. 

Swofford DL (2002). PAUP*: Phylogenetic analysis using parsimony (*and other 

methods), version 4. Sinauer Associates, Sunderland, Massachusetts. 

Thom C (1930). The Penicillia. Williams & Wilkins, Baltimore. 

Vilgalys R, Hester M (1990). Rapid identification and mapping of enzymatically 

amplified ribosomal DNA from several Cryptococcus species. Journal of 

Bacteriology 172: 4238–4246. 


ex Fries. Thesis, University of Utrecht. 

Vries GA de (1955). Cladosporium avellaneum de Vries, a synonym of Hormodendrum 

resinae Lindau. Antonie van Leeuwenhoek 21: 166–168. 


fungal ribosomal RNA sequences for phylogenetics. In: PCR Protocols: a guide 

to methods and applications. (Innis MA, Gelfand DH, Sninsky JJ, White TJ, 

eds.) Academic Press, San Diego, CA: 315–322. 


245

INDEX OF FUNGAL NAMES 

Arranged according to genus and then epithets. 

Synonymised names are italicised and page numbers of main entries are bolded. A superscript asterisk ( * ) points to pages with illustrations, 

a superscript c ( c ) to pages with a cladogram, a superscript t ( t ) to pages with a table and a superscript k ( k ) to pages with a key to genera or 

species. 

Acidomyces, 12 

Acidomyces richmondensis, 12 

Acladium biophilum, 85 

Acladium herbarum, 105 

Acrotheca acuta, 84 

Acrotheca cerophila, 72 

Acrotheca multispora, 84 

Acrothecium multisporum, 84 

Amorphotheca, 235, 242 

Amorphotheca resinae, 235–236 * , 238 t , 239–241, 242 c , 244 

Annulatascus triseptatus, 62 c , 64 c 

Antennaria cellaris, 75 

Anungitea, 54 k , 185, 203–204, 216 

Anungitea fragilis, 203–204 

Anungitea heterospora, 204 

Anungitea longicatenata, 54 k 

Anungitea rhabdospora, 204 

Anungitea uniseptata, 211 

Anungitopsis, 203–204, 216 

Anungitopsis amoena, 163, 204, 207 

Anungitopsis intermedia, 204, 209 

Anungitopsis speciosa, 186, 188 c , 190 t , 193 c , 203–204, 216 

Apiosporina, 205, 216 

Apiosporina collinsii, 189 c , 191 t , 194 c , 205 

Aporothielavia leptoderma, 36 c –37 c 

Ascochyta pisi, 36 c –37 c 

Ascocoryne cylichnium, 240 c 

Ascosphaera apis, 240 c 

Ascotaiwania hughesii, 62 c , 64 c 

Asperisporium, 31, 216 

Athelia epiphylla, 6 c –7 c , 36 c –37 c , 58, 62 c , 64 c , 188 c , 193 c 

Aureobasidium caulivorum, 158 t 

Batcheloromyces, 4, 12 k 

Batcheloromyces eucalypti, 2 t , 6 c –7 c , 12 

Batcheloromyces leucadendri, 2 t , 6 c –7 c , 12–13 * 

Batcheloromyces proteae, 2 t , 6 c –7 c , 12, 63 c , 65 c , 162 c 

Bispora, 52 k 

Bisporella citrina, 36 c –37 c , 240 c 

Blumeria graminis f. sp. bromi, 36 c –37 c 

Botryosphaeria ribis , 97 c , 240 c 

Botryosphaeria stevensii , 97 c 

Botrytis epichloës, 89 

Byssoascus striatisporus, 240 c , 242 c 

Cadophora elatum, 46 

Calicium, 236 

Calicium viride, 240 c 

Calycina herbarum, 242 c 

Capnobotryella, 11 k –12, 17 

Capnobotryella renispora, 2 t , 5, 6 c –7 c , 12 

Capnodium citri, 240 c 

Capnodium coffeae, 6 c –7 c 

Capnodium salicinum, 2 t , 6 c –7 c 

Capronia, 55, 92, 185, 203, 216, 241–243 

Capronia acutiseta, 188 c , 193 c , 243 c 

Capronia coronata, 62 c , 64 c , 243 c 

Capronia dactylotricha, 243 c 

Capronia epimyces, 243 c 

Capronia fungicola, 243 c 

Capronia hanliniana, 185, 212 

Capronia hystrioides, 185, 212 

Capronia mansonii, 6 c –7 c , 62 c , 64 c , 188 c , 193 c , 240 c , 243 c 

Capronia munkii, 188 c , 193 c , 243 c 

Capronia nigerrima, 243 c 

Capronia parasitica, 243 c 

Capronia pilosella, 188 c , 193 c , 243 c 

Capronia pulcherrima, 62 c , 64 c , 243 c 

Capronia semiimmersa, 243 c 

Capronia villosa, 241, 243 c 

Caproventuria, 185, 216, 205, 230 

Caproventuria hanliniana, 189 c , 194 c , 205, 212 

Caproventuria hystrioides, 185, 212 

Carpoligna, 92 

Carpoligna pleurothecii, 62 c , 64 c , 83 

Castanedaea, 52 k 

Catenulostroma, 12 k , 13–14, 164 

Catenulostroma abietis, 2 t , 6 c –7 c , 14 k –15, 17, 63 c , 65 c 

Catenulostroma castellanii, 2 t , 6 c –7 c , 36 c –37 c 

Catenulostroma chromoblastomycosum, 2 t , 6 c –7 c , 14 k –15 * , 16, 

36 c –37 c 

Catenulostroma elginense, 2 t , 6 c –7 c , 15 k –16 

Catenulostroma excentricum, 10, 14 k , 16 

Catenulostroma germanicum, 2 t , 6 c –7 c , 14 k , 16 * –17, 36 c –37 c 

Catenulostroma macowanii, 2 t , 6 c –7 c , 15 k , 17, 63 c , 65 c , 162 c 

Catenulostroma microsporum, 2 t , 6 c –7 c , 10, 15 k , 17 

Catenulostroma protearum, 14–15 k , 17 

Catenulostroma sp., 2 t 

Ceramothyrium carniolicum, 188 c , 193 c , 240 c 

Cercospora, 1, 31, 164 

Cercospora apii , 97 c 

Cercospora beticola , 97 c , 111 c 

Cercospora cruenta, 162 c 

Cercospora epicoccoides, 11 

Cercospora eucalypti, 26 

Cercospora penicillata, 1 

Cercospora zebrina, 162 c 

Cercosporella centaureicola, 2 t , 6 c –7 c , 63 c , 65 c 

Cercostigmina, 30–31 

Chaetomium globosum, 36 c –37 c 

Chaetomium homopilatum, 36 c –37 c 

Chloridium apiculatum, 68 

Chloridium indicum, 68, 72–73 

Chloridium minus, 76 

Chloridium musae, 57, 72 

Chloridium schulzerii, 84 

Cibiessia, 11 k , 17, 24–25, 30 

Cibiessia dimorphospora, 2 t , 6 c –7 c , 17, 63 c , 65 c 

Cibiessia minutispora, 2 t , 6 c –7 c 

Cibiessia nontingens, 2 t , 6 c –7 c 

Cistella acuum, 214 

246

Cladophialophora, 52 k , 54 k –55, 167, 185–188, 198–200, 204, 216, 

219–220, 226–228, 230–232 

Cladophialophora ajelloi, 231 

Cladophialophora arxii, 243 c 

Cladophialophora australiensis, 186, 187–188 c , 190 t , 193 c –194 * 

Cladophialophora bantiana, 187, 231, 243 c 

Cladophialophora boppii, 231, 243 c 

Cladophialophora brevicatenata, 185, 212 

Cladophialophora carrionii, 185, 187–188 c , 193 c , 219, 220 t –221 t , 

222–223, 224 c –226 c , 227–228 * , 229–230 * , 231–232, 243 c 

Cladophialophora cf. carrionii, 232 

Cladophialophora chaetospira, 54 k , 187–188 c , 190 t , 193 c , 195 * 

Cladophialophora devriesii, 231, 243 c 

Cladophialophora emmonsii, 243 c 

Cladophialophora hachijoensis, 185 

Cladophialophora hostae, 187, 188 c –189, 190 t , 193 c , 195 * –196 * , 

199, 204 

Cladophialophora humicola, 188 c –189, 190 t , 193 c , 196 * 

Cladophialophora minourae, 243 c 

Cladophialophora potulentorum, 188 c , 190 t , 193 c , 197 * –198 

Cladophialophora proteae, 188 c , 190 t , 193 c , 198 * 

Cladophialophora scillae, 188 c , 190 t , 193 c , 198–199 * , 204 

Cladophialophora sp., 223, 225, 243 c 

Cladophialophora sylvestris, 188 c , 190 t , 193 c , 200 * 

Cladophialophora yegresii, 185, 221 t , 224 c , 226 c , 227–228, 229 * – 

230 * , 231–232 

Cladoriella, 55 

Cladoriella eucalypti, 34 t , 36 c –37 c 

Cladosporium, 8, 21, 30, 33, 35, 38–39, 41, 44, 47, 49, 52 k , 55, 

57, 72, 95–96, 103, 105–106 * , 107, 113, 115–116, 137, 142, 

153–155, 157, 161–169, 173, 176, 181, 209, 227, 230, 237– 

238, 242, 244 

“Cladosporium” adianticola, 190 t , 193 c 

Cladosporium alneum, 116–117 

Cladosporium alopecuri, 142 

Cladosporium amoenum, 207 

Cladosporium antarcticum, 108 t , 111 c –112 c , 114 k –115, 116 * –117 * , 

154–155 

Cladosporium araguatum, 43 

Cladosporium argillaceum, 50 

Cladosporium arthoniae, 41, 116 

Cladosporium arthrinioides, 140 

Cladosporium arthropodii, 142 

Cladosporium asterinae, 55, 103 

Cladosporium avellaneum, 235, 237–238 t , 242–244 

Cladosporium breviramosum, 241–242 c , 244 

Cladosporium bruhnei, 2 t , 6 c –7 c , 36 c –37 c , 63 c , 65 c , 97 c , 107–108 t , 

110, 111 c –112 c , 114 k , 118 * –120 * , 145, 154–155, 158 t , 162 c –165 c , 

168 

Cladosporium brunneum, 133 

Cladosporium carpophilum, 205 

Cladosporium castellanii, 43, 44 

Cladosporium cellare, 75 

Cladosporium cerophilum, 72 

Cladosporium chlorocephalum, 95–96, 103 

Cladosporium cladosporioides, 2 t , 6 c –7 c , 36 c –37 c , 63 c , 65 c , 108 t , 

111 c –112 c , 113 k , 140, 147, 150, 153, 158 t , 162 c –165 c , 168, 174 

Cladosporium colocasiae, 162 c 

Cladosporium cubense, 19–20 

Cladosporium dominicanum, 158 t , 162 c –165 c , 166 * , 168 k –169, 

170 * , 175 

Cladosporium elatum, 46 

Cladosporium ferrugineum, 20–21 

Cladosporium fulvum, 55 

Cladosporium fusiforme, 158 t , 162 c –165 c , 166 * , 168 k –169, 171 * 

Cladosporium gracile, 133 

Cladosporium graminum, 133 

Cladosporium halotolerans, 158 t , 162 c –165 c , 166 * –167, 168 k –169, 

172 * –173, 175 

Cladosporium helicosporum, 18 

Cladosporium hemileiae, 55 

Cladosporium herbaroides, 108 t , 111 c –112 c , 114 k , 120, 121 * –123 * , 

154 

Cladosporium herbarum, 8, 33, 96–97 c , 101, 105, 107–108 t , 110, 

111 c –112 c , 113–114 k , 118, 120, 122, 124, 125 * –127 * , 133, 142, 

153–155, 158 t , 162 c , 164–166, 168, 181 

Cladosporium herbarum var. macrocarpum, 105, 129 

Cladosporium hordei, 118 

Cladosporium hypophyllum, 140 

Cladosporium iridis, 36 c –37 c , 97 c , 109 t , 111 c –112 c , 114 k , 125, 127– 

128 * , 135, 137, 153 

Cladosporium langeronii, 159 t , 162 c –165 c , 166 * –167, 168 k , 173– 

174 * 

Cladosporium lichenicola, 116 

Cladosporium licheniphilum, 116 

Cladosporium lichenum, 116 

Cladosporium macrocarpum, 97 c , 105, 107, 109 t –110, 111 c –112 c , 

114 k , 122, 129 * –130 * , 131 * –133, 153–155 

Cladosporium magnusianum, 133, 135 

Cladosporium musae, 55 

Cladosporium nigrellum, 209 

Cladosporium olivaceum, 237 

Cladosporium ossifragi, 97 c , 109 t –110, 111 c –112 c , 114 k , 133 * –134 * , 

135, 154 

Cladosporium oxysporum, 159 t , 162 c –165 c , 168 

Cladosporium paeoniae, 95–96, 101, 103 

Cladosporium paeoniae var. paeoniaeanomalae, 95–96 

Cladosporium pseudiridis, 109 t , 111 c –112 c , 114 k , 135, 136 * –137 * 

Cladosporium psoraleae, 116–117 

Cladosporium psychrotolerans, 159 t , 162 c –165 c , 166 * –167, 168 k , 

175–176 * 

Cladosporium ramotenellum, 107, 109 t , 111 c –112 c , 115 k , 137–138 * , 

139 * –140, 150, 154, 159 t , 163 c 

Cladosporium resinae, 235, 237–238 t 

Cladosporium rigidophorum, 21, 22 

Cladosporium salinae, 159 t , 161, 162 c –165 c , 166 * , 168 k , 175, 

176–177 * 

Cladosporium scillae, 198 

Cladosporium sinuosum, 109 t , 111 c –112 c , 114 k , 140 * –141 * , 142 

Cladosporium sp., 109 t , 111 c , 149, 159 t , 163 c , 165 c 

Cladosporium sphaerospermum, 2 t , 6 c –7 c , 63 c , 65 c , 153, 157, 159 t , 

162 c –165 c , 166 * , 167–168 k , 169, 173–175, 177–178 * , 181 

Cladosporium spinulosum, 109 t , 111 c –112 c , 114 k , 142 * , 145, 154– 

155, 160 t , 162 c –165 c , 166 * , 168 k , 179 * , 180–181 

Cladosporium staurophorum, 55 

Cladosporium strumelloideum, 23 

Cladosporium subinflatum, 109 t , 111 c –112 c , 114 k , 143 * –144 * , 145, 

154, 160 t , 163 c –165 c 

Cladosporium subnodosum, 150 

Cladosporium subtilissimum, 109 t , 111 c –112 c , 114 k , 145 * –146 * , 

147 * , 149–150, 154–155 

Cladosporium tenellum, 97 c , 109 t , 111 c –112 c , 115 k , 137, 140, 148 * – 

149 * , 150, 153–154 

247

Cladosporium tenuissimum, 160 t , 162 c –163 c , 168 

Cladosporium uredinicola, 2 t , 6 c –7 c , 36 c –37 c , 63 c , 65 c , 116, 162 c 

Cladosporium variabile, 97 c , 110 t , 111 c –112 c , 114 k , 131, 150 * –152 * , 

153–154 

Cladosporium velox, 160 t , 162 c –165 c , 166 * , 168 k , 180 * –181 

Clathrosporium intricatum, 189 c , 193 c 

Coccodinium, 28 

Coccodinium bartschii, 2 t , 4–5, 6 c –7 c , 28, 29 * 

Cochliobolus heliconiae, 240 c 

Colletogloeopsis, 24 

Colletogloeopsis blakelyi, 26 

Colletogloeopsis considenianae, 26 

Colletogloeopsis dimorpha, 26 

Colletogloeopsis gauchensis, 26 

Colletogloeopsis molleriana, 10 

Colletogloeopsis nubilosum, 10 

Colletogloeopsis stellenboschiana, 26 

Colletogloeopsis zuluensis, 26 

Colletogloeum nubilosum, 10 

Coniella, 25 

Conioscypha lignicola, 62 c , 64 c 

Conioscyphascus varius, 62 c , 64 c 

Coniosporium, 216 

Coniothecium macowanii, 17 

Coniothecium punctiforme, 17 

Coniothyrium palmarum, 34 t , 36 c –37 c 

Coniothyrium zuluense, 26 

Coremium, 236, 239 

Cryptadelphia groenendalensis, 62 c , 64 c 

Cryptadelphia polyseptata, 62 c , 64 c 

Cryptococcus neoformans, 232 

Cudonia lutea, 240 c 

Cylindrosympodium, 204 

Cylindrosympodium lauri, 189 c –190 t , 194 c , 204 * 

Cylindrosympodium variabile, 205 

Cyphellophora, 201 

Cyphellophora fusarioides, 201 

Cyphellophora guyanensis, 201 

Cyphellophora hylomeconis, 188 c , 190 t , 193 c , 200 * –201 

Cyphellophora laciniata, 188 c , 190 t , 193 c , 201 

Cyphellophora pluriseptata, 201 

Cyphellophora suttonii, 201 

Cyphellophora vermispora, 201 

Davidiella, 5 k , 8, 30, 33, 35, 44, 52 k , 92, 96, 106, 115, 153, 155, 

163, 166, 173, 230 

Davidiella allicina, 2 t , 108 t , 112 c , 115 k , 118, 119 * –120 * 

Davidiella macrocarpa, 109 t , 112 c , 115 k , 129–130 * , 132 * –133 

Davidiella macrospora, 109 t , 112 c , 125, 128 

Davidiella sp., 108 t , 110 t , 111 c –112 c , 155 

Davidiella tassiana, 8, 108 t , 112 c , 115 k , 120, 122, 124 * –125, 126 * , 

158 t , 163 c –165 c , 166, 168 

Davidiella variabile, 110 t , 112 c , 115 k , 151 * , 152–153 

Dematium epiphyllum var. chionanthi, 122, 124 

Dematium graminum, 129, 132 

Dematium herbarum, 8, 105, 122 

Dematium herbarum var. brassicae, 129, 132 

Dematium nigrum, 238 t –239 

Dematium vulgare var. foliorum, 129, 132 

Dematium vulgare var. typharum, 129, 132 

Dendryphion resinae, 238 t 

Denticularia, 31 

Devriesia, 11 k , 17, 43, 54 k –55, 164, 227, 230 

Devriesia acadiensis, 34 t , 36 c –37 c 

Devriesia americana, 34 t , 36 c –37 c , 42–43 * 

Devriesia shelburniensis, 34 t , 36 c –37 c , 43 

Devriesia staurophora, 5, 6 c –7 c , 17, 36 c –37 c , 63 c , 65 c , 162 c 

Devriesia thermodurans, 34 t , 36 c –37 c 

Dichocladosporium, 53 k , 55, 96, 103 

Dichocladosporium chlorocephalum, 36 c –37 c , 96, 97 c , 98 t* –99 * , 

100 * –101, 102 * 

Didymella bryoniae, 36 c –37 c 

Didymella cucurbitacearum, 36 c –37 c 

Didymellina macrospora, 126 

Digitopodium, 52 k , 55 

Diplococcium, 39 

Discosphaerina fagi, 162 c , 240 c 

Dissoconium, 5 k , 28, 61 

Dissoconium aciculare, 2 t , 6 c –7 c , 28, 63 c , 65 c 

Dissoconium commune, 2 t , 28 

Dissoconium dekkeri, 2 t 

Distocercospora, 31 

Dothidea insculpta, 162 c 

Dothidea ribesia, 162 c 

Dothistroma, 4 

Exophiala, 75, 92, 185, 201, 203, 216, 231 

Exophiala attenuata, 243 c 

Exophiala crusticola, 243 c 

Exophiala dermatitidis, 6 c –7 c , 62 c , 64 c , 188 c , 193 c , 201, 243 c 

Exophiala eucalyptorum, 188 c , 190 t , 193 c , 203 * 

Exophiala heteromorpha, 243 c 

Exophiala jeanselmei, 6 c –7 c , 62 c , 64 c , 188 c , 193 c 

Exophiala lecaniicorni, 243 c 

Exophiala mesophila, 201, 243 c 

Exophiala nishimurae, 243 c 

Exophiala oligosperma, 6 c –7 c , 188 c , 193 c , 243 c 

Exophiala phaeomuriformis, 201 

Exophiala pisciphila, 188 c , 193 c , 243 c 

Exophiala sp. 1, 188 c , 190 t , 193 c , 201–202 * 

Exophiala sp. 2, 188 c , 190 t , 193 c , 201–202 * 

Exophiala sp. 3, 188 c , 190 t , 193 c 

Exophiala spinifera, 243 c 

Fasciatispora petrakii, 37 c , 188 c , 193 c 

Fonsecaea, 231 

Fonsecaea monophora, 231 

Fonsecaea pedrosoi, 6 c –7 c , 62 c , 64 c , 188 c , 193 c , 219, 232, 240 c 

Friedmanniomyces, 11 k –12, 24 

Fumagospora capnodioides, 2 t 

Fusarium, 231 

Fusicladium, 17, 52 k , 54 k –55, 91, 96, 103, 163, 176, 185, 188–189, 

197–199, 203–204, 205, 211, 216 

Fusicladium africanum, 189 c –190 t , 194 c , 205–206 * , 211 

Fusicladium amoenum, 162 c , 189 c –190 t , 194 c , 206 * –207 

Fusicladium brevicatenatum, 212 

Fusicladium carpophilum, 189 c –190 t , 194 c 

Fusicladium caruanianum, 208 

Fusicladium catenosporum, 189 c , 191 t , 194 c 

Fusicladium convolvularum, 189 c , 191 t , 194 c , 207 * –208 

Fusicladium effusum, 161, 162 c –163, 189 c , 191 t , 194 c 

Fusicladium fagi, 189 c , 191 t , 194 c , 208 * –209 

“Fusicladium hyphopodioides”, 215 

Fusicladium intermedium, 189 c , 191 t , 194 c , 209 * 

Fusicladium mandshuricum, 189 c , 191 t , 194 c , 210 

Fusicladium matsushimae, 209 

Fusicladium oleagineum, 189 c , 191 t , 194 c 

248

Fusicladium phillyreae, 189 c , 191 t , 194 c 

Fusicladium pini, 189 c , 191 t , 194 c , 210 * , 211–212 

Fusicladium pomi, 189 c , 191 t , 194 c 

Fusicladium radiosum, 189 c , 191 t , 194 c 

Fusicladium ramoconidii, 189 c , 191 t , 194 c , 211 * –212 * 

Fusicladium rhodense, 189 c , 191 t , 194 c , 212–213 * 

Fusicladium scillae, 198–199 

Fusicladium sp., 212 

Fusicladosporium, 205 

Geomyces asperulatus, 242 c 

Geomyces pannorum, 242 c 

Glyphium elatum, 188 c , 193 c , 240 c 

Golovinomyces cichoracearum, 240 c –241 

Gomphinaria, 84 

Gomphinaria amoena, 84 

Graphiopsis chlorocephala, 96 

Hansfordia biophila, 85 

Hansfordia torvi, 85 

Haplographium chlorocephalum, 96 

Haplographium chlorocephalum var. ovalisporum, 96 

Haplotrichum, 54 k 

Helminthosporium, 239 

Helminthosporium variabile, 150 

Hendersonia grandispora, 11 

Heteroconium, 54 k , 187, 241 

Heteroconium chaetospira, 187, 216 

Heteroconium citharexyli, 187 

Heteroconium sp., 243 c 

Heterosporium, 33, 142, 153 

Heterosporium iridis, 125 

Heterosporium magnusianum, 133, 135 

Heterosporium ossifragi, 133, 135 

Heterosporium variabile, 150 

Heyderia abietis, 242 c 

Hormiactis, 51 k 

Hormoconis, 54 k , 235, 238, 243–244 

Hormoconis resinae, 34 t , 36 c –37 c , 235, 238 t , 240 c , 243 

“Hormodendron”, 237 

Hormodendrum, 237 

“Hormodendrum” elatum, 47 

Hormodendrum hordei, 118, 120 

Hormodendrum langeronii, 167, 173–174 

Hormodendrum olivaceum, 237 

Hormodendrum resinae, 54 k , 235, 237–238 t , 239–240 * , 241–244 

Hormodendrum viride, 237 

Hortaea, 12 k , 17 

Hortaea werneckii, 2 t , 6 c –7 c , 17 

Hyalodendriella, 46, 52 k 

Hyalodendriella betulae, 34 t , 36 c –37 c , 46–47 * 

Hyalodendron, 46, 51 k 

Iodosphaeria aquatica, 240 c 

Kirramyces, 24 

Kirramyces destructans, 26 

Kirramyces epicoccoides, 11 

Kirramyces eucalypti, 26 

Kumbhamaya, 201 

Lacazia loboi, 167 

Lecanosticta, 24 

Leptodontidium elatius, 242 c 

Leptospora rubella, 36 c –37 c 

Letendraea helminthicola, 240 c 

Loboa loboi, 159 t 

Lojkania enalia, 240 c 

Lophodermium pinastri, 240 c 

Lylea, 51 k 

Madurella mycetomatis, 232 

Magnaporthe grisea, 62 c , 64 c 

Melanchlenus eumetabolus, 243 c 

Melanchlenus oligospermus, 243 c 

Melanelia exasperatula, 240 c 

Metacoleroa, 205, 216 

Metacoleroa dickiei, 62 c , 64 c , 189 c , 194 c , 205 

Metulocladosporiella, 18, 51, 53 k , 55, 103 

Metulocladosporiella musae, 103, 188 c , 193 c , 243 c 

Metulocladosporiella musicola, 188 c , 193 c , 243 c 

Mollisia cinerea, 6 c –7 c 

Mycosphaerella, 1, 5 k , 8, 28, 30, 31–33, 45, 51 k , 55, 91, 96, 105, 

115, 155, 163, 164, 166, 205, 241–242, 244 

Mycosphaerella africana, 9, 36 c –37 c 

Mycosphaerella alchemillicola, 34 t 

Mycosphaerella alistairii, 8–9 

Mycosphaerella ambiphylla, 10 

Mycosphaerella associata, 8–9 

Mycosphaerella aurantia , 97 c 

Mycosphaerella bellula, 8, 10 

“Mycosphaerella” communis, 2 t , 6 c –7 c , 28, 63 c , 65 c 

Mycosphaerella cryptica, 8, 10 

Mycosphaerella dendritica, 8, 10 

Mycosphaerella endophytica, 63 c , 65 c 

Mycosphaerella eucalypti, 8 

Mycosphaerella excentrica, 8, 10 

Mycosphaerella fibrillosa, 10 

Mycosphaerella fimbriata, 8, 10 

Mycosphaerella flexuosa, 10 

Mycosphaerella gamsii, 10 

Mycosphaerella graminicola, 3 t , 6 c –7 c , 63 c , 65 c , 162 c 

Mycosphaerella grandis, 10 

Mycosphaerella gregaria, 63 c , 65 c 

“Mycosphaerella” hyperici, 158 t , 163 c –165 c , 173 

Mycosphaerella intermedia, 63 c , 65 c 

Mycosphaerella irregulariramosa, 36 c –37 c 

Mycosphaerella jonkershoekensis, 8, 10 

Mycosphaerella juvenis, 10 

Mycosphaerella latebrosa, 162 c 

“Mycosphaerella” lateralis, 2 t , 6 c –7 c , 28, 36 c –37 c , 63 c , 65 c , 162 c 

Mycosphaerella macrospora, 126 

Mycosphaerella madeirae, 63 c , 65 c 

Mycosphaerella marksii, 36 c –37 c , 63 c , 65 c 

Mycosphaerella maxii, 8, 10 

Mycosphaerella mexicana, 8, 10 

Mycosphaerella microspora, 10 

Mycosphaerella molleriana, 10 

“Mycosphaerella mycopappi”, 240 c –241, 244 

Mycosphaerella nubilosa, 8, 10 

Mycosphaerella ohnowa, 10 

Mycosphaerella parkii, 63 c , 65 c 

Mycosphaerella parkiiaffinis, 10 

Mycosphaerella parva, 10 

Mycosphaerella perpendicularis, 10 

Mycosphaerella pluritubularis, 10 

Mycosphaerella pseudafricana, 11 

Mycosphaerella pseudocryptica, 8, 11 

Mycosphaerella pseudosuberosa, 8, 11 

Mycosphaerella punctiformis, 6 c –7 c , 28, 36 c –37 c , 63 c , 65 c , 97 c , 

166 

249

Mycosphaerella quasicercospora, 11 

Mycosphaerella readeriellophora, 11 

Mycosphaerella secundaria, 11 

Mycosphaerella stigminaplatani, 30 

Mycosphaerella stramenticola, 11 

Mycosphaerella suberosa, 8, 11 

Mycosphaerella suttonii, 11 

Mycosphaerella tassiana, 122, 166 

Mycosphaerella toledana, 11 

Mycosphaerella tulasnei, 132 

Mycosphaerella vespa, 10 

Mycosphaerella walkeri, 63 c , 65 c 

Mycovellosiella, 31, 53 k 

Myrmecridium, 61 k , 84, 92 

Myrmecridium flexuosum, 59 t , 62 c , 64 c , 84–85, 86 * 

Myrmecridium schulzeri, 59 t , 62 c , 64 c , 68 * , 84–85 * 

Myrmecridium schulzeri var. schulzeri, 84 

Myrmecridium schulzeri var. tritici, 84 

Myxotrichum, 242 

Myxotrichum arcticum, 242 c 

Myxotrichum cancellatum, 242 c 

Myxotrichum carminoparum, 242 c 

Myxotrichum chartarum, 242 c 

Myxotrichum deflexum, 36 c –37 c , 242 c 

Myxotrichum resinae, 239 

Myxotrichum stipitatum, 242 c 

Napicladium ossifragi, 133, 135 

Neofabraea alba, 36 c –37 c , 240 c , 242 c 

Neofabraea krawtzewii, 242 c 

Neofabraea malicorticis, 36 c –37 c , 189 c , 193 c , 242 c 

Neofabraea perennans, 242 c 

Neoovularia, 87 

Nothostrasseria, 12 k 

Nothostrasseria dendritica, 2 t , 6 c –7 c , 10 

Ochrocladosporium, 46, 54 k –55 

Ochrocladosporium elatum, 34 t , 36 c –37 c , 46, 48 * –49 

Ochrocladosporium frigidarii, 34 t , 36 c –37 c , 47–49 * 

Oidiodendron, 241–242, 244 

Oidiodendron cerealis, 242 c 

Oidiodendron chlamydosporicum, 242 c 

Oidiodendron citrinum, 242 c 

Oidiodendron flavum, 242 c 

Oidiodendron griseum, 242 c 

Oidiodendron maius, 242 c 

Oidiodendron myxotrichoides, 242 c 

Oidiodendron periconioides, 242 c 

Oidiodendron pilicola, 242 c 

Oidiodendron rhodogenum, 242 c 

Oidiodendron setiferum, 242 c 

Oidiodendron tenuissimum, 242 c 

Oidiodendron truncatum, 242 c 

Ophiostoma stenoceras, 62 c , 64 c 

Paracercospora, 31 

Paracoccidioides loboi, 167 

Parahaplotrichum, 54 k 

Parapericoniella, 53 k , 55, 103 

Parapleurotheciopsis, 51, 53 k –54 k , 204 

Parapleurotheciopsis coccolobae, 51, 54 k 

Parapleurotheciopsis inaequiseptata, 34 t –35, 37 c , 51 

Passalora, 28, 31, 44–45, 53 k , 216 

Passalora arachidicola , 97 c 

Passalora daleae, 34 t , 36 c –37 c 

Passalora dodonaeae, 162 c 

Passalora eucalypti, 36 c –37 c 

Passalora fulva, 36 c –37 c , 55, 97 c , 162 c 

Passalora perplexa, 30 

“Passalora” zambiae, 2 t , 6 c –7 c , 28 

Paullicorticium ansatum, 6 c –7 c , 36 c –37 c , 58, 62 c , 64 c , 188 c , 193 c 

Penicillium, 237 

Penicillium olivaceum, 237 

Penidiella, 12 k , 17–18, 21–22, 24, 53 k 

Penidiella columbiana, 2 t , 6 c –7 c , 17–18 k , 19 * –20 * , 36 c –37 c 

Penidiella cubensis, 18 k , 19–21 * 

Penidiella nectandrae, 3 t , 18 k , 20–21 * , 36 c –37 c 

Penidiella rigidophora, 3 t , 18 k , 21, 22 * –23 * , 36 c –37 c 

Penidiella strumelloidea, 3 t , 18 k , 23, 23 * –24 * , 36 c –37 c , 53 k 

Penidiella venezuelensis, 3 t , 18 k , 22, 24–25 * , 36 c –37 c 

Periconia chlorocephala, 95–96, 101, 103 

Periconia ellipsospora, 96 

Periconia velutina, 64 

Periconiella, 17, 53 k , 57–58, 61 k –62, 63, 91 

Periconiella arcuata, 59 t , 63 c , 65 c –66 * , 68 * 

Periconiella levispora, 59 t , 63 c , 65 c , 67 * –68 

Periconiella musae, 57, 72 

Periconiella papuana, 60, 80 

Periconiella velutina, 17, 57, 59 t –63 c , 64–65 c , 66 * 

Pertusaria mammosa, 240 c 

Phaeoblastophora, 52 k 

Phaeococcomyces catenatus, 188 c , 193 c , 243 c 

Phaeococcomyces chersonesos, 243 c 

Phaeococcomyces nigricans, 241, 243 c 

Phaeoisariopsis, 4, 31 

Phaeophleospora, 4, 24 

Phaeophleospora destructans, 26 

Phaeophleospora epicoccoides, 11 

Phaeophleospora eucalypti, 26 

Phaeophleospora toledana, 11, 25 

Phaeoramularia, 31, 53 k , 227 

“Phaeoramularia” hachijoensis, 43, 185, 214 

Phaeoseptoria eucalypti, 11 

Phaeoseptoria luzonensis, 11 

Phaeosphaeria avenaria, 36 c –37 c 

Phaeotheca triangularis, 3 t , 6 c –7 c 

Phaeothecoidea, 11 k 

Phaeothecoidea eucalypti, 3 t , 6 c –7 c 

Phaeotrichum benjaminii, 240 c 

Phialophora, 231 

Phialophora americana, 188 c , 193 c , 240 c 

Phialophora verrucosa, 243 c 

Phlogicylindrium, 216 

Phlogicylindrium eucalypti, 37 c , 188 c , 193 c , 214 

Phoma herbarum, 36 c –37 c 

Phyllactinia guttata, 132 

Phyllactinia sp., 110 

Piceomphale bulgarioides, 242 c 

Pidoplitchkoviella terricola, 188 c , 193 c 

Pilidiella, 25 

Plectosphaera eucalypti, 37 c 

Pleomassaria siparia, 240 c 

Pleospora herbarum, 240 c 

Pleuroascus nicholsonii, 240 c 

Pleurophoma sp., 3 t 

Pleurophragmium acutum, 84 

Pleurophragmium tritici, 84 

Pleurotheciopsis, 18, 51 

250

Ramichloridium epichloës, 60, 89 

Pleurothecium, 61 k , 83, 92 

243 c Rhinocladiella sp., 60 t , 62 c , 64 c 

Pleurothecium obovoideum, 59 t , 62 c , 64 c , 80, 83 * , 92 

Pollaccia, 185, 205 

Pollaccia mandshurica, 210 

Pollaccia sinensis, 210 

Polyscytalum, 51 k , 55 k , 185, 200, 214, 216 

Polyscytalum fecundissimum, 188 c , 191 t , 193 c , 200, 214 * 

Polyscytalum griseum, 200 

Prathigada, 31 

Protoventuria alpina, 186, 189 c , 191 t , 193 c 

Pseudeurotium bakeri, 242 c 

Pseudeurotium desertorum, 242 c 

Pseudeurotium ovale var. milkoi, 242 c 

Pseudeurotium ovale var. ovale, 242 c 

Pseudeurotium zonatum, 242 c 

Pseudocercospora, 4, 12, 30–31, 164 

Pseudocercospora angolensis, 162 c 

Pseudocercospora epispermogonia, 63 c , 65 c 

Pseudocercospora paraguayensis, 36 c –37 c 

Pseudocercospora protearum, 162 c 

Pseudocercosporidium, 31 

Ramichloridium fasciculatum, 60, 75, 77 

Ramichloridium indicum, 59 t –61, 63 c , 65 c , 73 * 

Ramichloridium mackenziei, 58, 59 t –60 t , 75, 80, 83, 188 c , 193 c , 

243 c 

Ramichloridium musae, 57–59 t , 60–61, 63 c , 65 c , 70 * , 72–73, 91 

Ramichloridium obovoideum, 60, 83 

Ramichloridium pini, 58–59 t , 60–61, 63 c , 65 c , 74 

Ramichloridium schulzeri, 58, 84 

Ramichloridium schulzeri var. flexuosum, 60, 84 

Ramichloridium schulzeri var. schulzeri, 60 

Ramichloridium schulzeri var. tritici, 84 

Ramichloridium strelitziae, 59 t , 63 c , 65 c , 73 * –74 * 

Ramichloridium subulatum, 60, 89 

Ramularia, 51 k , 164 

Ramularia aplospora, 34 t , 36 c –37 c 

Ramularia endophylla, 28 

Ramularia miae, 6 c –7 c , 63 c , 65 c 

Ramularia pratensis var. pratensis, 3 t , 6 c –7 c , 63 c , 65 c 

Ramularia sp., 3 t , 6 c –7 c , 63 c , 65 c 

Ramularia torvi, 85 

Pseudocladosporium, 17, 43, 52 k , 54 k –55, 185, 186, 188–189, Ramulispora, 4, 31 

197–198, 204–205, 216, 227, 230 

Pseudocladosporium brevicatenatum, 205, 212 

Pseudocladosporium caruanianum, 208 

Pseudocladosporium hachijoense, 43, 212 

Pseudocladosporium matsushimae, 209 

Pseudocladosporium proteae, 198 

Pseudocyphellaria perpetua, 240 c 

Pseudodidymaria, 87 

Pseudomassaria carolinensis, 37 c 

Pseudomicrodochium, 201 

Pseudomicrodochium aciculare, 201 

Pseudotaeniolina, 11 k –12, 24 

Pseudotaeniolina convolvuli, 24 

Pseudotaeniolina globosa, 3 t , 6 c –7 c , 24 

Pseudovirgaria, 61 k , 87, 92 

Pseudovirgaria hyperparasitica, 59 t , 63 c , 65 c , 67 * , 86 * –87 

Psilobotrys schulzeri, 84 

Pycnostysanus, 238, 244 

Pycnostysanus azaleae, 238 

Pycnostysanus resinae, 237–238 t , 239 

Rachicladosporium, 38–39, 52 k , 55 

Rachicladosporium luculiae, 34 t , 36 c –37 c , 38 * –39 

Racodium, 75 

Racodium cellare, 75 

Racodium resinae, 235, 237–238 t , 239 

Racodium resinae (β) piceum, 236 

Racodium rupestre, 75 

Radulidium, 61 k , 89, 92 

Radulidium epichloës, 59 t , 63 c , 65 c , 87–88 * , 89 

Radulidium sp., 59 t , 63 c , 65 c 

Radulidium subulatum, 59 t , 63 c , 65 c , 70 * , 87–88 * , 89 

Ramichloridium, 17, 57–58, 60–61 k , 68, 75, 84, 89, 91–92 

Ramichloridium anceps, 60, 75, 188 c , 193 c , 243 c 

Ramichloridium apiculatum, 17, 57, 59 t –61, 63 c , 65 c , 68–69 * , 73 

Ramichloridium australiense, 59 t , 63 c , 65 c , 69 * –70 * 

Ramichloridium basitonum, 58, 60, 75, 77 

Ramichloridium biverticillatum, 59 t , 61, 63 c , 65 c , 71 * –72, 91 

Ramichloridium brasilianum, 59 t , 63 c , 65 c , 70 * –71 * , 72 

Ramulispora sorghi, 162 c 

Rasutoria pseudotsugae, 63 c , 65 c 

Rasutoria tsugae, 63 c , 65 c 

Readeriella, 11 k –12 k , 17, 24–25, 30 

Readeriella blakelyi, 6 c –7 c , 26 

Readeriella brunneotingens, 3 t , 6 c –7 c , 26–27 * 

Readeriella considenianae, 6 c –7 c , 26, 63 c , 65 c 

Readeriella destructans, 3 t , 6 c –7 c , 26 

Readeriella dimorpha, 6 c –7 c , 26 

Readeriella epicoccoides, 6 c –7 c , 11, 30 

Readeriella eucalypti, 3 t , 6 c –7 c , 26 

Readeriella gauchensis, 3 t , 6 c –7 c , 26 

Readeriella mirabilis, 3 t , 6 c –7 c , 24–25, 27 * , 30, 97 c 

Readeriella molleriana, 3 t , 10 

Readeriella novaezelandiae, 6 c –7 c , 97 c 

Readeriella nubilosa, 10 

Readeriella ovata, 3 t , 6 c –7 c 

Readeriella pulcherrima, 6 c –7 c , 26 

Readeriella readeriellophora, 11, 25–27 * 

Readeriella stellenboschiana, 3 t , 6 c –7 c , 26 

Readeriella toledana, 11 

Readeriella zuluensis, 3 t , 6 c –7 c , 26 

Repetophragma goidanichii, 62 c , 64 c 

Retroconis, 46 

Retroconis fusiformis, 34 t , 36 c –37 c , 46 

Rhinocladiella, 57–58, 60–61 k , 75–76, 92 

Rhinocladiella anceps, 59 t , 62 c , 64 c , 75 * –76 

Rhinocladiella apiculata, 68 

Rhinocladiella atrovirens, 62 c , 64 c , 75–76, 243 c 

Rhinocladiella basitona, 59 t , 62 c , 64 c , 76 * –77, 243 c 

Rhinocladiella cellaris, 75 

Rhinocladiella elatior, 89 

Rhinocladiella fasciculata, 59 t , 62 c , 64 c , 77 * 

Rhinocladiella indica, 68, 73 

Rhinocladiella mackenziei, 62 c , 64 c , 78 * , 80, 83, 92 

Rhinocladiella obovoidea, 83 

Rhinocladiella phaeophora, 60 t , 62 c , 64 c , 76 

Rhinocladiella schulzeri, 84 

Ramichloridium cerophilum, 59 t –61, 63 c , 65 c , 71 * –72, 73, 241, Rhinocladiella similis, 243 c 

251

Rhinotrichum multisporum, 84 

Rhizocladosporium, 50–51, 54 k –55, 204 

Rhizocladosporium argillaceum, 34 t , 36 c –37 c , 50 * –51 

Rhodoveronaea, 61 k , 89–90, 92 

Rhodoveronaea varioseptata, 60 t , 62 c , 64 c , 70 * , 90–91 

Rhytisma acerinum, 240 c 

Saccharomyces cerevisiae, 240 c 

Sarcoleotia turficola, 36 c –37 c 

Satchmopsis brasiliensis, 189 c , 193 c 

Schizothyrium, 5 k , 28, 30 

Schizothyrium acerinum, 28 

Schizothyrium pomi, 6 c –7 c 

Scolicotrichum iridis, 125 

Scorias spongiosa, 6 c –7 c 

Seifertia, 52 k , 238, 244 

Seifertia azaleae, 235, 238–239, 240 c , 241–242, 244 

Septocylindrium chaetospira, 187 

Septonema, 54 k 

Septonema chaetospira, 187 

Septoria, 164, 173 

Septoria pulcherrima, 26 

Septoria tritici, 3 t , 6 c –7 c , 63 c , 65 c 

Setosphaeria monoceras, 240 c 

Sonderhenia, 24 

Sorocybe, 52 k , 54 k , 236–238, 242–244 

Sorocybe resinae, 235, 237 * –238 t , 239–240 c , 241–243 c , 244 

Spadicoides minuta, 187 

Spathularia flavida, 240 c 

Sphaerella allicina, 118 

Sphaerella cryptica, 10 

Sphaerella molleriana, 10 

Sphaerella nubilosa, 10 

Sphaerella tassiana, 8, 122 

Sphaeria allicina, 118 

Spilocaea, 185, 205 

Spilomyces dendriticus, 10 

Sporidesmium pachyanthicola, 63 c , 65 c 

Sporocybe, 236 

Sporocybe resinae, 237–238 t , 239 

Sporotrichum anceps, 76 

Sporotrichum nigrum, 238 t 

Stagonospora pulcherrima, 26 

Staninwardia, 12 k , 26 

Staninwardia breviuscula, 26 

Staninwardia suttonii, 5, 6 c –7 c , 26, 36 c –37 c , 63 c , 65 c 

Stenella, 31, 44–45, 53 k , 75, 227 

Stenella araguata, 16, 19, 24, 43, 44 * –45 * , 75 

“Stenella” cerophilum, 36 c –37 c 

Stenellopsis, 31 

Stigmidium, 30 

Stigmidium schaereri, 30 * 

Stigmina, 4, 12, 30–31 

Stigmina eucalypti, 12 

Stigmina platani, 12 

Strigopodia resinae, 236 

Stysanopsis resinae, 238 t 

Stysanus resinae, 237–238 t , 239 

Subramaniomyces, 51, 54 k , 204 

Subramaniomyces fusisaprophyticus, 34 t –35, 37 c , 51 

Subramaniomyces simplex, 51 

Sympodiella, 204 

Sympoventuria, 185, 204 

Sympoventuria capensis, 189 c , 191 t , 194 c , 205 

Taeniolella, 41, 52 k 

Taeniolina, 52 k 

Taeniolina scripta, 17 

Teratosphaeria, 5 k , 8, 28, 30–31, 43, 45, 92, 96 

Teratosphaeria africana, 8 

Teratosphaeria alistairii, 6 c –7 c , 9, 36 c –37 c , 63 c , 65 c 

Teratosphaeria associata, 9 

Teratosphaeria bellula, 3 t , 6 c –7 c , 10 

Teratosphaeria cryptica, 6 c –7 c , 10, 36 c –37 c 

Teratosphaeria dendritica, 2 t , 10 

Teratosphaeria excentrica, 10 

Teratosphaeria fibrillosa, 3 t –4, 6 c –7 c , 8–9 * , 10, 63 c , 65 c 

Teratosphaeria fimbriata, 10 

Teratosphaeria flexuosa, 6 c –7 c , 10 

Teratosphaeria gamsii, 10 

Teratosphaeria jonkershoekensis, 10 

Teratosphaeria juvenis, 36 c –37 c 

Teratosphaeria maxii, 6 c –7 c , 10 

Teratosphaeria mexicana, 3 t , 6 c –7 c , 10 

Teratosphaeria microspora, 2 t , 8, 10, 15 k , 97 c 

Teratosphaeria molleriana, 3 t , 6 c –7 c , 10, 36 c –37 c , 63 c , 65 c 

Teratosphaeria nubilosa, 3 t , 6 c –7 c , 10, 162 c 

Teratosphaeria ohnowa, 4 t , 6 c –7 c , 10 

Teratosphaeria parkiiaffinis, 10 

Teratosphaeria parva, 6 c –7 c , 10, 63 c , 65 c 

Teratosphaeria perpendicularis, 10 

Teratosphaeria pluritubularis, 10 

Teratosphaeria proteaarboreae, 8 

Teratosphaeria pseudafricana, 11 

Teratosphaeria pseudocryptica, 11 

Teratosphaeria pseudosuberosa, 2 t , 6 c –7 c , 11 

Teratosphaeria quasicercospora, 11 

Teratosphaeria readeriellophora, 6 c –7 c , 11, 63 c , 65 c , 97 c 

Teratosphaeria secundaria, 4 t , 6 c –7 c , 11 

Teratosphaeria sp., 2 t , 4 t , 6 c –7 c 

Teratosphaeria stramenticola, 11 

Teratosphaeria suberosa, 6 c –7 c , 11, 63 c , 65 c 

Teratosphaeria suttonii, 3 t , 6 c –7 c , 11, 36 c –37 c 

Teratosphaeria toledana, 6 c –7 c , 11, 63 c , 65 c 

Terfezia gigantea, 240 c 

Thedgonia ligustrina, 34 t , 36 c –37 c 

Thielaviopsis basicola, 159 t 

Thyridium, 84 

Thyridium vestitum, 60, 62 c , 64 c 

Thysanorea, 61 k , 63, 80, 92 

Thysanorea papuana, 60 t , 62 c , 64 c , 68 * , 78 * –79 * , 80, 188 c , 193 c 

Torrendiella eucalypti, 36 c –37 c 

Torula pinophila, 239 

Toxicocladosporium, 39, 53 k , 55 

Toxicocladosporium irritans, 34 t , 36 c –37 c , 39–40 * , 41 

Trematosphaeria heterospora, 240 c 

Trichosphaeria pilosa, 6 c –7 c 

Trimmatostroma, 5, 14, 164 

Trimmatostroma abietina, 14–15, 97 c 

Trimmatostroma abietis, 14–15, 97 c 

Trimmatostroma betulinum, 3 t , 5, 6 c –7 c 

Trimmatostroma elginense, 16 

Trimmatostroma excentricum, 10 

Trimmatostroma macowanii, 17 

Trimmatostroma microsporum, 10 

Trimmatostroma protearum, 17 

252

Trimmatostroma salicis, 3 t , 5, 6 c –7 c , 13 * , 97 c 

Uwebraunia, 28 

Venturia, 43, 52 k , 55, 185–186, 204–205, 216 

Venturia aceris, 189 c , 191 t , 194 c 

Venturia alpina, 189 c , 191 t , 194 c 

Venturia anemones, 189 c , 191 t , 194 c 

Venturia asperata, 62 c , 64 c 

Venturia atriseda, 189 c , 191 t , 194 c 

Venturia aucupariae, 189 c , 191 t , 194 c 

Venturia carpophila, 62 c , 64 c , 189 c –190 t , 194 c 

Venturia cephalariae, 189 c , 191 t , 194 c 

Venturia cerasi, 189 c , 191 t , 194 c 

Venturia chlorospora, 62 c , 64 c , 189 c , 191 t , 194 c 

Venturia crataegi, 189 c , 191 t , 194 c 

Venturia dickiei, 205 

Venturia ditricha, 189 c , 191 t , 194 c 

Venturia fraxini, 189 c , 192 t , 194 c 

Venturia hanliniana, 62 c , 64 c , 205, 212 

Venturia helvetica, 189 c , 192 t , 194 c 

Venturia hystrioides, 189 c , 192 t , 194 c , 212–213 * 

Venturia inaequalis, 62 c , 64 c , 189 c , 191 t , 194 c 

Venturia lonicerae, 189 c , 192 t , 194 c 

Venturia macularis, 189 c , 192 t , 194 c 

Venturia maculiformis, 189 c , 192 t , 194 c 

Venturia mandshurica, 191 t , 210 

Venturia minuta, 189 c , 192 t , 194 c 

Venturia nashicola, 189 c , 192 t , 194 c 

Venturia polygonivivipari, 189 c , 192 t , 194 c 

Venturia populina, 189 c , 192 t , 194 c 

Venturia pyrina, 62 c , 64 c , 189 c , 192 t , 194 c 

Venturia saliciperda, 189 c , 192 t , 194 c 

Venturia sp., 189 c , 192 t , 194 c 

Venturia tremulae, 191 t 

Venturia tremulae var. grandidentatae, 189 c , 192 t , 194 c 

Venturia tremulae var. populialbae, 189 c , 192 t , 194 c 

Venturia tremulae var. tremulae, 189 c , 192 t , 194 c 

Venturia viennotii, 189 c , 192 t , 194 c 

Veronaea, 57–58, 60–61 k , 76, 80, 87, 89, 91, 92 

Veronaea apiculata, 68 

Veronaea botryosa, 60 t , 62 c , 64 c , 76, 80–81 * , 188 c , 193 c 

Veronaea compacta, 60 t , 62 c , 64 c , 81–82 * , 83 

Veronaea harunganae, 92 

Veronaea indica, 73 

Veronaea japonica, 60 t , 62 c , 64 c , 74 * , 82 * 

Veronaea musae, 57, 72 

Veronaea parvispora, 76 

Veronaea simplex, 60, 91 

Veronaea verrucosa, 73 

Veronaeopsis, 61 k , 91–92 

Veronaeopsis simplex, 60 t , 62 c , 64 c , 74 * , 90 * –91, 189 c , 194 c 

Verrucisporota, 31, 45 

Verrucocladosporium, 41, 53 k , 55 

Verrucocladosporium dirinae, 34 t , 36 c –37 c , 41–42 * 

Vibrissea flavovirens, 6 c –7 c 

Vibrissea truncorum, 6 c –7 c 

Virgaria, 87 

Websteromyces, 52 k 

Wentiomyces, 91 

Wentiomyces javanicus, 91 

Westerdykella cylindrica, 240 c 

Xenomeris juniperi, 63 c , 65 c 

Xylographa vitiligo, 240 c 

Xylohypha, 52 k 

Zasmidium, 45, 61, 74–75 

Zasmidium cellare, 60 t –61, 63 c , 65 c , 74 * –75 

Zeloasperisporium, 214, 216 

Zeloasperisporium hyphopodioides, 189 c , 191 t , 194 c , 214–215 * 

Zygosporium, 5 k 

253

The genus Cladosporium and similar dematiaceous ... - CBS - KNAW

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