Evolution of Fungi and Fungal-Like Organisms (The Mycota, 14)

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The Mycota Edited by K. Esser

The Mycota I

Growth, Differentiation and Sexuality 1st edition ed. by J.G.H. Wessels and F. Meinhardt 2nd edition ed. by U. Kuës and R. Fischer

II

Genetics and Biotechnology 1st and 2nd edition ed. by U. Ku¨ck

III

Biochemistry and Molecular Biology 1st and 2nd edition ed. by R. Brambl and G. Marzluf

IV

Environmental and Microbial Relationships 1st edition ed. by D. Wicklow and B. So¨derstro¨m 2nd edition ed. by C.P. Kubicek and I.S. Druzhinina

V

Plant Relationships 1st edition ed. by G. Carroll and P. Tudzynski 2nd edition ed. by H.B. Deising

VI

Human and Animal Relationships 1st edition ed. by D.H. Howard and J.D. Miller 2nd edition ed. by A.A. Brakhage and P.F. Zipfel

VII

Systematics and Evolution 1st edition ed. by D.J. McLaughlin, E.G. McLaughlin, and P.A. Lemke† 2nd edition ed. by D. McLaughlin and J.W. Spatafora

VIII Biology of the Fungal Cell 1st and 2nd edition ed. by R.J. Howard and N.A.R. Gow IX

Fungal Associations Ed. by B. Hock

X

Industrial Applications 1st edition ed. by H.D. Osiewacz 2nd edition ed. by M. Hofrichter

XI

Agricultural Applications Ed. by F. Kempken

XII

Human Fungal Pathogens Ed. by J.E. Domer and G.S. Kobayashi

XIII

Fungal Genomics Ed. by A.J.P. Brown

XIV Evolution of Fungi and Fungal-Like Organisms Ed. by S. Po¨ggeler and J. Wo¨stemeyer XV

Physiology and Genetics: Selected Basic and Applied Aspects Ed. by T. Anke and D. Weber

The Mycota A Comprehensive Treatise on Fungi as Experimental Systems for Basic and Applied Research Edited by K. Esser

XIV

Evolution of Fungi and Fungal-Like Organisms

Volume Editors: S. Po¨ggeler and J. Wo¨stemeyer

Series Editor Professor Dr. Dr. h.c. mult. Karl Esser Allgemeine Botanik Ruhr-Universita¨t 44780 Bochum, Germany Tel.: +49 (234)32-22211 Fax.: +49 (234)32-14211 e-mail: [email protected] Volume Editor Professor Dr. Stefanie Po¨ggeler Institut fu¨r Mikrobiologie und Genetik Georg-August-Universita¨t Go¨ttingen Grisebachstr. 8 37077 Go¨ttingen, Germany Tel.: +49 (551) 391 3930 Fax: +49 (551) 391 0123 e-mail: [email protected] Professor Dr. Johannes Wo¨stemeyer Institut fu¨r Mikrobiologie Friedrich-Schiller-Universita¨t Jena Neugasse 24 07743 Jena, Germany Tel.: +49 (3641) 949310/1 Fax: +49 (3641) 949312 e-mail: [email protected]

ISBN 978-3-642-19973-8 e-ISBN 978-3-642-19974-5 DOI 10.1007/ 978-3-642-19974-5 Springer Heidelberg Dordrecht London New York Library of Congress Control Number: 2011932566 # Springer-Verlag Berlin Heidelberg 2011 This work is subject to copyright. All rights are reserved, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilm or in any other way, and storage in data banks. Duplication of this publication or parts thereof is permitted only under the provisions of the German Copyright Law of September 9, 1965, in its current version, and permission for use must always be obtained from Springer. Violations are liable to prosecution under the German Copyright Law. The use of general descriptive names, registered names, trademarks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. Cover design: Erich Kirchner and WMXDesign GmbH, Heidelberg, Germany Printed on acid-free paper Springer is part of Springer Science+Business Media (www.springer.com)

Karl Esser (born 1924) is retired Professor of General Botany and Director of the Botanical Garden at the Ruhr-Universita¨t Bochum (Germany). His scientific work focused on basic research in classical and molecular genetics in relation to practical application. His studies were carried out mostly on fungi. Together with his collaborators he was the first to detect plasmids in higher fungi. This has led to the integration of fungal genetics in biotechnology. His scientific work was distinguished by many national and international honors, especially three honorary doctoral degrees.

Stefanie Po¨ggeler (born 1963) studied Biology at the Ruhr-Universita¨t in Bochum (Germany). In 1993, she graduated with a thesis on intronencoded polypeptides in plastids and mitochondria under the supervision of Prof. Ulrich Ku¨ck. She later completed her ‘‘Habilitation’’ at the Ruhr-Universita¨t Bochum in 2000 and was awarded the Venia Legendi in Botany. Between 2001 and 2003 she had a stand-in assistant professorship in Botany at the WilhelmsUniversita¨t in Mu¨nster (Germany). In 2006 she was appointed as professor for Genetics of Eukaryotic Microorganisms at the Georg-August Universita¨t Go¨ttingen (Germany). Her work focuses on the analysis of mating type genes and sexual development in filamentous ascomycetes. In a second line of research, she is interested in the evolution of fungal inteins.

Johannes Wo¨stemeyer (born 1951) studied Biology and Chemistry at Justus-LiebigUniversita¨t Gießen and Ruhr-Universita¨t Bochum (Germany). He obtained his PhD with a study on sporulation and gene expression in Bacillus megaterium in 1975. As a postdoc, he worked on transposon genetics in maize and on artificial genes for blood pressure hormones at the universities of Cologne and Hamburg. Since 1982 he concentrated on fungal genetics at the Technical University in Berlin and worked especially on biochemical and genetic analysis of sexual development in Mucor-like fungi. He qualified for a professorship by Habilitation in 1989. After moving to ‘‘Institut fu¨r Genbiologische Forschung’’, an institute cooperatively run by Schering Company and the Senate of Berlin, he added studies in plant pathology to his research interests. In 1993, he accepted a professorship at Friedrich-Schiller-Universita¨t in Jena, where he holds the chair for General Microbiology and Microbial Genetics. Major research interests are molecular biology of sexual communication in fungi, horizontal gene transfer, and the development of in vitro manipulation systems for zygomycetes.

.

Series Preface

Mycology, the study of fungi, originated as a sub discipline of botany and was a descriptive discipline, largely neglected as an experimental science until the early years of this century. A seminal paper by Blakeslee in 1904 provided evidence for self incompatibility, termed “heterothallism”, and stimulated interest in studies related to the control of sexual reproduction in fungi by mating-type specificities. Soon to follow was the demonstration that sexually reproducing fungi exhibit Mendelian inheritance and that it was possible to conduct formal genetic analysis with fungi. The names Burgeff, Kniep and Lindegren are all associated with this early period of fungal genetics research. These studies and the discovery of penicillin by Fleming, who shared a Nobel Prize in 1945, provided further impetus for experimental research with fungi. Thus began a period of interest in mutation induction and analysis of mutants for biochemical traits. Such fundamental research, conducted largely with Neurospora crassa, led to the one gene: one enzyme hypothesis and to a second Nobel Prize for fungal research awarded to Beadle and Tatum in 1958. Fundamental research in biochemical genetics was extended to other fungi, especially to Saccharomyces cerevisiae, and by the mid1960s fungal systems were much favored for studies in eukaryotic molecular biology and were soon able to compete with bacterial systems in the molecular arena. The experimental achievements in research on the genetics and molecular biology of fungi have benefited more generally studies in the related fields of fungal biochemistry, plant pathology, medical mycology, and systematics. Today, there is much interest in the genetic manipulation of fungi for applied research. This current interest in biotechnical genetics has been augmented by the development of DNA-mediated transformation systems in fungi and by an understanding of gene expression and regulation at the molecular level. Applied research initiatives involving fungi extend broadly to areas of interest not only to industry but to agricultural and environmental sciences as well. It is this burgeoning interest in fungi as experimental systems for applied as well as basic research that has prompted publication of this series of books under the title The Mycota. This title knowingly relegates fungi into a separate realm, distinct from that of either plants, animals, or protozoa. For consistency throughout this Series of Volumes the names adopted for major groups of fungi (representative genera in parentheses) areas follows: Pseudomycota Division: Division:

Oomycota (Achlya, Phytophthora, Pythium) Hyphochytriomycota

Eumycota Division: Division:

Chytridiomycota (Allomyces) Zygomycota (Mucor, Phycomyces, Blakeslea)

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Division: Subdivision: Class: Class: Subdivision: Class: Class:

Series Preface

Dikaryomycota Ascomycotina Saccharomycetes (Saccharomyces, Schizosaccharomyces) Ascomycetes (Neurospora, Podospora, Aspergillus) Basidiomycotina Heterobasidiomycetes (Ustilago, Tremella) Homobasidiomycetes (Schizophyllum, Coprinus)

We have made the decision to exclude from The Mycota the slime molds which, although they have traditional and strong ties to mycology, truly represent nonfungal forms insofar as they ingest nutrients by phagocytosis, lack a cell wall during the assimilative phase, and clearly show affinities with certain protozoan taxa. The Series throughout will address three basic questions: what are the fungi, what dothey do, and what is their relevance to human affairs? Such a focused and comprehensive treatment of the fungi is long overdue in the opinion of the editors. A volume devoted to systematics would ordinarily have been the first to appear in this Series. However, the scope of such a volume, coupled with the need to give serious and sustained consideration to any reclassification of major fungal groups, has delayed early publication. We wish, however, to provide a preamble on the nature off ungi, to acquaint readers who are unfamiliar with fungi with certain characteristics that are representative of these organisms and which make them attractive subjects for experimentation. The fungi represent a heterogeneous assemblage of eukaryotic microorganisms. Fungal metabolism is characteristically heterotrophic or assimilative for organic carbon and some nonelemental source of nitrogen. Fungal cells characteristically imbibe or absorb, rather thaningest, nutrients and they have rigid cellwalls. The vast majority of fungi are haploid organisms reproducing either sexually or asexually through spores. The spore forms and details on their method of production have been used to delineate most fungal taxa. Although there is amultitude of spore forms, fungal spores are basically only of two types: (i) asexual spores are formed following mitosis (mitospores) and culminate vegetative growth, and (ii) sexual spores are formed following meiosis (meiospores) and are borne in or upon specialized generative structures, the latter frequently clustered in a fruit body. The vegetative forms of fungi are either unicellular, yeasts are an example, or hyphal; the latter may be branched to form an extensive mycelium. Regardless of these details, it is the accessibility of spores, especially the direct recovery of meiospores coupled with extended vegetative haploidy, that have made fungi especially attractive as objects for experimental research. The ability of fungi, especially the saprobic fungi, to absorb and grow on rather simple and defined substrates and to convert these substances, not only into essential metabolites but into important secondary metabolites, is also noteworthy. The metabolic capacities of fungi have attracted much interest in natural products chemistry and in the production of antibiotics and other bioactive compounds. Fungi, especially yeasts, are important in fermentation processes. Other fungi are important in the production of enzymes, citric acid and other organic compounds as well as in the fermentation of foods. Fungi have invaded every conceivable ecological niche. Saprobic forms abound, especially in the decay of organic debris. Pathogenic forms exist with both plant and animal hosts. Fungi even grow on other fungi. They are found in aquatic as well as soil environments, and their spores may pollute the air. Some are edible; others are

Series Preface

ix

poisonous. Many are variously associated with plants as copartners in the formation of lichens and mycorrhizae, as symbiotic endophytes or as overt pathogens. Association with animal systems varies; examples include the predaceous fungi that trap nematodes, the micro fungi that grow in the anaerobic environment of the rumen, the many insect associated fungi and the medically important pathogens afflicting humans. Yes, fungi are ubiquitous and important. There are many fungi, conservative estimates are in the order of 100,000 species, and there are many ways to study them, from descriptive accounts of organisms found in nature to laboratory experimentation at the cellular and molecular level. All such studies expand our knowledge of fungi and of fungal processes and improve our ability to utilize and to control fungi for the benefit of humankind. We have invited leading research specialists in the field of mycology to contributeto this Series. We are especially indebted and grateful for the initiative and leadership shown by the Volume Editors in selecting topics and assembling the experts. We have all been a bit ambitious in producing these Volumes on a timely basis and there in lies the possibility of mistakes and oversights in this first edition. We encourage the readership to draw our attention to any error, omission or inconsistency in this Series in order that improvements can be made in any subsequent edition. Finally, we wish to acknowledge the willingness of Springer-Verlag to host this project, which is envisioned to require more than 5 years of effort and the publication of at least nine Volumes. Bochum, Germany Auburn, AL, USA April 1994

KARL ESSER PAUL A. LEMKE Series Editors

.

Addendum to the Series Preface

During the Fourth International Mycological Congress in Regensburg (1989) while relaxing in a beer garden with Paul Lemke (USA), Dr. Czeschlik (Springer-Verlag) discussed with us the possibility to publish a series about Fungi. We both were at first somewhat reserved, but after a comprehensive discussion this idea looked promising. We decided to name this new series The Mycota. Then Paul Lemke and I created a program involving seven volumes covering a wide area of Mycology. The first volume was presented in 1994 at the Fifth International Mycological Congress in Vancover (Canada). The other volumes followed step by step. After the early death of Paul Lemke (1995) I proceeded alone as Series Editor. However for Vols. X-XII I received support by Joan Bennett. Since evidently the series was well accepted by the scientific community and since the broad area of Fungi was not completely covered, it was decided to proceed with eight more volumes. In addition, second editions of eight volumes were published and three more are in preparation. I would like to thank Springer-Verlag, represented by Drs. Czeschlik and Schlitzberger for their support and cooperation. Bochum, Germany March 2011

KARL ESSER

.

Volume Preface

Based on sequence analyses of many different genes, the past decade has provided us with a profound knowledge of fungal systematics and phylogeny. In addition, several sequences of complete fungal genomes have been revealed; and several others will soon be finished. Now it is feasible to address questions concerning the origin of the fungal kingdom and fungal evolution at a level of analytical refinement that have never been possible before. The Mycota XIV provides a selection of state of the art reviews dealing with major aspects of fungal evolution. The four sections comprise Evolutionary roots of fungi (Chapters 1–3), Evolution of signaling in fungi and fungal-like organisms (Chapters 4–6), Evolution of mutualistic systems and metabolism in fungi (Chapters 7–11) and Evolutionary mechanisms and trends (Chapters 12–13). Fungi are among the oldest eukaryotic organisms in the living world and thus efforts are being made towards understanding their history and importance on the way towards the origin of their more recent sister group, the Metazoa (Chapter 1). Chapter 2 describes the basal fungal lineage Microsporidia, a group of unusual intracellular parasites that infect a wide range of animal cells. This chapter also discusses evidence that links Microsporidia to fungi and the evolutionary processes that have driven the transformation of these highly specialized cells from a free-living eukaryotic lineage. The diversity of the Fungi has been estimated to be as much as 5.1 million species; however only a small fraction of this number is currently described. The uncultered majority is overlooked. Chapter 3 reviews the progress made in identifying fungal diversity using direct molecular approaches from a range of different environments and discusses what these data mean for the fungal tree of life. Dictyostelids were once grouped with fungi because their fruiting structures are superficially similar. However, both protists evolved quite distinct strategies to reach this mode of species propagation. Chapter 4 describes the cell communication systems that cause dictyostelid amoebas to aggregate and that regulate cell type specialization and cell movement during fruiting body formation. In fungi, mating usually occurs between morphologically identical partners that are distinguished only by their mating type. Recognition of a compatible mating partner is accomplished by a pheromone/receptor system. Chapters 5 and 6 describe the current knowledge of the structure, evolution and function of pheromone/receptor systems in filamentous ascomycetes and basidiomycetes. Members of the phylum Glomeromycota, an evolutionary lineage considerably older than the Asco- and Basidiomycota, form symbioses with photoautotrophs and are thought to be among the oldest living asexual eukaryotes. They influence most terrestrial ecosystems by forming arbuscular mycorrhiza, an intimate, mutualistic

xiv

Volume Preface

symbiosis. Chapter 7 highlights evolutionary and co-evolutionary aspects of the Glomeromycota and gives an overview of the current approaches characterizing, describing and detecting arbuscular mycorrhiza fungi as foundations for future understanding their biogeography and related evolutionary aspects. Fruiting body evolution follows a complex pattern, which we are only beginning to understand. Chapter 8 highlights the phylogenetic distribution of ascomatal traits (ascoma type, ascoma development, ascus structure) across the currently accepted classes in the Ascomycota. A particular focus of this chapter is the intertwined evolutionary history of lichenized and non-lichenized species in the Ascomycota. The class Dothideomycetes is a recently defined taxon within the phylum Ascomycota, with approximately 20,000 member species. Many species are important phytopathogens employing diverse pathogenicity strategies. Chapter 9 compares the currently available genome sequences, including their mitochondrial sequences to describe the remarkable plasticity of Dothideomycetes, characterized by extensive rearrangement of gene order and orientation. Recent advances in genome sequencing indicate that the number of genes potentially involved in the biosynthesis of secondary metabolites is much larger than the number of known metabolites. These findings raise the questions of how and why fungi evolved so many genes for the production of secondary metabolites. Chapter 10 gives an overview of secondary metabolite genes in different fungal groups, addresses some of the principles of the regulation of expression of these genes and summarizes what is known about the biological functions of secondary metabolites in fungi. Chapter 11 offers a survey of the gene family of carbonic anhydrases, enzymes that are capable to rapidly accelerate the spontaneous and reversible interconversion from carbon dioxide to bicarbonate. To date, fungal carbonic anhydrases have been identified in the genomes of ‘basal Fungi’, ascomycetes, basidiomycetes and also within the group of non-fungal organisms. Duplication events, which have obviously occurred at several times, seem to be the major force in the evolutionary history of this gene family and might reflect numerous specialization events. Sexual reproduction in fungi is regulated by relatively small genomic regions containing the mating-type loci. Chapter 12 describes the genomic traits and evolutionary features of the mating-type loci and the mating-type chromosomes in model systems of filamentous ascomycetes. The main focus of this chapter lies in recent scientific advances from studies in Neurospora. Metabolism is one of the key characteristics of life, implying that evolution always had an impact on metabolic processes. Studies on fungal metabolism have traditionally concentrated on metabolites of specific interest, namely mycotoxins, pathogenicity factors, antibiotics and other compounds displaying interspecific activity. Chapter 13 offers an overview on several theories on metabolic evolution in fungi. We are very grateful to the authors for their contribution to The Mycota XIV. All are extremely busy and we appreciate their willingness to commit valuable time to this project. We hope that readers enjoy reading this volume of The Mycota. We thank Dr. Andrea Schlitzberger of the Springer Verlag for her support during the preparation of this volume.

Go¨ttingen and Jena, Germany March 2011

STEFANIE PO¨GGELER JOHANNES WO¨STEMEYER Volume Editors

Contents

Evolutionary Roots of Fungi 1 The Protistan Origins of Animals and Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 MARTIN CARR, SANDRA L. BALDAUF 2 Microsporidia – Highly Reduced and Derived Relatives of Fungi . . . . . . . . . . . . . . . . 25 BRYONY A.P. WILLIAMS, PATRICK J. KEELING 3 Environmental DNA Analysis and the Expansion of the Fungal Tree of Life . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 37 MEREDITH D.M. JONES, THOMAS A. RICHARDS Evolution of Signalling in Fungi and Fungal-Like Organisms 4 Evolution of Signalling and Morphogenesis in the Dictyostelids . . . . . . . . . . . . . . . . . 57 PAULINE SCHAAP 5 Function and Evolution of Pheromones and Pheromone Receptors in Filamentous Ascomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 73 STEFANIE PO¨GGELER 6 Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 97 URSULA KUËS, TIMOTHY Y. JAMES, JOSEPH HEITMAN Evolution of Mutualistic Systems and Metabolism in Fungi 7 Evolution of the ‘Plant-Symbiotic’ Fungal Phylum, Glomeromycota . . . . . . . . . . . 163 ARTHUR SCHU¨ßLER, CHRISTOPHER WALKER 8 Fruiting Body Evolution in the Ascomycota: a Molecular Perspective Integrating Lichenized and Non-Lichenized Groups . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 187 IMKE SCHMITT 9 Genomic and Comparative Analysis of the Class Dothideomycetes . . . . . . . . . . . . 205 JAMES K. HANE, ANGELA H. WILLIAMS, RICHARD P. OLIVER 10 Evolution of Genes for Secondary Metabolism in Fungi . . . . . . . . . . . . . . . . . . . . . . . 231 INES TEICHERT, MINOU NOWROUSIAN

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Contents

11 Carbonic Anhydrases in Fungi and Fungal-Like Organisms – Functional Distribution and Evolution of a Gene Family . . . . . . . . . . . . . . . . . . . . . . . 257 SKANDER ELLEUCHE Evolutionary Mechanisms and Trends 12 Evolution of Mating-Type Loci and Mating-Type Chromosomes in Model Species of Filamentous Ascomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 277 CARRIE A. WHITTLE, HANNA JOHANNESSON 13 Evolution of Special Metabolism in Fungi: Concepts, Mechanisms, and Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 293 CHRISTINE SCHIMEK Biosystematic Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 331 Subject Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 337

List of Contributors

SANDRA L. BALDAUF (e-mail: [email protected]) Department of Systematic Biology, Evolutionary Biology Centre, University of Uppsala, Norbyva¨gen 18D, 752-36 Uppsala, Sweden MARTIN CARR (e-mail: [email protected], +113 3432596, Fax: +113 3432835) LC Miall Building, Faculty of Biological Sciences, University of Leeds, Clarendon Way, Leeds, LS2 9JT, UK SKANDER ELLEUCHE (e-mail: [email protected], +49 40 42878 3334, Fax: +49 40 42878 2582) Technische Universita¨t Hamburg-Harburg, Institut fu¨r Technische Mikrobiologie, Kasernenstrasse 12, 21073 Hamburg, Germany JAMES K. HANE CSIRO Plant Industry, CELS Floreat, Perth, WA 6014, Australia JOSEPH HEITMAN (e-mail: [email protected]) Department of Molecular Genetics and Microbiology, Duke University Medical Center, 322 CARL Building, Research Drive, Durham, NC 27710, USA TIMOTHY Y. JAMES (e-mail: [email protected]) Department of Ecology and Evolutionary Biology, University of Michigan, 830 N. University, Ann Arbor, MI 48109, USA HANNA JOHANNESSON (e-mail: [email protected]) Department of Evolutionary Biology, Uppsala University, Uppsala, Sweden MEREDITH D. M. JONES (e-mail: [email protected], +44 207 942 6320, Fax: +44 207 942 5054) Natural History Museum, Cromwell Road, London, SW7 5BD, UK PATRICK J. KEELING (e-mail: [email protected], +1 604 822 4906, Fax: +1 604 822 6089) Canadian Institute for Advanced Research, Botany Department, University of British Columbia, 3529–6270 University Boulevard, Vancouver, British Colombia, V6T 1Z4, Canada

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URSULA KUËS (e-mail: [email protected], +49 551 392 2705) Division of Molecular Wood Biotechnology and Technical Mycology, University of Go¨ttingen, Bu¨sgen-Institute, Bu¨sgenweg 2, 37077 Go¨ttingen, Germany MINOU NOWROUSIAN (e-mail: [email protected], +49 234 322 4588, Fax: +49 234 321 4184) Lehrstuhl fu¨r Allgemeine und Molekulare Botanik, Ruhr-Universita¨t Bochum ND 6/165, Universita¨tsstrasse 150, 44780 Bochum, Germany RICHARD P. OLIVER (e-mail: [email protected], +618 9266 7872, Fax: +618 9266 2021) Department of Environment and Agriculture, Curtin University, Perth, WA 6102, Australia STEFANIE PO¨GGELER (e-mail: [email protected], +49 551 391 3930, Fax: +49 551 391 0123) Genetics of Eukaryotic Microorganisms, Institute of Microbiology and Genetics, Georg-August University Go¨ttingen, Grisebachstrasse 8, 37077 Go¨ttingen, Germany THOMAS A. RICHARDS (e-mail: [email protected]) Natural History Museum, Cromwell Road, London, SW7 5BD, UK PAULINE SCHAAP (e-mail: [email protected], +44 1382 388078, Fax: +44 1382 345386) College of Life Sciences, University of Dundee, MSI/WTB/JBC Complex, Dow Street, Dundee DD1 5EH, UK CHRISTINE SCHIMEK (e-mail: [email protected], +49 3641 949327, Fax: +49 3641 949312) Department of General Microbiology and Microbe Genetics, Institute of Microbiology, Friedrich-Schiller-Universita¨t Jena, Neugasse 24, 07743 Jena, Germany IMKE SCHMITT (e-mail: [email protected], +49 69 7542 1855, Fax: +49 69 7542 1800) Goethe University Frankfurt and Biodiversity and Climate Research Centre, Senckenberganlage 25, 60325 Frankfurt am Main, Germany ARTHUR SCHU¨ßLER (e-mail: [email protected], +49 89 2180 74730, Fax: +49 89 2180 74702) Genetics, Department of Biology, University of Munich (LMU), Grosshaderner Strasse 2–4, 82152 Martinsried, Germany INES TEICHERT Lehrstuhl fu¨r Allgemeine und Molekulare Botanik, Ruhr-Universita¨t Bochum ND 6/165, Universita¨tsstrasse 150, 44780 Bochum, Germany CHRISTOPHER WALKER Royal Botanic Garden Edinburgh, 20A Inverleith Row, Edinburgh, EH3 5LR, UK; and School of Earth Sciences and Environment, University of Western Australia, 35 Stirling Highway, Crawley, WA 6009, Australia


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CARRIE A. WHITTLE Department of Evolutionary Biology, Uppsala University, Uppsala, Sweden ANGELA H. WILLIAMS Faculty of Health Sciences, Murdoch University, Perth, WA 6150, Australia BRYONY A. P. WILLIAMS (e-mail: [email protected]) School of Biosciences, Geoffrey Pope Building, University of Exeter, Stocker Road, Exeter, EX4 4QD, UK

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Evolutionary Roots of Fungi

.

1

The Protistan Origins of Animals and Fungi

MARTIN CARR1, SANDRA L. BALDAUF2

CONTENTS I. II. III. IV. V. VI. VII.

VIII.

IX.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Opisthokonta . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nucleariida. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Metazoa . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Radiata . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Bilateria. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ichthyosporea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Rhinosporideacae . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Ichthyophonae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Choanoflagellida. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Codosigid Morphology . . . . . . . . . . . . . . . . . . . . . . B. Salpingoecid Morphology. . . . . . . . . . . . . . . . . . . . C. Acanthoecidae. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Incertae Sedis Opisthokont Protists. . . . . . . . . . . . A. Aphelidea. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Capsaspora . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Ministeria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Corallochytrea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Phylogeny of Opisthokonta . . . . . . . . . . . . . . . . . B. Sister-Group to Opisthokonta . . . . . . . . . . . . . . . C. Determining Opisthokont Diversity. . . . . . . . . D. Reconstructing Opisthokont Evolution and the Origins of Multicellularity . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

3 5 5 7 8 9 9 10 10 11 12 14 14 14 15 15 15 16 16 17 17 17 17 17 18

I. Introduction Fungi and Metazoa constitute two of the major multicellular eukaryotic lineages and a large body of robust data confirms that they are close relatives (Baldauf and Palmer 1993; Burki et al. 2007; Hackett et al. 2007; Hampl et al. 2009; Wainright et al. 1993). Along with three major groups of protists – nucleariids, ichthyosporeans and choanoflagellates – and four enigmatic incertae sedis 1

LC Miall Building, Faculty of Biological Sciences, University of Leeds, Clarendon Way, Leeds, LS2 9JT, UK; e-mail: M.Carr@leeds. ac.uk 2 Department of Systematic Biology, Evolutionary Biology Centre, University of Uppsala, Norbyva¨gen 18D, Uppsala 75236, Sweden; e-mail: [email protected]

groups – Aphelidea, Capsaspora, Corallochytrea and Ministeria – Fungi and Metazoa make up the phylogenetic supergrouping Opisthokonta (Adl et al. 2005; Cavalier-Smith 1987). The ichthyosporeans, nucleariids and incertae sedis groups were placed together in the taxon Mesomycetozoa by Adl et al. (2005). Whilst the Opisthokonta is a well established grouping, Mesomycetozoa remains controversial as it is paraphyletic (Fig. 1.1) and not universally recognised. Therefore each of the mesomycetozoan lineages will be discussed separately. The relationships among the opisthokont groups, and those of the opisthokonts with other eukaryotic supergroups, are slowly becoming clearer (Fig. 1.1). We discuss here the major taxonomic groups within Opisthokonta and their relationships with each other. It now seems clear that the deepest bifurcation within the opisthokonts resulted in two major lineages, now referred to as Holozoa and Holomycota. The Holomycota is composed of the Fungi and nucleariids (Lara et al. 2010). Metazoa, the choanoflagellates, ichthyosporeans and the four incertae sedis protists are collectively known as Holozoa (Lang et al. 2002; Ruiz-Trillo et al. 2004; Steenkamp et al. 2006). Taxon-rich phylogenies of the opisthokonts are frequently ribosomal DNA (rDNA) studies. This is because these genes are multi-copy in eukaryotes and possess highly conserved regions that facilitate the binding of universal PCR primers – two factors which allow relatively rapid and easy DNA sequencing. The same, or similar, protocols can therefore be employed to produce orthologous sequences across a broad variety of taxa.

Whilst there are many advantages in producing large-scale phylogenies based on rDNA sequences, it is often necessary to use sequences from multiple genes when attempting to resolve deep branches. Combining small- and largesubunit rDNA sequences (Medina et al. 2003; Moreira et al. 2007) is one simple approach to Evolution of Fungi and Fungal-Like Organisms, The Mycota XIV S. Po¨ggeler and J. Wo¨stemeyer (Eds.) © Springer-Verlag Berlin Heidelberg 2011

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Martin Carr and Sandra L. Baldauf

Fig. 1.1. Simplified four-gene phylogeny of the opisthokonts. The tree shown was derived by Bayesian inference, using MrBayes 3.1.1 (Ronquist and Huelsenbeck 2003), based on a combined rDNA (SSU, LSU), a-tubulin (tubA), 90-kDa heat-shock protein (hsp90) dataset. The 6415 aligned nucleotide positions were analysed using the GTR+I+G substitution model, with separate partitions for rRNA, first + second codon positions and third positions. Support values are also shown from a maximum

likelihood tree derived using RAxML 7.0.3 (Stamatakis et al. 2005) utilizing the GTRCAT substitution model. Bayesian inference posterior probabilities (biPP) and maximum likelihood bootstrap percentages (mlBP) values are given above and below branches respectively. Branches are drawn proportional to the number of nucleotide substitutions per site as indicated by the scale bar at the lower left. Dotted lines indicate taxa of uncertain phylogenetic position (modified from Carr et al. 2008a)

creating phylogenies from more than one gene. However, the two genes are together in the genome, transcribed as a single unit (Perry et al. 1970) and function in the same biosynthetic pathway. It is therefore likely that both subunits will be affected by similar evolutionary forces (natural selection, local mutation bias, etc.) and may, in effect, act as a single locus. A preferential method for producing robust opisthokont phylogenies is to use multiple ribosomal and protein-coding genes in conjunction; such sequences can be produced using targeted PCR with degenerate primers (Carr et al. 2008a; Ruiz-Trillo et al. 2004; Steenkamp et al. 2006) or expressed sequence tag (EST) libraries (Patron et al. 2007; Philippe et al. 2004; Shalchian-Tabrizi et al. 2008). The sequencing of whole genomes and EST libraries allows the construction of datasets composed of many thousands of aligned amino acid positions. This field, termed phylogenomics, allows the powerful analyses of deep phylogenetic relationships (Burki et al. 2008; Minge et al. 2009; Philippe et al. 2009). At present, the sequencing of either whole

genomes or EST libraries is still a relatively expensive procedure and as a result genomic data are publicly available for less than ten opisthokont protists. Sparse taxon sampling is a difficult issue in producing accurate phylogenies. Whilst a wealth of molecular data exists for many of the major lineages of Fungi and Metazoa, there are data for far fewer opisthokont protists. Nonetheless multiple protein-coding gene sequences are available for 15 species of choanoflagellates, two species of ichthyosporeans, a single nucleariid and three of the incertae sedis holozoan taxa (Capsaspora owczarzaki, Corallochytrium limacisporum, Ministeria vibrans). Poorly sampled taxa can lead to species being present on isolated long branches; this, in turn, may lead to problems when reconstructing phylogenies due to the phenomenon of long-branch attraction (Hendy and Penny 1989). When distantly related sequences share a relatively high number of characters due to convergence rather than ancestry the true phylogenetic signal may


be overwhelmed. Long-branch effects can also be produced by unequal rates of evolution; therefore, when possible, it is advisable to screen taxa and pick those most suitable for phylogenetic reconstruction. This however is not always possible, particularly in the case of the opisthokont protists where some lineages are only represented by a single known species.

II. Opisthokonta The Opisthokonta was originally postulated by Cavalier-Smith, partly based on the presence of a posteriorly directed flagellum common to both Fungi and Metazoa. However as the depth of diversity in opisthokont lineages has been uncovered, it has become apparent that there are no recognised universal morphological characters unique to this group. The posterior flagellum is not present in all opisthokont groups; loss of the flagellum must have occurred on multiple occasions within Fungi (James et al. 2006a), as well within the nucleariids, ichthyosporeans, Capsaspora, Corallochytrea and possibly ministeriids. Throughout the opisthokonts, the morphology of mitochondrial cristae is predominantly flat (Adl et al. 2005), but this appears to be a plastic trait, with lamellar, tubular and discoidal cristae also present (Amaral-Zettler et al. 2001; Ragan et al. 1998). Nonetheless, these characteristics are widespread across the opisthokonts and may point to the ancestral state of the group. The abilities to produce amoeboid cells and also to engulf particles by phagocytosis are present in all of the major lineages. Moreover, Metazoa is the only major opisthokont lineage that does not contain species with cell walls and it has been suggested that the last common ancestor of the opisthokonts also possessed the potential to produce a cell wall (Mendoza et al. 2002). Nearly all described eukaryotes can be assigned to one of six super-groupings (for a review, see Adl et al. 2005), namely Opisthokonta, Amoebozoa, Archaeplastida, Chromalveolata, Excavata and Rhizaria. Of these, Amoebozoa appear to form a super-assemblage with the opisthokonts (Burki et al. 2007; Stechmann and Cavalier-Smith 2003). Opisthokonta and Amoebozoa both contain species possessing a single flagellum, giving rise

5

to the name Unikonta for the super-assemblage (Stechmann and Cavalier-Smith 2003). Under this hypothesis, the other eukaryotic groups, termed bikonts, may have ancestrally possessed two flagella (Cavalier-Smith 2009). The enigmatic Apusozoa are also proposed to be the closest group to the opisthokonts (Kim et al. 2006), however if the biflagellate Apusozoa are the true sister-group to the opisthokonts, then the unikonts must be a polyphyletic grouping. Competing hypotheses however suggest that the apusozoans may alternatively be an early branching lineage within either the amoebozoans or bikonts (Minge et al. 2009).

The uncertainty over the sister-group of the opisthokonts may only be resolved by increasing the number of apusozoan and amoebozoan taxa for which multiple gene sequences are available.

III. Fungi Fungi are a large, diverse group of heterotrophs, which predominantly employ an absorptive mode of nutrition. Species often produce multinucleate hyphae and cell walls that comprise both b-glucan and chitin (Cavalier-Smith 1998, 2001; Kirk et al. 2001; Tehler 1988). When present, mitochondrial cristae are flat. Fungal classification and phylogeny is currently in a state of flux, with many traditional taxa (e.g., Zygomycota) now considered redundant. At its simplest level, Fungi can be divided into the subkingdom Dikarya (formerly Neomycota; Cavalier-Smith 1998) and nine basal groups (Blastocladiomycota, Chytridiomycota, Entomophthoromycotina, Glomeromycota, Kickxellomycotina, Microsporidia, Mucoromycotina, Neocallimastigomycota, Zoopagomycotina; Hibbett et al. 2007). Dikarya is characterised by species possessing pairs of unfused haploid nuclei (dikaryons). The taxon is composed of the phyla Ascomycota and Basidiomycota and comprises the majority (~98%) of described fungal species (James et al. 2006a). The two phyla form a well supported monophyletic group within Fungi (Fig. 1.2; James et al. 2006a; Liu et al. 2006). In contrast to Dikarya, the relationships between the basal lineages of Fungi are not, at present, well resolved. Several genera (e.g., Basidiobolus, Olpidium, Rozella) are not currently associated

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Fig. 1.2. Simplified four-gene phylogeny of the Fungi and Nucleariida. The tree was derived by a maximum likelihood method, using RAxML 7.0.3, from partial sequences of rRNA (SSU and LSU) tubA and hsp90. The 6411 aligned nucleotide positions were analysed using the GTRCAT substitution model, with separate partitions for rRNA, first + second codon positions and third positions. Support values are also shown from a maximum likelihood tree derived using MrBayes 3.1.1 utilizing the GTR+I+G

substitution model; mlBP and biPP values are given above and below branches respectively. The long-branch and phylogenetically unstable subphyla Entomophthoromycotina, Kickxellomycotina and Zoopagomycotina have been omitted. Branches are drawn proportional to the number of nucleotide substitutions per site as indicated by the scale bar at the lower left. Dotted lines indicate taxa of uncertain phylogenetic position

with any higher taxa (Hibbett et al. 2007). Chytridiomycota was long considered the most basal lineage within Fungi, with the Zygomycota viewed as the sister-group to Dikarya. Molecular phylogenies have confirmed that the former chytridiomycetes do indeed fall at the base of the fungal tree (Fig. 1.2; James et al. 2006a; Tehler et al. 2000), but show that the traditional taxon is polyphyletic comprising at least four major lineages: Rozella spp., the “core chytrids”, Blastocladiomycota and Olpidium spp. (Fig. 1.2; James et al. 2006a, b). Of these, it appears that Rozella forms the earliest branching clade within Fungi (Lara et al. 2010). In fact, recent evidence indicate that it is a member of a large clade, the “cryptomycota”, which is known almost exclusively from metagenomic data (Jones et al. 2011). The former Chytridiomycota is now formally separated into the Chytridiomycota, Neocallimastigomycota and Blastocladiomycota (Hibbett et al. 2007; James et al. 2006b). Zygomycota also appears to be a polyphyletic assemblage (Fig. 1.2; James et al. 2006a; Keeling 2003; Voigt and Wo¨stemeyer 2001). A recent

revision of fungal classification considered the Zygomycota to be a redundant taxon (Hibbett et al. 2007) and divided the former group into one phylum (Glomeromycota) and four subphyla incertae sedis (Entomophthoromycotina, Mucormycotina, Kickxellomycotina and Zoopagomycotina). Within the former zygomycete groups, the glomeromycetes have been tentatively placed as the closest lineage to Dikarya (James et al. 2006a; Voigt and Wo¨stemeyer 2001). This result however may be heavily biased by a reliance on rDNA sequences, since phylogenies based only on protein-coding genes place the Mucormycotina as the sister-group to Dikarya (Lee and Young 2009; Liu et al. 2009). The position of Microsporidia within Fungi remains unclear (see Chapter 2 in this volume). Liu et al. (2006) considered Microsporidia as a sister-group to all other fungi; however, using a larger dataset, James et al. (2006a) placed these highly reduced obligate parasites as an early branching lineage and the sister-group to Rozella spp (see Chapter 3 in this volume). A more derived


position, within the former zygomycetes, was proposed on the basis of protein-coding genes (Gill and Fast 2006; Keeling 2003) and genome-wide conserved gene order (synteny; Lee et al. 2008). A robust phylogeny of Fungi is vital in order to understand how the group evolved. Due to the widespread presence of a posteriorly positioned flagellum in the basal fungal lineages (James et al. 2006a), parsimony argues that it was present in the last common ancestor of Fungi and that flagella loss has subsequently occurred. However, due to competing phylogenies the number of losses that have occurred is uncertain. Liu et al. (2006) presented a phylogeny that only required a single loss of the flagellum, whereas other phylogenies (James et al. 2006a; Tanabe et al. 2005) highlight multiple flagella losses. Although considered one of the major multicellular kingdoms, unicellular species are found across the fungal tree. Only unicellular forms are know for the early branching Microsporidia and Rozella. In addition, secondary reversions to unicellularity (exemplified by the ascomycete yeasts) appear to have evolved on multiple occasions within the kingdom (James et al. 2006a). Only a minority of described fungal species are marine and it is unclear from phylogenetic reconstructions whether the last common ancestor of Fungi was a marine or freshwater organism (James et al. 2006a). It does appear however that the radiation of the major extant fungal lineages occurred in the terrestrial environment and, in fact, it has been proposed that the presence of glomeromycetes on land was essential for terrestrial colonisation by green plants (James et al. 2006a; Pirozynski and Malloch 1975). Despite the recent, and continuing, advances using multigene phylogenies, a complete robust tree of Fungi remains elusive. At the time of writing, the Deep Hypha initiative (http://ocid. nacse.org/research/deephyphae/) is in the process of producing a large-scale phylogeny by sequencing seven loci from 300 basal fungal taxa and 1200 Dikarya species.

IV. Nucleariida Nucleariids are a small group of mainly free-living amoeboid protists. The protoplast (cell body) appears as a spherical or flattened elongated amoeba with radiating filopodia (Amaral-Zettler

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et al. 2001). Flagellated cells have not been observed in any of the stages in the life cycles of any nucleariid. All described species thus far have been isolated from freshwater (Maldonado 2004). Most described species are algivorous or bacterivorous (Patterson 1984). One potential exception to this is Nuclearia pattersoni, which was discovered living in the gills of freshwater fish (Dykov et al. 2003). However the authors did not ascertain whether the association between the amoebae and host is parasitic or commensal. There is considerable variation within the ultrastructure of species in the group (for a review, see Yoshida et al. 2009). Two types of mitochondrial cristae have been reported; most species have discoidal cristae (Amaral-Zettler et al. 2001) but Nuclearia pattersoni exhibits flattened cristae (Dykov et al. 2003). The majority of known species have small (15–25 mm in diameter), uninucleate cells; in contrast Nuclearia delicatula has larger (50 mm) cells with 2–12 nuclei (Yoshida et al. 2009). Cyst formation has been observed in Nuclearia pattersoni and Nuclearia simplex, but both Nuclearia delicatula and Nuclearia moebiusi appear incapable of producing cysts. Additionally, a mucoidal extracellular matrix, absent from Nuclearia moebiusi, is present in Nuclearia delicatula, Nuclearia pattersoni and Nuclearia simplex (Dykov et al. 2003; Patterson 1984).

The taxonomic classification of the nucleariids has been reappraised over recent years through the use of molecular phylogenetics. Due to the morphology of their mitochondrial cristae, they were initially placed within the Filosea, a subgroup of Amoebozoa (Cavalier-Smith 1993; Page 1987). The monophyly of Filosea was questioned by Patterson (1999) on the basis that filopodia are likely to have evolved on multiple occasions within eukaryotes, as well as the unexpected presence of three mitochondrial cristal morphologies in the group. Amaral-Zettler et al. (2001) confirmed the polyphyly of Filosea using small-subunit rDNA sequences, which strongly supported the inclusion of nucleariids within Opisthokonta and not Amoebozoa. This study also highlighted polyphyly within the sequenced nucleariids, showing that one species (Nuclearia sp. ATCC 30864) did not group with the other taxa. This species was later reclassified as Capsaspora owczarzaki and moved to Mesomycetozoa (Hertel et al. 2002; see below). Early small-subunit rDNA studies did not robustly resolve the position of the nucleariids within the opisthokonts

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(Amaral-Zettler et al. 2001; Cavalier-Smith and Chao 2003; Hertel et al. 2002). Subsequently, multiple-gene phylogenies firmly placed nucleariids as the sister-group to Fungi (Moreira et al. 2007; Ruiz-Trillo et al. 2004; Steenkamp et al. 2006).

generated from this genome sequence may be a useful aid to understanding the early evolution of Fungi.

A recent five-gene study by Brown et al. (2009) identified the slime mold Fonticula alba as a member of the Holomycota. Their phylogenies grouped F. alba together with the nucleariids with strong support. Brown et al. (2009) termed this new group the nuclearioid amoebae and their phylogenies placed them as the sister-group of Fungi. F. alba is an amoeboid protist, which undergoes aggregation to form a conical multicellular fruiting body. Cells at the tip of the fruiting body encyst and form spores, creating a prominent apical bulge. Mature spores are then forcibly ejected from the bulge and germinate to produce amoeboid cells (Deasey 1982; Worley et al. 1979). If the placement of F. alba is confirmed through additional studies, multicellularity appears to have evolved on at least two occasions within Holofungi. The aggregative form of multicellularity employed by F. alba is distinct from multicellularity in Fungi and it will be of significant interest to compare the relevant genetic pathways within the two lineages.

V. Metazoa

The placement of the nucleariids at the base of Fungi highlights the considerable morphological and ecological evolution that has occurred since they last shared a common ancestor. Flagellated cells are widespread in basal Fungi (James et al. 2006a), therefore flagellar loss in the nucleariids was clearly independent from loss in Fungi. Amoebal motility however has been observed in Fungi (Loytynoja and Milikovitch 2001; Mendoza et al. 2002) so it is quite likely that the common ancestor of the two lineages could produce amoeboid cells. Two feeding modes have been documented in the nucleariids. In Nuclearia delicatula filopodia have been shown to penetrate damaged algal cells (Cann 1986). However, most species employ phagocytosis, as does the parasitic basal fungal species Rozella polyphagi which phagocytoses the organelles of its host (Powell 1984). This suggests that the last common ancestor of nucleariids and Fungi may have fed by means of phagotrophy (Lara et al. 2010). Both lineages comprise predominantly freshwater species; therefore it is quite possible that the last common ancestor of both groups was a freshwater or terrestrial organism (Cavalier-Smith 2009). At the time of writing, the genome of Nuclearia simplex is being sequenced. The data

The Kingdom Metazoa comprises multicellular phagotrophs with fibrous connective tissue (normally collagens) between epithelial layers (CavalierSmith 1998). Mitochondrial cristae are flat or, more rarely, tubular (Adl et al. 2005). Metazoans typically possess greatly reduced mitochondrial genomes (~16 kb), with a highly conserved set of 37 genes (Lang et al. 1999; Dellaporta et al. 2006; Signorovitch et al. 2007). This is in sharp contrast to the known mitochondrial genomes of opisthokont protists. The choanoflagellate Monosiga brevicollis has a 76.6-kb genome that encodes 55 genes and the ichthyosporean Amoebidium parasiticum has a mitochondrial genome of more than 200 kb on hundreds of linear chromosomes (Burger et al. 2003). In addition to the transfer of genes from the mitochondrion to the nucleus, the reduction in mitochondrial genome size in Metazoa is the result of the loss of introns and the reduction of intergenic regions. Within this compact genome as much as 98% of the DNA codes for either proteins or catalytic RNAs (Signorovitch et al. 2007). Interestingly, parallel gene loss and genome reduction have also occurred in Fungi (Burger et al. 2003), but fungal species do not show such uniformly compact genomes (Brown et al. 1985; Bullerwell et al. 2003; Cardoso et al. 2007; Cummings et al. 1990; Woo et al. 2003). The number of recognised metazoan phyla ranges from 21 to 38 (e.g., Cavalier-Smith 1998; Farabee 2002; Hyman 1940; Philippe and Telford 2006) and most phyla can be placed in one of two major subdivisions – Radiata and Bilateria (for a review, see Telford 2006). Radiata (diploblasts) have a body comprising two tissue layers, the endoderm (gut) and ectoderm (skin). Bilateria (triploblasts) have a third layer, the mesoderm, located between the endoderm and ectoderm and a body plan that exhibits bilateral symmetry. The mesoderm may form a coelom (fluid-filled body cavity). Metazoans that possess a coelom do not form a monophyletic group; therefore it would appear that it is either an ancestral character,


which has been lost on multiple occasions within Bilateria, or it has independently evolved on multiple occasions. Recently published evidence for the evolution of the bilaterian gut on multiple occasions has come from the expression patterns of developmental genes (Hejnol and Martindale 2008).

A. Radiata The Radiata consists of three principal phyla: Porifera (sponges), Cnidaria (corals, jellyfish, sea anemones) and Ctenophora (comb jellies). Additionally two further phyla have been placed within Radiata through molecular phylogenies. Placozoa currently contains a single described species, Trichoplax adhaerens, although culture independent rDNA libraries indicate that the phylum possesses significantly more diversity (Voigt et al. 2004). T. adhaerens is morphologically the simplest extant metazoan known with only four somatic cell types. Each individual has an upper and lower layer of epithelia that surround a loose collection of contractile fibre cells and gland cells (Schierwater 2005). Molecular phylogenies support the placement of Placozoa within Radiata, but its precise relationship to the other lineages remains uncertain (Dellaporta et al. 2006; Erpenbeck et al. 2007; Signorovitch et al. 2007; Srivastava et al. 2008; Voigt et al. 2004; Wallberg et al. 2004; Wang and Lavrov 2007).

Myxozoa includes over 2000 species of sporeforming parasites (Jimeńez-Guri et al. 2007; Monteiro et al. 2002). The group was long considered protozoan rather than Metazoa. However molecular phylogenies – using both rDNA and protein-coding genes – have placed the lineage within Metazoa (Anderson et al. 1998; Smothers et al. 1994; Wallberg et al. 2004) as highly derived cnidarians (Jimeńez-Guri et al. 2007). Molecular phylogenies suggest that Radiata is almost certainly a paraphyletic assemblage, but have yet to determine the relationships within the group. Porifera has traditionally been considered as the most basal metazoan lineage on the basis of their simple body plan and most molecular phylogenies confirm this (Baurain et al. 2007; Jimeńez-Guri et al. 2007; Wallberg et al. 2004;). The monophyly of Porifera has been questioned in a number of studies (Borchiellini et al. 2001;

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Haen et al. 2007; Medina et al. 2001; Wallberg et al. 2004), raising the possibility that all extant metazoans may have evolved from a sponge-like ancestor. Ctenophora appears to be the next branching metazoan lineage, although one recent large-scale EST phylogeny by Dunn et al. (2008) placed the ctenophores at the base of the metazoans. The Cnidaria are recovered as the sister-group to Bilateria (Jimeńez-Guri et al. 2007; Medina et al. 2001; Wallberg et al. 2004). Recent work by Matus et al. (2006) showed that cnidarians possess many of the developmental genes previously thought to be unique to Bilateria, leading them to speculate that cnidarians were ancestrally bilaterally symmetrical and have subsequently evolved a more simplified body plan. B. Bilateria The monophyletic lineage Bilateria is further subdivided into the deuterostomes and protostomes. The defining characteristics of these two subdivisions are the fates of the primary embryonic opening (blastopore). In deuterostomes the blastopore develops into the anus (with the mouth developing from a secondary opening), whilst in the protostomes the blastopore typically develops into the mouth (for a review, see Telford 2006). Bourlat et al. (2006) recently revised the make up of Deuterostomia, showing that it consists of four phyla (Chordata, Hemichordata, Echinodermata, Xenoturbellida). Protostomia is made from the subdivisions Ecdysozoa and Lophotrochozoa (Aguinaldo et al. 1997; Anderson et al. 2004; Jimeńez-Guri et al. 2007; Mallatt et al. 2004). Ecdysozoa comprises organisms that undergo ecdysis (the moulting of cuticle: panarthropods, nematodes, nematomorphs, kinorhynchs, loriciferans, priapulids; see Halanych 2004; Philippe and Telford 2006). Lophotrochozoa has no single unifying morphological characteristic, but was proposed by Halanych et al. (1995) on the basis of molecular phylogenetics. Lophotrochozoans (platyhelminthes, phoronids, sipunculids, nemerteans, gastrotrichs, cycliophorans, entoprocts, gnathostomulids, rotifers, acanthocephalans, molluscs, annelids, nemertines, brachiopods, dicyemids, myzostomids) possess either a lophophore (a structure of ciliated tentacles around the mouth) or trochophore larvae (planktonic larvae with a distinctive band of cilia). Metazoan phylogenomics has yet to resolve the relationships within Lophotrochozoa, but indicate that neither the trochozoans nor lophophorates form monophyletic groups (Bourlat et al. 2008; Dunn et al. 2008).

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VI. Ichthyosporea The ichthyosporeans are an ecologically and morphologically diverse group of predominantly parasitic protists. The first indications of this lineage came from Spanggaard et al. (1996) and Ragan et al. (1998), with the latter naming the group the DRIP clade. This was an acronym based on the first known members – Dermocystidium, rosette agent, Ichthyophonus and Psorospermium. Cavalier-Smith (1998) later placed these protists in Ichthyosporea; Herr et al. (1999) however recommended the name Mesomycetozoa to reflect the phylogenetic position of the group, that is, taxa intermediate between Fungi and Metazoa. This recommendation was taken up by Mendoza et al. (2001) and the composition of the group was further amended by Adl et al. (2005) to include several non-ichthyosporeans (e.g., Capsaspora, Corallochytrium, Ministeria, the nucleariids). Acceptance of Mesomycetozoa as a group is not universal, particularly as, under the most recent taxon description, it is paraphyletic (RuizTrillo et al. 20006; Shalchian-Tabrizi et al. 2008; Steenkamp et al. 2006). Membership of Ichthyosporea is based almost exclusively on phylogenetic trees, since there are no known unifying morphological, ultrastructural or ecological characteristics. Many species now recognised as ichthyosporeans were originally classified as fungi, although their placement was usually controversial (Lichtwardt 1986; Trotter and Whisler 1965). For example, Amoebidiales and Eccrinales were previously classified as trichomycetes but are now clearly ichthyosporeans based on rDNA phylogeny (Benny and O’Donnell 2000; Cafaro 2005).

Parasitism is the prevalent strategy among known ichthyosporeans, although symbiosis and saprotrophy are also observed (Mendoza et al. 2002). Protoplasts show a great variety of forms, with flagellated and amoeboid cells widespread and many species having cysts, sporangia or hyphal stages (Mendoza et al. 2002). Mitochondrial cristae are predominantly flat but, unlike its close congeners, Ichthyophonus hoferi is reported to have tubular cristae (Ragan et al. 1998). No ichthyosporeans have been confirmed as planktonic although two species, Sphaeroforma arctica

and LKM51, are thought to have stages in their life cycles which are free-living in the water column (Mendoza et al. 2002; van Hannen et al. 1999). Ichthyosporea has been subdivided into two groups (Fig. 1.3), based on phylogenetic trees and differences in their life cycles (Cafaro 2005; Mendoza et al. 2002).

A. Rhinosporideacae Initially erected as Dermocystida by CavalierSmith (1998), this group mainly comprises parasites of fish, mammals and birds (Mendoza et al. 2002). As with many ichthyosporeans, species now recognised as rhinosporideacids were misidentified as Fungi, as well as apicomplexans, haplosporeans and cyanobacteria (Ahluwalia et al. 1997; Mendoza et al. 2002). Most described species conform to a parasitic life cycle, in which the infectious dispersal agent is a flagellated zoospore. Upon infecting a new host the zoospore encysts and produces a walled sporangium (cyst) in the host. The sporangia then increase in size to 200–400 mm in diameter, whilst cells undergo division to produce thousands of zoospores. Zoospores range across 7–15 mm in diameter (Herr et al. 1999) and are either released directly into the original host tissue or into the environment to infect a new host (for a review of the life cycle, see Mendoza et al. 2002). Infections caused by members of Rhinosporideacae vary in both host and tissue affected. Dermocystidium sp. within fish can be highly virulent, with sporangial numbers overwhelming the gills and resulting in asphyxia (Mendoza et al. 2002). The rosette agent also infects fish, but in the kidneys and spleen (Harrell et al. 1986). Outside of fish hosts, Rhinosporidium seeberi causes rhinosporidiosis in humans (a disease resulting in granulated nasal polyps; Herr et al. 1999) and Amphibiothecum penneri infects the subcutaneous tissue of frogs and toads (Feldman et al. 2005; Jay and Pohley 1981).

Flagellum loss appears to have occurred on at least one occasion within Rhinosporideacae as no flagellated cells have been observed in Rhinosporidium seeberi. Tubular mitochondrial cristae were initially described in the rosette agent (Arkush et al. 1998); however this appears to have been a misidentification and all known rhinosporideacids have flat cristae (Mendoza et al. 2002).


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Fig. 1.3. Simplified four-gene phylogeny of the major holozoan groups. The tree was derived by a maximum likelihood method, using RAxML 7.0.3, from partial sequences of rRNA (SSU, LSU) tubA and hsp90. The 6386 aligned nucleotide positions were analysed using the GTRCAT substitution model, with separate partitions for rRNA, first + second codon positions and third positions. Support values are also shown from a maximum likeli-

hood tree derived using MrBayes 3.1.1 utilizing the GTR+I +G substitution model; mlBP and biPP values are given above and below branches respectively. The long-branch and phylogenetically unstable Capsaspora owczarzaki, Corallochytrium limacisporum and Ministeria vibrans have been omitted. Branches are drawn proportional to the number of nucleotide substitutions per site as indicated by the scale bar at the lower left

B. Ichthyophonae

Amoebidium parasiticum is a non-pathogenic symbiont which attaches itself to the external exoskeleton of its insect hosts (Benny and O’Donnell 2000). Two species, Sphaeroforma arctica and LKM51, are also suspected to be saprotrophs, with the latter also possibly planktonic (Mendoza et al. 2002). Both flagellated and amoeboid cells are unknown in a number of ichthyophonid species. Anurofeca richardsi, originally assigned to the parasitic algal genus Prototheca (Kruger 1894), forms spherical cells in the guts of anuran (frog, toad) larvae. Within these cells a small number (200 species (Carr 2008a). Some choanoflagellate species may form colonies, under suitable conditions. These colonies may be plate-like assemblages on a substratum or free-floating globules of cells (Fig. 1.4A). Globate colonies are held together by a gelatinous matrix which resembles the mesohyl of some poriferans (Buss 1987). As a result of these similarities a genus within the choanoflagellates was erected under the name Protospongia (later amended to Proterospongia). Despite initial speculation (Kent 1880), the Proterospongia colonies are not an evolutionary intermediate step between choanoflagellates and the multicellular metazoans. Rather it appears to be an ephemeral stage in the life cycle of codosigid/salpingoecid species, with no choanoflagellate species having a solely colonial existence (Carr et al. 2008a). The taxonomic position of the choanoflagellates has been the source of much debate and speculation.


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Fig. 1.4. (A) Colony of the choanoflagellate Salpingoeca amphoridium. The characteristic flagella and collar face outward to maximise predation of bacteria. (B) The nudiform choanoflagellate Savillea parva surrounded by its silica lorica. Pictures with kind permission from Barry Leadbeater

They were proposed as a close relative to Metazoa over 140 years ago as a result of the similarity between their protoplasts and the feeding cells (choanocytes) of poriferans (Haeckel 1874; James-Clark 1866, 1868). However they have also been considered to be algae (Bourrelly 1968; Chadefaud 1960), highly reduced members of Metazoa (Maldonado 2004) and even a paraphyletic assemblage ancestral to both Fungi and Metazoa (Cavalier-Smith 1987).

Molecular phylogenetics has slowly answered this question by first placing choanoflagellates firmly within Opisthokonta (Wainright et al. 1993) and then allied with both Metazoa and Ichthyosporea (Amaral-Zettler et al. 2002; Baker et al. 1999; Cavalier-Smith and Chao 2003; King and Carroll 2001; Lang et al. 2002; Lavrov et al. 2005; Medina et al. 2003; Ruiz-Trillo et al. 2004; Steenkamp et al. 2006). Most early rDNA studies showed a sister-grouping between the choanoflagellates and ichthyosporeans. However, recent multigene phylogenies show choanoflagellates forming a robust sister-grouping with Metazoa (Carr et al. 2008a; Ruiz-Trillo et al. 2008; Shalchian-Tabrizi et al. 2008). This sister group relationship allows some tentative conclusions to be drawn about the nature of the last common ancestor (LCA) of choanoflagellates and metazoans. This ancestor was almost certainly a marine protist. It also probably possessed a flagellum surrounded by a collar of microvilli that would have been employed for filter feeding. Since a sedentary lifestyle appears to

have been ancestral in choanoflagellates (Carr et al. 2008a) and, since adult poriferans are also sedentary, the last ancestor of both choanoflagellates and Metazoa was probably also a sedentary organism employing a flagellate swimming stage for dispersal. The ability to form colonies is present in two of the three major clades of choanoflagellates (Carr et al. 2008a). It is therefore possible that the last common ancestor of the choanoflagellates and Metazoa could also form colonies, and thus that this feature was already present in the LCA of Metazoa. The genome of the marine choanoflagellate Monosiga brevicollis has recently been sequenced (King et al. 2008). Comparative genomics now allows a greater understanding of the differences and similarities in the genetic makeup of choanoflagellates and metazoans (Abedin and King 2008; Carr et al. 2008b; Kodner et al. 2008; Manning et al. 2008). Earlier sequencing of ESTs from three species of choanoflagellate (M. brevicollis, M. ovata, Proterospongia sp. 50818) genome indicated the of presence proteins, such as cadherins, receptor tyrosine kinases and C-type lectins, thought to be unique to Metazoa (King and Carroll 2001; King et al. 2003). At the time of writing, the roles of these proteins in the choanoflagellate cells have yet to be determined.

Although molecular phylogeny indicates that only one of the major divisions of choanoflagellates in fact corresponds to one of the three traditional families, the taxonomy of the group has yet to be formally revised.

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A. Codosigid Morphology The codosigid morphology (choanoflagellate clades 1 and 2; Carr et al. 2008) is found in two of the three major clades of choanoflagellate, and used to define the traditional family Codosigidae (Carr et al. 2008). This morphology consists of a periplast of a fine mucilaginous cup, known as the glycocalyx, which extends from the cell to form a stalk of carbohydrate microfibrils (Leadbeater and Morton 1974). Cell division occurs within the glycocalyx, as it has a flexible nature. The daughter cell may then disperse by temporarily using its flagellum for locomotion. Alternatively, the juvenile may remain attached to the parental cell stalk, which can result in multiple head cells being present on a single stalk. This is not observed in species with the salpingoecid morphology or members of the Acanthoecidae, because both possess rigid periplasts which prevent simple lateral cell division (Carr et al. 2008a; see below). The glycocalyx appears be an ancestral character of the choanoflagellates which has been retained in all extant species. All other choanoflagellates possess additional periplast structures, so the codosigid morphology is defined by the absence of additional structures rather than the possession of any unique feature. (Carr et al. 2008a).

B. Salpingoecid Morphology The salpingoecid morphology (choanoflagellate clades 1 and 2; Carr et al. 2008) is defined by the presence of an additional thick organic periplast called a theca. Like the glycocalyx, the theca is formed from carbohydrate microfibrils but the latter has a considerably more rigid structure. This rigidity prevents simple cell division within the periplast, so, unlike in codosigids, the cell becomes amoeboid and partially exits the theca prior to cell division. Molecular studies indicate that the codosigid and salpingoecid morphologies are dispersed between Clade 1 and 2 choanoflagellates, being absent only in Clade 3 (traditional Acanthoecidae; Carr et al. 2008a). Clades 1 and 2 themselves form a strongly supported monophyletic group (Carr, Leadbeater and Baldauf, unpublished data). Since the theca is a complex mulit-layered structure, it is unlikely to have arisen multiple times independently. The Salpingoecidae should therefore be

considered a paraphyletic family, with the polyphyletic Codosigidae nested within it. Under this scenario the theca appears to have been the ancestral periplast of sedentary cells in this group; the theca has therefore been reduced, or lost, in species designated to Codosigidae. The unification of the salpingoecids and codosigids into a single taxonomic group was previously proposed by Cavalier-Smith and Chao (1997), under the name Craspedida.

C. Acanthoecidae Unlike the Codosigidae and Salpingoecidae, which were erected in the 1880s, the Acanthoecidae was described relatively recently (Norris 1965). The group is characterized by a highly distinct periplast, the lorica, which consists of a complex basket of silica strips surrounding the protoplast (described in detail by Leadbeater et al. 2009). The lorica is constructed from two layers of strips; an external layer of longitudinal strips is held in position by an internal layer of horizontal strips, which can be in the form of rings or helices. Organic microfibrils are frequently arranged between the strips to create an enclosed column for water to funnel through. Some acanthoecids are sedentary, attached to the substratum by a peduncle which extends from the lorica. However in contrast to the other choanoflagellates many acanthoecids are pelagic species. The lorica is a complex structure that varies across the group. Two major types are known and these are also associated with differences in cell division and the mechanism of lorica construction (Fig. 1.3; Carr et al. 2008a; Leadbeater et al. 2008, 2009). Nudiformes species have helical horizontal strips, but are never found with rings of strips in their lorica (Fig. 1.4B). In contrast tectiforms are always found with rings and only certain species possess helical strips. All acanthoecids fall into one of these two groups, which form monophyletic sister-groups (Fig. 1.3; Leadbeater et al. 2008). In common with codosigid and salpingoecid species, nudiform acanthoecids produce swimming dispersal cells. The length of the dispersal stage in nudiforms is considerably shorter than that in other choanoflagellates and cells rapidly settle to produce their lorica. In contract, in tectiform species, the entire set of lorica strips is


pre-assembled so that the daughter cell erects its own lorica immediately after separating from the parent cell. Therefore free-swimming juvenile dispersal cells are never produced (Leadbeater 1979). Acanthoecidae is the only choanoflagellate family whose members are exclusively marine (Pettitt et al. 2002). They also include the only known pelagic species. Of the two major divisions of the group, the nudiform is the smaller with fewer than ten known species. These mainly inhabit specialised ecological niches such as bacterial biofilms. In contrast the tectiforms are the most speciose division of the choanoflagellates and inhabit a broad range of habitats. Tectiforms are the only choanoflagellates that do not lead sedentary lives; many are free-floating, with some present in the water column whilst others are benthic and drift in water currents (Carr et al. 2008a).

VIII. Incertae Sedis Opisthokont Protists A. Aphelidea The parasitic Aphelidea make up one of the more neglected groups of opisthokonts. They were initially classed as phycomycete fungi (Bozarth 1972; van Etten 1991) and then proposed as members of Rhizopoda (Gromov 2000), before being placed in Ichthyosporea by Cavalier-Smith et al. (2004). Unlike many taxa classed as ichthyosporeans they do not parasitise metazoans, but are intracellular parasites of algae (Adl et al. 2005). Immature dispersal cells can be either flagellated or amoeboid and are released from the host to invade new host cells. The phylogenetic position of the group within the opisthokonts is uncertain since nucleotide sequence data are currently unavailable. B. Capsaspora At present the genus Capsaspora is only represented by a single species, C. owczarzaki. Capsaspora owczarzaki was long considered to be a nucleariid, despite a number of morphological differences (Owczarzak et al. 1980), and accordingly placed in Amoebozoa rather than the Opisthokonta. Based on morphological and phylogenetic differences to other designated Nuclearia species, Hertel et al. (2002) erected the new genus Capsaspora to accommodate this amoeboid opisthokont.

15

The morphological similarities to Nucleariida are only superficial, with a suite of characteristics distinguishing C. owczarzaki. Like the nucleariids the cells of C. owczarzaki are amoeboid with long unbranching filopodia, however the latter are considerably smaller (3–7 mm in diameter) than any described nucleariid (Hertel et al. 2002). C. owczarzaki lacks a mucoidal extracellular matrix and possesses flat mitochondrial cristae rather than the discoidal cristae seen in most nucleariids (Amaral-Zettler et al. 2001). A further major difference between C. owczarzaki and Nucleariida is in the mode of nutrition. Nucleariids mainly engulf their prey via phagocytosis whilst C. owczarzaki uses a feeding peduncle (Owczarzak et al. 1980). The complete life cycle of C. owczarzaki has yet to be observed, but at present no stages have been shown to produce flagellated cells (Hertel et al. 2002). In contrast to the free-living nucleariids, C. owczarzaki exists as an endosymbiont of the pulmonate snail Biomphalaria glabrata (Stibbs et al. 1979). B. glabrata strains may also harbour the parasitic trematode Schistosoma mansoni and it has been shown that the presence of C. owczarzaki offers a degree of immunity to this infection (Stibbs et al. 1979). Capsaspora cells present in the snail haemolymph attack and kill trematode sporocysts; this protection comes at a certain cost, as C. owczarzaki may also act as a parasite of its snail host (Ruiz-Trillo et al. 2006).

The phylogenetic position of C. owczarzaki remains contentious. One major problem is that no close relative has been identified, therefore in small-scale phylogenies C. owczarzaki tends to be found on an unstable, long branch within Holozoa (Amaral-Zettler et al. 2001; Carr et al. 2008a; Cavalier-Smith and Chao 2003; Hertel et al. 2002; Ruiz-Trillo et al. 2004). Recent phylogenomic studies have produced robustly supported, yet contrasting positions for C. owczarzaki. Two large-scale multigene studies (Burger et al. 2009; Shalchian-Tabrizi et al. 2008) placed C. owczarzaki as a near relative of the sedentary marine protist Ministeria vibrans (see below). In order to accommodate this relationship, both Capsaspora and Ministeria were placed in a newly erected group, termed Filasterea (Shalchian-Tabrizi et al. 2008). This putative taxon is characterised by a naked protoplast which exhibits long tapering tentacles that contain an internal skeleton of actin microfilaments.

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Both studies placed the Filasterea as closer relatives of the Metazoa and Choanoflagellida than to the other members of Holozoa. Although the monophyly of Filasterea has yet to be tested in other studies, numerous published phylogenies have also positioned C. owczarzaki as a closer relative to the choanoflagellates and metazoans than the ichthyosporeans (Dunn et al. 2008; Philippe et al. 2009; Ruiz-Trillo et al. 2006, 2008). In contrast to this position, Ruiz-Trillo et al. (2008) also produced a 110-nuclear-protein phylogeny that robustly placed C. owczarzaki with the ichthyosporeans Amoebidium parasiticum and Sphaeroforma arctica at the base of Holozoa. Data from the draft C. owczarzaki genome, recently produced by the Broad Institute, may finally allow an accurate placement of this enigmatic protist within Holozoa.

C. Ministeria The genus Ministeria comprises two known species, M. vibrans and M. marisola, of which only the former has been successfully maintained in culture (Patterson et al. 1993). The group is of considerable interest with respect to both its unusual morphology and phylogenetic position; despite this it remains one of the more poorly studied groups in Opisthokonta. Unlike all other known opisthokont lineages, Ministeria does not exhibit either amoeboid or flagellated cells. The spherical protoplast of M. vibrans attaches to the substratum via a peduncle, which may under certain conditions vibrate. There has been speculation that the peduncle may be a modified flagellum (Cavalier-Smith and Chao 2003), but this remains to be confirmed. Ministeria protoplasts are small (5 bp from all previously known genotypes. Phylogenetic analyses demonstrated 200 previously undescribed phylotypes, clustering into 45 clades throughout the fungal phylogeny.

This one publication increased the number of described yeasts by 30%, and it demonstrates the importance of exploring differing environments to enable a greater understanding of fungal diversity (Suh et al. 2005). Our understanding of fungal evolutionary complexity is expanding as researchers use increasingly direct environmental DNA methods

to investigate microbial diversity. Recent work using environmental ITS sequencing of soil samples has suggested a further upward revision of global fungal diversity from 1.5 to 3.5–5.1 million species, based on conservative 97% ITS similarity clustering (O’brien et al. 2005). Independent of the accuracy of these estimates it suggests that less than 2–6% of fungal species are currently described. Furthermore, of the 77269 species described (Hawksworth 2001) to date 478 fungal genome sequence projects are listed as complete or in progress (Gold database http:www.geno mesonline.org). This demonstrates the effort required to narrow the huge diversity gap between the fungi isolated, cultured, and studied, and fungal communities present in natural environments. In this chapter we will review the progress made in identifying fungal diversity using direct molecular approaches from a range of different environments and begin to determine what these data mean for the fungal tree of life (see Chapter 1 in this volume).

II. Environmental DNA Analysis, the Clone Library Approach Traditionally researchers have investigated fungal diversity by collecting macroscopic reproductive organs or by using culture based isolation techniques. However, such approaches have several limitations, they preferentially sample the few species that are easily propagated into culture and which therefore grow and reproduce at high rates in rich growth media, or species which have larger body sizes and possess distinctive morphologies. This raises a number of difficulties for the study of fungi, primarily because a large proportion of fungal diversity resides in soil and sedimentary environments, highly complex and heterogeneous environments which limiting direct microscopic observations (Arnold et al. 2000; O’brien et al. 2005). An additional problem arises because the majority of fungi that remain uncultured are likely to include huge amounts of cryptic diversity indistinguishable using microscopy of environmental samples. Further complications occur because similar morphotypes can often branch in distant and paraphyletic positions on the fungal tree of life (James et al. 2006a). This

Environmental DNA Analysis and the Expansion of the Fungal Tree of Life

makes classifications based on environmental sample observations extremely difficult and therefore encourages the increasing use of molecular approaches (e.g., Anderson and Cairney 2004; Brock et al. 2009; Seifert et al. 2007; White et al. 1990). Molecular analyses of microbial diversity from environmental samples began with the sequencing of 5S and 16S rRNA gene markers from environmental DNA samples (e.g., Giovannoni et al. 1990; Hugenholtz et al. 1998; Pace 1997). This work borrowed many components from earlier research by Woese and colleagues who developed the use of ribosomal RNA encoding genes to reconstruct phylogeny (e.g Olsen and Woese 1993; Winker and Woese 1991). They followed the logic that all cellular life possessed a ribosome with both small and large subunit ribosomal RNA components and that phylogeny of these rDNA sequences can be used to reconstruct a tree of life (Olsen and Woese 1993; Winker and Woese 1991; see Chapters 1, 2 in this volume). Woese and colleagues were encouraged to focus on the ribosomal RNA encoding genes because the rRNA molecules fold to form a complex secondary structure with loops and helix regions (e.g., Olsen and Woese 1993; Van de Peer et al. 1997; Winker and Woese 1991; Wuyts et al. 2000) which means these gene sequences were composed of regions with a mixture of fast and slow rates of character variation. Consequently, these molecules seemed particularly suited to defining evolutionary relationships, among both relatively ancient and recently derived branches. A second advantage of this molecular characteristic was that polymerase chain reaction (PCR) primers could be developed which were either specific to narrow taxonomic groups, or were comprehensive for a wide diversity of species. This inspired the early use of rDNA sequences for investigating evolutionary relationships among microbes and rDNA sequences were soon among the best-sampled gene databases for identifying the placement of microbial lineages in molecular phylogenies (Cole et al. 2003; Sogin et al. 1986; Maidak et al. 1996; Woese et al. 1990). This approach was adapted for the investigation of microbial diversity present in environmental samples by using PCR amplification of ribosomal RNA encoding genes from environmental DNA samples (see Olsen et al. 1986; Pace 1997). This method is now commonly called environmental clone library analysis or environmental

39

gene library analysis, and has been widely applied to a range of environments and evolutionary groups (e.g., Bass et al. 2007; Giovannoni et al. 1990; Moon-van der Staay et al. 2001; O’brien et al. 2005; Richards et al. 2005; Stoeck and Epstein 2003). This approach follows a six-step methodology with numerous variants (Fig. 3.1, left column): Step 1 involves the extraction of DNA from an environmental sample either as a filtrate from an aquatic sample or directly from sediment or soil samples. We note standard approaches to environmental DNA extraction in many published environmental clone library analysis often do not encompass chemical or physical preparations specifically tailored to the preparation of fungal cells, especially among general eukaryotic molecular diversity studies. Consequently, it is possible that many of these studies have failed to sample true fungal diversity because the DNA preparation protocol lacks chemical or physical cell lysis steps to release DNA from cells enclosed within robust chitin-rich fungal cell walls. As such, future adaptation of this technology for the study of fungal microbes should consider experimentation with additional chemical and physical cell lysis steps. Step 2 involves PCR amplification of environmental DNA using primers designed to target a taxonomic unit of interest. A great body of work is now available on the universal amplification of small subunit (SSU) ribosomal RNA (rRNA) genes of both prokaryotes and eukaryotes, enabling environmental clone library analyses from a range of different environments (e.g., Giovannoni et al. 1990; Moon-van der Staay et al. 2001; Olsen et al. 1986; Pace 1997; Richards et al. 2005; Stoeck and Epstein 2003). A large number of additional studies have also demonstrated amplification conditions specific for fungal SSU rDNA sequences (e.g., Bass et al. 2007; Anderson and Cairney 2004; Anderson et al. 2003; Jebaraj et al. 2009; O’brien et al. 2005; Vandenkoornhuyse et al. 2002) and a selection of these approaches are discussed in more detail below. Step 3 involves the use of a standard cloning approach and involves the ligation of the PCR product into a plasmid vector. During step 4 the population of vectors carrying a selection of different inserts is transformed into host Escherichia coli cells and grown on selective media (step 5). For step 6, subsets of the transformed cells are then picked and the plasmid insert is sequenced

40

Meredith D.M. Jones and Thomas A. Richards Environmental Clone Library analysis of Community Diversity

Environmental 454 amplicon analysis of Community Diversity

Step 1. Extract DNA

Step 1. Extract DNA

Step 2. rDNA PCR amplification

Step 2. rDNA PCR amplification

Step 3. Ligation of amplicons into plasmids

454 Sequencing A. One DNA fragment per bead association and isolation process

Step 4. Transformation of plasmids into E. coli cells

B. Bead based PCR reaction mixture in oil emulsion C. Individual beads are centrifuged into individual picolitre-sized wells (formed by a fibre-optic slide) and sequenced using pyrophosphatebased sequencing

Step 5. Culture of plasmid containing E. coli

Step 6. Clones picked and sequenced using Sanger dideoxy sequencing method

X Y Z

Community structure and diversity parameters identified using sequence analysis

Fig. 3.1. Schematic showing comparison of environmental gene library and 454 amplicon diversity analyses; two methods used for investigating microbial diversity in environmental samples. 454 amplicon analysis allows large-

scale sequencing of an environmental sample without the need of a cloning step minimising some sources of sampling error and radically increasing the magnitude of sequences retrieved


using a standard Sanger dideoxy sequencing approach. Steps 3–6 can all convey considerable sampling biases, which can mean that the results of the clone library sequencing profiles are not equivalent to the true community profile. Finally the resulting sequences are checked for evidence of chimaeras formed during the multitemplate PCR using manual alignment checks (e.g., Berney et al. 2004; DeSantis et al. 2006; Hugenholtz and Huber 2003), partial treeing analyses (Robison-Cox et al. 1995) and/or an increasing number of bioinformatic tools (e.g., Cole et al. 2003; Huber et al. 2004). The remaining sequences are then identified in an alignment of sequences from known taxa and phylogenies are calculated to identify the branching position of the environmental sequences. This approach has been used for both prokaryotes and eukaryotes to place many previously unrecognised branches on the tree of life, in many cases redefining our understanding of the evolutionary complexity of eukaryotic groups among the highest taxonomic grades (Bass and Cavalier-Smith 2004; Dawson and Pace 2002; Edgcomb et al. 2002; Lo´pez-Garcıá et al. 2001; Moon-van der Staay et al. 2001; Richards et al. 2005; Stoeck et al. 2006), although these results have been the subject of much debate and revision (Berney et al. 2004; Cavalier-Smith 2004; Richards and Bass 2005). Furthermore, such techniques have demonstrated that poorly recognised groups are important ecosystem components (Bass and Cavalier-Smith 2004; Massana et al. 2004; Moreira and Lo´pez-Garcia 2002; Chambouvet et al. 2008). For example, early applications of environmental clone library analysis focusing on eukaryotic microbial diversity has demonstrated that protist diversity in marine environments encompasses numerous hitherto unsampled groups of marine stramenopiles (MAST) and alveolates (Lo´pez-Garcıá et al. 2001; Massana et al. 2002, 2004; Moonvan der Staay et al. 2001) radically expanding both groups. Follow up work suggested the novel stramenopile groups consisted of at least 12 separate clades, now labelled as MAST groups 1 to 12 (Massana et al. 2004). While the novel alveolate sequences fall into five extremely diverse clades, which branch close to the mainly photosynthetic dinoflagellate protists (e.g., Guillou et al. 2008). The bulk of these microbial forms remain uncultured, and therefore many aspects of their biology and ecology remain relatively unknown.

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But where does this leave our understanding of fungal biodiversity and how it fits on to the fungal tree of life? Environmental fungal clone library analyses have generally focused on regions within the rRNA gene array, with a mixture of approaches and sequence targets, with some researchers focusing on the SSU rDNA sequence whilst others focus on the internal transcribed spacer (ITS) regions. The two ITS regions are sections of DNA located between the 18S and 5.8S rRNA genes and between the 5.8S and 28S rRNA genes, the variable nature of these regions relative to the flanking rRNA genes enables increased accuracy when assigning sequences to genus and species level classifications within wellsampled groups (Bruns and Gardes 1993; Gardes and Bruns 1993; Horton and Bruns 2001). Assignment is further facilitated by an increasingly wellsampled database of sequences (Buchan et al. 2002; James et al. 2006a; O’brien et al. 2005), although it is important to note that some database sequence classifications may be erroneous and rates of ITS variation can vary between taxonomic groups hindering classifications using these data (Nilsson et al. 2006, 2008; Vilgalys 2003). The ITS regions are commonly amplified using either the universal ITS1 and ITS4 primers, fungal specific primers ITS1f and ITS4f, or increasingly taxon-specific variants, e.g., ITS1b and ITS4b that are basidiomycete-specific (Gardes and Bruns 1993; Horton and Bruns 2001; Anderson et al. 2003).

Molecular studies based around the amplification of the ITS marker from environmental DNA have reported much fungal diversity (e.g., Buchan et al. 2002; O’brien et al. 2005). However, with much of this research focused on ITS1-5.8S-ITS2 comparisons, results are often biased towards previously sampled groups (Anderson and Cairney 2004). Whilst this approach is useful for determining species diversity and can be used for ecosystem comparisons when targeting well defined taxonomic groups, it is of limited value for inferring higher-level phylogenetic relationships and identifying novel groups. This is because phylogenies based on ITS sequences demonstrate weak resolution among phyla level relationships (Horton and Bruns 2001). Therefore, some researchers have taken a different approach and have instead focused on sampling across the SSU rDNA region in order to investigate novel fungal diversity

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Meredith D.M. Jones and Thomas A. Richards

among higher taxonomic groups (e.g., Anderson et al. 2003; Bass et al. 2007; Jebaraj et al. 2009; Vandenkoornhuyse et al. 2002). However, it is important to stress both approaches have benefits and they are not mutually exclusive, with some authors calling for a combination of the ITS and the SSU approaches (O’brien et al. 2005). This is achievable by using a forward primer in the SSU region and a reverse primer in the LSU region, enabling amplification from the SSU, through the ITS1, 5.8S, ITS2, and LSU regions. This approach enables multi gene phylogenetic analyses, which can improve tree support and assignment of environmental sequences to taxonomic classifications (e.g., Porter et al. 2008; Jones et al. 2011). We shall now briefly review how the application of environmental clone library techniques has altered our understanding of the fungal tree of life. We will focus mainly on studies that have sampled the SSU region to evaluate the complexity of the fungal phylogeny at higher taxonomic levels. Eukaryotic SSU ribosomal DNA environmental gene sampling methods have consistently demonstrated a broad range of novel fungal lineages indicating that many potentially key evolutionary branches and environmentally important fungal microbes remain unexplored. This review is not an exhaustive account of all the literature in this area but focuses on key studies which have provided progress towards a fungal phylogeny that accounts for the large complexity of fungal forms present in natural environments that have currently not been isolated and cultured.

A. Fungal Molecular Diversity in Aquatic Environments using a Clone Library Approach 1. Marine Fungi In terrestrial ecosystems fungi perform critical roles in breaking down complex biomolecules and recycling nutrients. This process is very important because it underpins the wider ecosystem, but it is much less clear which organisms perform equivalent roles in marine environments. Culture-based fungal isolation studies of marine environments (e.g., Burgaud et al. 2009; Damare and Raghukumar 2008) have suggested the diversity of fungi present is limited, with only 467

described marine fungal species, belonging to 244 genera, equivalent to ~0.6% of the described fungal species (Kis-Papo 2005). Culture based studies of marine fungi have demonstrated that ‘pink basidiomycete yeasts’ are the most common fungal isolate. Marine yeasts are generally associated with accumulations of nutrient concentrations, e.g., pollution, plankton blooms, and macro algae (Kohlmeyer and Kohlmeyer 1979). Fungi are therefore suggested to play a critical role in detritus processing in marine environments (Mann 1988; Raghukumar 2004) providing essential nutrients to the wider food web such as lysine and methionine amino acids, various vitamins, and sterols (Mann 1988). For example, it is now clear that Crustacea require the polyunsaturated fatty acid, docosahexaenoic acid, for growth (Harrison 1990) which is made accessible to benthic food webs by actions of detritus microbes (Raghukumar 2004). In 2005, during the early days of eukaryotic SSU rDNA clone library analysis, Richards and Bass conducted a meta-analysis of 13 environmental gene library studies encompassing 49 separate SSU rDNA environmental clone libraries and including a total of 1077 environmental sequences from soils, freshwater and marine samples. Of these sequences 124 (11.5%) clustered within, or close to, the Fungi (Richards and Bass 2005). Interestingly, this meta-analysis suggested that although fungi were present in aquatic sediments, low-oxygen aquatic environments, freshwater samples, and soils; the fungi appeared to be almost entirely absent from surface marine waters from both coastal sites and the open ocean. This pattern was confirmed by a separate marine specific meta-analysis which included 23 coastal water libraries (1349 clones) and 12 open sea libraries (826 clones) where only 16 fungal clones, equivalent to 0.8% of marine SSU rDNA sequences, were recovered (Massana and Pedro´sAlio´ 2008). This pattern is a little surprising as fungi are thought to be a major contributor to the decomposition of woody and herbaceous substrates and animal remains in coastal and surface marine environments (Kohlmeyer and Kohlmeyer 1979; Mann 1988; Newell 1996) raising questions of the significance of fungal communities in these marine ecosystems. A study by Jebaraj et al. (2009) examined the diversity of fungi in oxygen-depleted regions of the Arabian Sea using clone library construction.


Multiple primer sets, two fungal specific and one set routinely used for eukaryotic diversity studies, were used to amplify the SSU from samples collected at 25 m depth (Jebaraj et al. 2009). Comparisons between fungal diversity detected using the fungal specific primer sets compared with the universal eukaryotic primers revealed a greater diversity detected within the fungal specific clone library. This result highlights the importance of using multiple primer sets to negate PCR biases and demonstrates that a proportion of fungal diversity is often missed when using ‘universal’ primer sets, potentially caused by the preferential detection of alternative taxonomic groups (Jebaraj et al. 2009; Stoeck et al. 2006). Phylogenetic analysis of SSU clones identified 48 distinct fungal phylotypes (clustered at 99% sequence similarity): 27 branching within the ascomycete radiation, 20 branching within the basidiomycete, while only a single highly unique phylotype was detected among the ‘lower fungi’ branching as a sister to mortierellales taxa (traditionally classified as a zygomycete). A number of the ascomycete and basidiomycete sequences formed highly novel branching positions in the phylogeny and in several cases clustered with additional environmental sequences recovered from oxygen-depleted habitats (Jebaraj et al. 2009). For example, several sequences branched within a clade previously termed the ‘hydrothermal and/or anaerobic fungal group’ identified as part of a study of deep-sea eukaryotic environmental clone libraries (Lo´pezGarcia et al. 2007). No ‘chytrid’ sequences were identified from this study, suggesting the possibility these taxonomic groups have a low diversity in these marine environments or alternatively the PCR primers or DNA sampling methodology used for this study were collectively biased towards the ascomycetes and basidiomycetes.

2. Deep-Sea Environments The majority of the planet is covered by ocean with an average depth of ~3200 m (Gage and Tyler 1991) deep-sea environments therefore represent the largest and most underexplored habitats on Earth. The deep-sea environment is comprised of a variety of habitats such as lownutrient water, sediments, and in some areas hydrothermal vents associated with high temperature and extreme pH. Preliminary exploration of

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these environments is redefining our understanding of both marine biodiversity and the tree of life, with an average of two new species discovered within deep-sea waters each month (Fisher et al. 2007), yet we are far from understanding many of the ecological processes occurring in deep-sea environments. Using SSU rDNA environmental PCR, followed by clone library analyses, Bass et al. (2007) investigated the composition of fungal communities within deep-sea sediments and water column samples ranging from depths of 500–4200 m and including several hydrothermal vent samples. The sequences recovered showed deep-sea fungal communities to have low diversity, dominated by ascomycete and baidiomycete forms that branched closest to yeast taxonomic groups on phylogenetic trees. Many of these phylotypes were also shown to branch close to known pathogens, perhaps indicative of the presence of fungal pathogens of deep-sea animals and consistent with additional observations suggesting fungi constitute important pathogens in deep-sea ecosystems (Van Dover et al. 2007). The phylogenies reported by Bass et al. (2007) included additional marine environmental SSU rDNA sequences and showed seven clusters of highly unique environmental sequences branching within the opithokonts, six specifically within the fungal radiation, indicating the presence of unknown fungal forms in marine environments. A further study by Le Calvez et al. (2009) using an environmental SSU rDNA clone library approach to study deep-sea hydrothermal vent ecosystems also demonstrated the recovery of several novel fungal lineages. These lineages included three ‘unknown phylotypes’ branching within the basidiomycete radiation, and a further two ‘unknown phylotypes’ branching with ‘chytrid’ taxa (Le Calvez et al. 2009). The primer set used in this study was different from that chosen by Bass et al. (AU2–AU4; Vandenkoornhuyse et al. 2002) but both studies demonstrate the identification of several novel fungal lineages and that deep-sea fungal communities, although relatively non-diverse, are dominated by a large proportion of species branching among the ascomycete and basidiomycete yeast species. This observation is consistent with molecular analysis of eukaryotic diversity in marine sediments, which has demonstrated yeast species can dominate deepsea microbial eukaryotic communities (Takishita et al. 2006).

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3. Freshwater Fungi A number of papers report a SSU rDNA clone library approach to investigate the eukaryotic microbial community of freshwater environments, from either direct environmental DNA extractions (Amaral Zettler et al. 2002; Berney et al. 2004; Lefe`vre et al. 2007; Lefranc et al. 2005; Slapeta et al. 2005) or environmental DNA extracted from experimental detritus ecosystems, created from dead algae which have been inoculated with lake water microbial communities (van Hannen et al. 1999). In contrast to many of the equivalent marine studies (Massana and Pedro´sAlio´ 2008), freshwater studies have reported a high proportion of fungal sequences among the eukaryotic clones recovered (e.g., 19%, Lefe`vre et al. 2008; 23%, Lefe`vre et al. 2007; 33%, Berney et al. 2004; or 25%, Lepe`re et al. 2006). Furthermore, in contrast to the majority of sequences recovered from Fig. 3.2. Bar chart showing comparisons of the numbers of fungal phylotypes recovered during multiple studies of freshwater and marine samples. Whilst the differing clone library sizes, methodologies, and primers mean that comparisons cannot be directly drawn between the two environment types. Dikarya appear to dominate the marine environment, whilst ‘lower fungi’ appear dominant in the freshwater samples

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marine environments (e.g., Bass et al. 2007; Jebaraj et al. 2009), a large proportion of the fungal freshwater sequences branch among ‘lower fungi’ and/or ‘chytrid’ clades, where the majority of sequences recovered from marine environments are Dikarya (Fig. 3.2). These results suggest a large and underexplored diversity of ‘lower fungal’ groups in freshwater environments (Lefe`vre et al. 2007, 2008; Lefranc et al. 2005; Slapeta et al. 2005) and suggests a contrasting pattern of fungal diversity between freshwater and marine environments although further and equivalent sampling is required to investigate this pattern. 4. Novel Basal Branches in the Fungal Radiation Interestingly, freshwater studies have also identified a large and complex clade of environmental

Freshwater fungi identified using 3 different general eukaryotic primer sets

Marine fungi identified using 6 different primer sets (5 fungi-specific)

Dikarya

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“Lower fungi”

30

20

10

Le Calvez et al. (2009) hydrothermal vents

Bass et al. (2007) deep-sea

Jeberaj et al. (2009) oxygen depleted Arabian sea environments

Lefèvre et al. (2008) lake water picoeukaryotes 2

Lefèvre et al. (2007) lake water picoeukaryotes 1

Lefranc et al. (2005) lake water columns

Berney et al. (2004) stream sediment

0


sequences that form one of the deepest branches in the Fungi (e.g., Berney et al. 2004; Lefe`vre et al. 2007, 2008). van Hannen et al. (1999) first identified this group from sequencing DNA recovered from an experimental detritus system composed of dead cultured algae seeded with lake water microbial communities. Of the four fungal-like sequences recovered in this experiment, three formed this deep-branching clade, often named LKM11 after the first clone sequenced from this phylogenetic group (van Hannen et al. 1999). Follow up sequencing of freshwater environments have considerably expanded the diversity of this clade (Lefe`vre et al. 2007, 2008; Lefranc et al. 2005) while Lepe`re et al. (2006) have used terminal restriction fragment length polymorphism to demonstrate that the LKM11 group are highly abundant in the lake water column and linked to differential algal population abundances. Further phylogenetic analysis, with additional sampling, suggests that the LKM11 group is extremely diverse and branches with the intracellular parasitic genus Rozella, suggested to be the first branch in the fungal radiation (James et al. 2006a; Jones et al. 2011; Lara et al. 2010; see Chapter 1 in this volume). This is an interesting relationship because the genus Rozella comprises of intracellular parasites which do not form a standard fungal cell wall and are also capable of phagotrophy (Held 1981). These characteristics represent distinct differences to standard fungal phenotypic characteristics which have a rigid chitin wall, feed by osmotrophy, and cannot perform phagotrophy, thus calling into question whether the Rozella genus and perhaps the LKM11 clade of environmental sequences should be classified within the true Fungi, or whether they represent an intermediate form (Lara et al. 2010; Liu et al. 2009). Jones et al. (2011) used a FISH strategy to reveal that cells of this group have cyst, flagellated zoospore, and epibiotic lifecycle stages. Furthermore, co-staining with cell wall specific markers revealed that these cells lack chitin/cellulose-rich walls during the observed stages of their life-cycle, a characteristic previously considered one of the defining features of the fungal kingdom. Jones et al. name this group the cryptomycota (hiddenfungi), in anticipation of formal classification. Interestingly, both the cryptomycota and the chytrid-like sequences recovered from freshwater environments all group most closely to fungi known to parasitise algae, other fungi, or protists

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(Held 1981; Lefe`vre et al. 2007, 2008; Lefranc et al. 2005). These observations have led some authors to re-evaluate the functional role of microbial eukaryotes, and especially fungal parasites in freshwater environments, suggesting they play a very significant role in the microbial loop of freshwater ecosystems (Lefe`vre et al. 2008).

B. Fungal Diversity in Soil and Plant Environments using the Clone Library Approach A number of studies have used environmental clone library analyses to investigate the presence and diversity of fungal taxa in soils and plantassociated environments (Buchan et al. 2002; Horton and Bruns 2001; Vandenkoornhuyse et al. 2002), with the aim to identify important ecological agents associated with plant derived ecosystems. Other studies have focused on analysing the diversity of eukaryotes (Lawley et al. 2004; Lesaulnier et al. 2008) and fungi (O’brien et al. 2005; Schadt et al. 2003) from a range of soil habitats in order to investigate the complexity of these communities. Together this work has demonstrated that fungi represent a significant fraction of soil communities as identified using general eukaryotic environmental gene library studies (e.g., 20%, Lawley et al. 2004; 28–30%, Lesaulnier et al. 2008). Schadt et al. (2003) used environmental clone library analysis to reveal microbial activity in snow-covered tundra soil, an environment which until recently was thought to harbour only inactive forms (Schadt et al. 2003). Clone libraries generated from total DNA revealed this community comprised largely of ascomycetes, of which 40% clustered into novel and distinct branches, indicating a large diversity of previously unidentified ascomycete groups. These newly identified phylotypes split into two major clades, branching separately within the ascomycete radiation. Using comparisons of clone abundance, Schadt et al. (2003) demonstrated the abundances of these two groups appeared to be coupled to opposite seasons, confirming an active and dynamic fungal community to be present in snow covered soils. Porter et al. (2008) further examined group I of these novel organisms by investigating soil samples using group specific nested PCR primers to amplify the SSU-ITS1-5.8SITS2-LSU regions of the rRNA gene array. This approach allowed the construction of soil clone group 1 libraries

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from a broad range of geographic and environmental locations, and demonstrated the presence of the group within alpine and forest tundra, tropical, and forest soils from a variety of locations around America, Europe, and Australia. Phylogenetic analyses of the recovered 2.4-kb sequences firmly rooted the group within the ascomycetes (Porter et al. 2008). It is also of note that two thirds of the taxa detected in this study were in low abundance; indicating saturation of the library was not achieved. Therefore, it is likely the diversity of the ascomycete soil clone group 1 is far greater than currently sampled.

Fungi are commonly associated with plant roots where they form mutualistic mycorrhizal associations. Vandenkoornhuyse et al. (2002) used SSU clone library generation to reveal novel fungal forms present on the roots of the grass Arrhenatherum elatius (Vandenkoornhuyse et al. 2002). From 200 clones, 49 different sequences were obtained distributed across all traditionally classified fungal phyla (1 ‘chytrid’, 8 zygomycetes, 16 basidiomycetes, 25 ascomycetes). Interestingly, Vandenkoornhuyse et al. (2002) note this diversity pattern is in stark contrast to culture isolation based protocols that recover only ascomycete taxa (Arnold et al. 2000), suggesting fast growth and a preference for nutrient-rich media is a bias for these ascomycetes. Of these sequences, only seven showed greater than 99% identity to known species, while a large proportion of the remaining sequences formed four novel phylogenetic clusters that branched within the basidiomycete and ascomycete radiation. These results again demonstrate numerous additional branches within the fungal phylogeny raising questions regarding the ecological implications of these novel species and the role of these novel groups in the mycorrhizae and general soil habitats.

C. Limitations of the Clone Library Approach Whilst environmental gene library techniques have proved to be very useful, revealing numerous previously undetected fungal groups, the generation of clone libraries is known to result in biases and to suffer from a number of experimental limitations. The majority of the studies discussed use single primer sets and sample 400 MY ago, exemplify the need for molecular data to permit meaningful AMF characterisation.

C. Multi-Gene Analysis and Analyses of Protein Coding Genes James et al. (2006) investigated the evolution of fungal lineages through a multi-locus phylogenetic analysis using super-matrix (nucleotide and amino acid sequences) and individual gene analyses. This presently provides the most comprehensive picture of fungal evolution, based on six gene regions for the SSU rRNA, LSU rRNA, 5.8S rRNA, EF1a, and two RNA polymerase II subunits (RPB1, RPB2). Although resulting from a large international effort, it included just five species of Glomeromycota (Scutellospora heterogama, Funneliformis mosseae (synonym Glomus

Arthur Schu¨ßler and Christopher Walker

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FAMILY

Glomerales Glomeraceae

Claroideoglomeraceae Diversisporales Diversisporaceae Acaulosporaceae Entrophosporaceae 2 Gigasporaceae Pacisporaceae

GENUS Glomus Funneliformis Rhizophagus Sclerocystis Claroideoglomus Redeckera Diversispora Otospora 1 Acaulospora Entrophospora Gigaspora Scutellospora Racocetra 3 Pacispora

Archaeosporales Geosiphonaceae Ambisporaceae Archaeosporaceae

Geosiphon Ambispora Archaeospora

Paraglomerales Paraglomeraceae

Paraglomus

Fig. 7.1. Schematic phylogenetic relationships based on rDNA phylogenies and a molecular phylogeny-based systematics of the phylum Glomeromycota (for references, see text; see also Schu¨ßler and Walker 2010, Kru¨ger et al. 2011). Notes 1–3 are as follows. 1 The genus Otospora (Palenzuela et al. 2008) was placed in the Diversisporaceae based on two partial non-overlapping SSU rDNA sequences obtained from soil trap culture specimens. It may be correct but, due to some morpho-taxonomic

discrepancies, we suggest its phylogenetic position should be treated with caution pending verification. 2 A recently erected family (Sieverding and Oehl 2006) containing one genus with two Entrophospora species, neither of which is phylogenetically characterised. 3 The genus Racocetra was the only one accepted in a revision (Morton and Msiska 2010) which rejected the proposal (Oehl et al. 2008) to split the genus Scutellospora into three families and six genera

mosseae), Rhizophagus irregularis (synonym G. irregulare) DAOM197198, Ge. pyriformis, Paraglomus occultum) and such multi gene datasets are lacking for several main phylogenetic AMF lineages. Nevertheless, the study provided sound support for earlier results (Schu¨ßler et al. 2001) positioning the Glomeromycota as a monophyletic clade and also, with weak support, as a sister group to the Ascomycota and Basidiomycota lineage (the Dikarya). This sister group relationship recently has been questioned, mainly based on the first sequenced mitochondrial genome from Rhizophagus irregularis isolate 494 (Lee and Young 2009) that indicated that AMF have a common ancestor with the zygospore forming Mortierellales. Confirmation, however, will only be possible when data from more species and additional nuclear genes is forthcoming. Regardless of this, the split between the ancestral Glomeromycota and their closest relatives took place at least 600 MY ago, and the phylum level for AMF is well defined and accepted. Some recent detailed phylogenetic analyses of the gene for b-tubulin

from a larger species sampling confirmed the intra-phylum topologies found in rDNA analyses (Morton and Msiska 2010). A few further proteins were studied for phylogenetic inference, including those for a-tubulin, H+-ATPase, elongation factor 1-a (EF-1a) and actin. A study of genes coding for the elongation factor EF1a (tef) and actin (act) in combination with the SSU rDNA phylogeny (Helgason et al. 2003) challenged the SSU rDNA phylogeny, suggesting that the Acaulosporaceae clustered together with the Glomeraceae, not the Gigasporaceae. However, the taxon coverage was very low and a phylogenetic signal in the sequences studied poor. Tanabe et al. (2003) proved that tef sequences are not useful for the reconstruction of deep branches within the Fungi. Moreover, Bayesian analyses can give erroneously high support to incorrect tree topologies, particulary for short and highly variable sequences. This might be exemplified by the Ascomycota placement as a sister group to a lineage comprising the Mucorales and a Glomeromycota–Mortierellales–Basidiomycota clade which renders the Dikarya (Asco- and Basidiomycota) nonmonophyletic in Helgason et al. (2003). Several publications described the analysis of H+ATPase sequences. Ferrol et al. (2000) reported a gene family coding for five plasma membrane H+-ATPase

Evolution of the ‘Plant-Symbiotic’ Fungal Phylum, Glomeromycota isoforms in Funneliformis mosseae (GmHA1-5) and assumed an origin of the variants by duplication and horizontal gene transfer (HGT). In the second study (Requena et al. 2003) a large H+-ATPase gene family was not supported because only one additional H+-ATPase gene, GmPMA1, could be isolated. The fact that from F. mosseae genomic DNA only GmPMA1 and GmHA5 could be isolated and that the GmHA1-4 sequences clustered within the Ascomycota indicated their contaminant origin, which later was confirmed in a study of Corradi et al. (2004a). Requena et al. (2003) therefore rejected the view that AMF form large H+-ATPase gene families. In the study of Corradi et al. (2004a) the H+-ATPase tree topology gave unexpected results, for example Basidiomycota as sister group of plants (with high bootstrap support) and failed to resolve the fungi as a natural group showing that H+-ATPase genes are not useful markers for AMF evolution. Another common fungal gene family is that coding for the P-type II ATPases, crucial for the cell to adjust to changing external K+, Ca2+ and Na2+ concentrations. Corradi et al. (2006) reported four sub-families of P-type II ATPases from Rh. irregularis DAOM197198, Rh. diaphanus and Rh. proliferus. They summarised that specific gene duplications as well as indel mutations among coding regions caused considerable variation in AMF individuals within populations. Tubulin genes (tub) are abundant and often used for phylogenetic inference. Two studies (Corradi et al. 2004a, b) revealed that several earlier published tub sequences were derived from contaminants. Such contaminant sequences are a serious problem in AM research and were also identified for rRNA genes (Redecker et al. 1999; Schu¨ßler 1999). Unfortunately, GenBank still rejects third party correction or tagging of erroneous annotation (Bidartondo et al. 2008), so such errors will further multiply. The tub analyses were the first ones dealing with a protein coding gene that covered more than four AMF species. A tree topology from a-tub conflicting with SSU rDNA phylogenies placed the Acaulosporaceae as a sister group to Diversisporales–Glomerales. However, the b-tub gene tree was congruent with the SSU rDNA phylogenies, but AMF sequences clustered together with those from Chytridiomycota at the base of the fungal phylogenetic tree. The authors stated that this is probably biased, because the tub genes of the Glomeromycota and the Chytridiomycota have a much lower mutation rate than those of the Asco-, Basidio- and Zygomycota and the Microsporidia. Also the Microsporidia placement within the zygomycetes can be considered to be probably an artefact (see James et al. 2006). An interesting question raised was why the a- and b-tub genes evolve at such different speeds in the different lineages. The molecular distances (Keeling 2003; Corradi et al. 2004a, b) show that sequences from Choanoflagellata and Animalia evolve at similar rates to those of Glomeromycota and Chytridiomycota. It seems that the tubulins of the other fungal lineage evolved extraordinarily rapidly, which might be explained by the transition of lifestyles. The chytridiomycotan aquatic lifestyle, including flagellate stages, has remained unchanged for a very long time. An acceleration of tubulin evolution in other lineages could be correlated with the loss of the 9+2 microtubule (flagellum) structures. But

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AMF also do not possess flagella. Did the tubulin of the Asco-, Basidio-, and (some) Zygomycetes evolve so rapidly, because these fungi adapted to changing terrestrial habitats and lifestyles including saprobism, parasitism, and mutualism? From fossil evidence, the Glomeromycota, in contradistinction to the remaining terrestrial fungal lineages, have not changed their stable symbiotic lifestyles and habitat for at least 450 MY, possibly >600 MY. Perhaps the slower evolving tubulin cytoskeleton is correlated with morphological and functional stability over a long geological time (the same AMF morpho-species are found on many continents), as also is the assumed asexual lifestyle.

IV. The Origin of Glomeromycota and the First Land Plants The AMF are obligate biotrophs with vascular land plants and also many lycopods and bryophytes and phylogenetic studies have increased the understanding of evolutionary aspects of the AM symbiosis. A. The Fossil Record In relation to land plant evolution, unequivocal embryophyte traces are more than 470 MY old (Early Middle Ordovician; from eastern Gondwana), representing diverse cryptospore assemblage and sharing characters with those of extant liverworts (Rubinstein 2010, see also Wellman et al. 2003). The diversity of the assemblage implies an earlier, maybe even Cambrian, origin of embryophytes. Early vascular plants existed ~420 MY ago (Middle Silurian; Stewart and Rothwell 1993; Cai et al. 1996). Therefore, a minimum age of 420 MY for the liverwort–vascular plant divergence must be assumed and, based on the fossil record, bryophyte like land plants already were present 510–470 MY ago. Also the AMF have an ancient fossil record history of >460 MY. The earliest reliable evidence for AM in seed plants occurs in the form of non-septate hyphae, vesicles, arbuscules and clamydospores in silicified roots of the Triassic cycad Antarcticycas schopfii (Stubblefield et al. 1987; Phipps and Taylor 1996). Many of the oldest and best preserved fungal fossils in association with plants are known from the Rhynie chert, radiometrically dated to the early Devonian (¼ Pragian). These ~400-MY-old fossils include the earliest known direct fossil evidence for Glomeromycota forming

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symbiosis with the early vascular land plants, in Aglaophyton major (Remy et al. 1994), and later this plant was shown also to contain well preserved Scutellospora- and Acaulospora-like spores (Dotzler et al. 2006, 2009). The very well defined arbuscules and spores in Rhynie chert plant cells doubtless are from plant-symbiotic AMF and were the first direct evidence of a 400-MY-old AM symbiosis, The oldest known fossils representing terrestrial fungi are from ~460-MY-old Ordovician dolomite rock of Wisconsin. These fossil hyphae and spores resemble modern AMF (Redecker 2000a) and it was concluded that terrestrial AMF already existed at a time when the land flora most likely consisted only of bryophyte-like ‘lower’ plants, 460 MY ago. Altogether, these data can be taken as strong support for the hypothesis of Pirozynsky and Malloch (1975) that AMF symbioses played a crucial role on the colonisation of the land by plants. The recent discovery of fossil Scutellospora-like spores (carrying a subtending bulbous base and germination shields) from the Early Devonian Rhynie chert may indicate that the family Gigasporaceae is older than originally thought. Phipps and Taylor (1996) dated the Gigasporaceae to >240 MY ago (based only on perceived similarity of mycorrhizas with those of extant gigasporacean fungi), but from the fossil spores named Scutellosporites devonicus, the family may have evolved by the Pragian (400 MY ago; Dotzler et al. 2006). Does this mean that all the radiation, even to the younger families in the AMF, already had taken place by then? In debate on molecular clock data for AMF, the characters attributed to Scutellosporites devonicus could considerably influence interpretation when character states based on molecular clock data are fitted. However, there is another option to explain the occurrence of Scutellospora-like fossils 400 MY ago. Recently it was shown that most of the structures resembling germination shields in the 400 MY old Rhynie Chert fossils are from acaulosporoid spores (Dotzler et al. 2009). Such shield structures occur in Scutellospora, Pacispora, and Acaulospora species, which are all in the Diversisporales, but perhaps also in some members of the more ancient Archaeosporales. The occurrence of Scutellospora- and Acaulosporalike organisms 400 MY ago therefore shows that either the major diversification within the Diversisporales (which are much younger than the Archaeosporales and Paraglomerales) already had occurred more than 400 MY ago, or that germination shields are plesiomorphic (‘primitive’ characters, sharing a character state with an ancestral lineage) rather than apomorphic (‘advanced’ characters, distinguishing an organism from others sharing the same ancestor). Moreover, there is a published account of germination in Ambispora (Archaeosporales), which shows a rather complex folded germ-shield like structure (Spain et al. 2006). Germination shield type structures therefore

seem to be symplesiomorphies occurring across widely divergent orders in the Glomeromycota, and Scutellospora/Acaulospora/Ambispora-like AMF fossils with germination shields may display the morphology of ancestral AM-forming members of the Glomeromycota. This would fit to the situation that the evolutionarily more advanced AMF groups (e.g., Rhizophagus in the Glomerales, and Gigaspora in the Diversisporales) are reduced rather than advanced in morphological complexity.

B. Molecular Clock Estimates During the past 15 years molecular clock estimates of the origin of the Glomeromycota have varied by a factor of three and there are no widely accepted values for the age of the main fungal lineages. The known fossil record may only show the minimum age of a lineage. For groups lacking a robust fossil record, molecular clock estimates can be expected to be too recent because fossil calibration points, which are main factors when estimating divergence times (Taylor and Berbee 2006), may not reveal the true age of a lineage. The recently discovered fossils (Dotzler et al. 2006, 2009) could be very valuable for re-calibrating molecular estimates, and Scutellosporites devonicus could support a considerable backdating of the origin of the Gigasporaceae, and with it most other glomeromycotan lineages. However, the possible symplesiomorphy of germination shield-like structures introduces an element of doubt because it is as yet unclear which of the present taxa they represent. In general, the fossil and molecular clock estimates together usually are interpreted to reveal the AMF evolution as parallel with land plants, suggesting that both evolved more or less concurrently, but molecular clock scenarios clearly indicate that glomeromycotan fungi evolved much earlier than land plants. The actual origin of the land plant lineage is difficult to interpret. A recent molecular clock study (Smith at al. 2010) suggested an ‘origin of land plants’ around ~477 MY, but this dating in fact refers to the split between bryophytes and the remaining lineages, not to the (presumably earlier) origin of the land plant lineage itself. Extant bryophytes have maintained a ‘semi-aquatic lifestyle’ for >450 MY and bryophyte-like plants may have existed during much of the transition from the aquatic to land habitat. It may well be possible that the poor fossil record and a restricted global occurrence of the first land plants hide 50 or

Evolution of the ‘Plant-Symbiotic’ Fungal Phylum, Glomeromycota

100 MY (or even more?) of evolution of bryophyte-like first land plants. The fossil record is too sparse to be used as evidence against the theories of Heckman et al. (2001) and Hedges and Kumar (2003). Both, fossil and molecular estimates presently still have limitations, and maybe the truth lies somewhere in between. We leave this point moot, but indicate the distinct implications in Fig. 7.2. The first molecular estimates, for the age of the Glomeromycota, from SSU rDNA of 12 AMF (Simon et al. 1993), assumed land plant origins to have been about 420 MY ago and estimated a radiation of Glomeromycota 460–350MY ago. Later studies pushed back the origins of major organism groups into the Precambrian. Redecker et al. (2000a) dated AMF to about 610 MY ago, and Berbee and Taylor (2001) to about 650 MY ago, with an origin of land plants about 600 MY ago. The origin of the Fungi was dated to about 970 MY ago in the latter study. Published protein sequence based estimates (Heckman et al. 2001; Hedges and Kumar 2003) dated the origin of the Fungi to 1400 My ago and that of AMF to about 1200 MY ago. These numbers appear very high, however, in the same study the split in between chlorophyte algae and the lineage leading to the land plants was dated to >1000 MY ago and that between the bryophytes and the vascular plant lineage to 450–700 MY ago, with a likely colonisation of land by plants about 600 MY ago. The last numbers are debatable, but an origin considerably earlier than 470 MY is likely (see also Rubinstein 2010).

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V. Glomeromycota and Land Plant Evolution Molecular analyses and the fossil record indicate that the AM association evolved as a symbiosis, facilitating the adaptation of plants to the terrestrial environment (Schu¨ßler 2002). Pirozynski and Malloch (1975) already suggested a mycotrophic origin of land plants. It appears obvious that the study of land plants without considering their symbiotic lifestyle precludes an understanding of their evolution. The key for this understanding might be to also consider non-land plant associations of fungi in the Glomeromycota. Irrespective of calibration points used for molecular clock analyses, colonisation of land by fungi is always dated to have taken place 50–200 MY earlier than the origin of land plants. Therefore, the earliest possible terrestrial glomeromycotan association could have taken place in ecosystems lacking such land plants. This raised the hypothesis that a symbiotic lifestyle of AMF with photoautrophic organisms could have had evolved before AM symbioses itself originated (Schu¨ßler 2002). It is conceivable that the AM symbiosis then evolved by recruiting already existing fungal symbiotic mechanisms, perhaps derived

zygomycetes Ascomycota Basidiomycota Glomerales Diversisporales

II I

Paraglomerales Archaeosporales

800 1200

700 1000

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Fig. 7.2. The evolution of AMF and land plants shown at putative timescales. The dark lines and numbers roughly correspond to the age of the lineage splits as suggested in the more conservative analyses (e.g., Berbee and Taylor 2001), the grey lines to the more extreme values (Heckman et al. 2001). The putative symbiosis partners (cyanobacteria, green algae, bryophyte-like plants) as well as the known AMF hosts (Aglaophyton, gymnosperms, angiosperms) are shown. Fossil Glomus-like (glomoid) spores

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0

(point I; Redecker et al. 2000a) and fossil scutellosporoid and acaulosporoid spores (point II; Dotzler et al. 2006, 2009) are indicated. The tree is assembled from phylogenetic, fossil, and molecular clock data from the sources noted in the text. The branching topology between certain zygomycetes (e.g., Mortierella, Rhizopus), Glomeromycota, and Dikarya is not yet convincingly resolved and, therefore, shown as polytomy

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from associations similar to the Geosiphon–Nostoc symbiosis (Schu¨ßler 2002). A recruitment of existing ‘symbiosis modules’ is also known from much later in land plant evolution inventing completely new features for the plants, namely the ability to fix atmospheric N2 through the root nodule symbioses (RNS) with bacteria. This symbiosis evolved by recruitment of a signalling pathway that regulates AM formation. The result are ‘common sym genes’ (Parniske 2008) essential for intracellular accommodation of symbionts in both, the AM and RNS symbioses. Similar recruitments of, or adaptations to symbiotic mechanisms on the fungal side could have resulted in a ‘sudden’ potential for plants to exploit the land habitat. A rapid diversification of the main AMF lineages could then have followed. Another interesting aspect of the AM is that there must be some very basic mechanisms of plant–microorganism interactions present among the different ‘ranks’ of AM associations. Some AMF (e.g., Claroideoglomus claroideum; synonym Glomus claroideum) can be symbiotic with such widely divergent photoautotrophs as hornworts and vascular plants (Schu¨ßler 2000). The Geosiphon pyriformis fungus, beside its symbiosis with cyanobacteria, perhaps also forms AM with vascular plants and/or bryophytes (Schu¨ßler, unpublished data). Obviously there are the same ancestral and general mechanisms involved in hornworts and flowering plants, contrasting with more recent and more specific plant–pathogen interaction mechanisms. Recently, Wang et al. (2010) showed for three common sym genes that they are functionally conserved within all major plant lineages and that a Medicago truncatula common sym gene mutant (dmi3) could be functionally complemented with the homologous gene of liverworts and hornworts. This demonstrates the highly conserved function and by that also the key role of AM for land plant evolution. An evolution similar to ‘arms races’ and stepwise gene for gene relationships in host–parasite interactions is not possible for the mutualistical and extremely stable AM association. What we should search for, to find the mechanistic keys to the AM associations, are fundamental differences between AM forming vascular plants, lycophytes, liverworts and non-AM-forming plants like Arabidopsis and mosses. The genome data to run such comparisons are available, and it was already indicated that

certain genes required for AM are lacking in mosses (Wang et al. 2010; Zhang et al. 2010). It also must be considered in plant sciences that mechanisms of nutrient acquisition by land plants since their origin co-evolved with the AMF. Relatively few evolutionary advanced plants have secondarily adapted to other modes of nutrient uptake, often to secondary mycorrhiza types such as the ecto-, ericoid-, or orchid-mycorrhizas. Fewer than 5% of land plants have completely lost mycorrhizal associations, and this number might even be much lower if associations of plants currently considered as non-mycorrhizal, for example with certain fungi from the Sebacinales (Yadav et al. 2010), turn out to be mutualistic and therefore mycorrhizas. Still, approximately 80% of land plant species form AM. In the light of this knowledge, the use of Arabidopsis thaliana as a model plant for nutrient uptake studies is somewhat unfortunate. It does not form arbuscular mycorrhiza and therefore is rather atypical for the normal way of plant nutrition. For members of vascular plant families, it is much easier to list those that lack AMF than those that have them (see Smith and Read 2008). Even some of the ‘non-mycorrhizal’ families contain AM hosts, including several aquatic plants (e.g., Miller et al. 1999, Nielsen et al. 2004). AM is also rare or absent from Orchidaceae and Ericales that form symbioses with ascomycetes and basidiomycetes. Ectomycorrhizas (ECM) are formed by gymnosperms (e.g., most, if not all Pinaceae, a few species in the Cupressaceae and Gnetaceae) and a few woody perennial angiosperms in the Fagaceae, Betulaceae, Salicaceae, Juglandaceae, Ulmaceae, Dipterocarpaceae, Sarcolaenaceae, Rosaceae, Caesalpininioidea, Tiliaceae, and Myrtaceae (Strullu-Derrien and Strullu 2007). Most ECM forming families also contain some AM hosts, Eucalyptus and Populus trees form ECM and AM in parallel or succession, and the seedlings of many trees first form an AM before switching to ECM. As an example, members of the Dipterocarpaceae were often considered to be an example of ECM forming tropical trees, but many have AM associations (several species in parallel to ECM). In general, the specialisation of an entire very old group of fungi as obligate symbionts is unexpected. Members of all the other fungal phyla inhabit a wide spectrum of ecological niches and include pathogens and symbionts, as well as free-living saprobes. It is possible that AMF associations with photoautotrophs, during the colonisation of land, affected the Precambrian climate (Heckman et al. 2001). On the other hand, many researchers doubt such an early existence of land plants. It seems certain that the AMF lineage is older than that of land plants, although it is often suggested that molecular analyses and fossil studies indicate that AMF ‘have their origin with the first

Evolution of the ‘Plant-Symbiotic’ Fungal Phylum, Glomeromycota land plants’ (e.g., Rosendahl 2008). This opens evolutionary scenarios for discussion, for example what were the AMF life strategies before land plants evolved? AMF diverged into their present-day main lineages (four orders) very early and the ancestral lineages (Paraglomerales, Archaeosporales) might have evolved before the land plants, their present day symbiotic partners, existed. Possibly Glomeromycota already radiated during their early associations with bryophyte-like plants or even with more ancestral partners (Fig. 7.2). Presently all main AMF lineages form symbioses with all main land plant lineages, perhaps with the exception of the mosses, but these are not yet studied in detail.

A. The Geosiphon–Nostoc Endosymbiosis The Geosiphon–Nostoc symbiosis is the only known fungal endosymbiosis with cyanobacteria and the fungus belongs to the Glomeromycota. Details of ecological aspects, partner recognition, structure of the symbiosis, etc., are described in Schu¨ßler and Kluge (2001) and Schu¨ßler and Wolf (2005). The photoautotrophic partner, Nostoc, broadens the spectrum of glomeromycotan symbionts beyond vascular plants and bryophytes to prokaryotic partners, and raises interesting questions about the origin of the AM. The symbiosis is small and therefore rarely recognized in nature and its cultivation is difficult. Whereas the photobiont (Nostoc punctiforme) can be cultured without its fungal partner, the fungus is an obligate symbiont living together with its cyanobacterial partner on the surface and in the upper layer of humid, nutrient poor soils. When a Geosiphon hypha comes into contact with a specific, symbiosis compatible Nostoc stage, the latter can be incorporated by the hyphal tip. The irregular structure formed thereafter swells and forms a unicellular ‘bladder’, about 1–2 mm long, appearing on the soil surface. Inside a bladder the cyanobacteria are photosynthetically active, supplying the fungus with carbohydrates whilst the Nostoc obtains its inorganic nutrients via the fungus (Schu¨ßler et al. 2008). From current knowledge it can be hypothesised that the Geosiphon–Nostoc endosymbiosis is an AM-like association in which the eukaryotic plant partner is replaced by a photoautotrophic prokaryote. It also shows a symbiotic organisation that perhaps existed as an early evolutionary stage of AM symbioses, when plants had not yet colonised land and cyanobacteria were prominent under the prevailing environmental conditions.

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B. Evolutionary Scenarios for Land Plant Evolution It is most likely that early land plants could not successfully colonise the land habitat and further evolve to larger organisms that efficiently explore atmosphere and soil substrate, without AMF associations. Evolutionary scenarios that conceivably led to the vascular plant AM include that: 1. ‘Precursor AMF’ evolved as saprobes or parasites of the very early land plants, later becoming symbiotic, giving rise to a symbiotically living highly successful plant lineage and since then co-evolving with the land plants (monophyletic origin of the symbiosis after land plants originated). 2. Primitive land plant lineages evolved independently from glomeromycotan fungi and later evolved symbioses with AMF several times (polyphyletic origin of the symbiosis after land plants originated). 3. Glomeromycotan (or their precursor) fungi and green algae already formed symbioses in the (semi-)aquatic environment and colonised land together from the beginning (origin of the AM as green algal symbiosis). 4. Symbioses similar to the Geosiphon-Nostoc association evolved before plants colonised land and the fungal ‘symbiotic mechanisms’ were later recruited by algae or plants to establish AM symbiosis (origin of the fungal symbiotic AM-like mechanisms in fungus–cyanobacteria symbioses). Points (1) and (2) are related and have been considered mechanisms for the origin of AM symbioses, although the hypothesis of Pirozynski and Malloch (1975) already went further, as represented by (3). Presently, the molecular clock data may favour possibilities (3) and (4), implying that neither saprobism nor parasitism are necessary steps in the evolution of the terrestrial AM. As indicated in (3) and (4) it is conceivable that AM-like symbioses existed, before land plants appeared. In relation to point (4), present-day cyanobacteria symbioses are widespread in extremely diverse organisms and one Nostoc clade is especially ‘symbiosis competent’. In the damp or semi-aquatic habitats where ‘ancestral AMF’ perhaps began to colonise land, cyanobacteria would have been the prominent photoautotrophic organisms. Therefore, glomeromycotan fungi could have lived symbiotically with cyanobacteria (Geosiphon-like?) much earlier than in

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AM with land plants (Schu¨ßler 2002; Fig. 7.2). The discovery of a fossil or recent green algal ‘AM symbiosis’, as well as of Geosiphon-like associations would not be too surprising. There does, however, appear to be a dilemma. The molecular data do not exclude that some lineages within the AMF (at least the Paraglomerales and Archaeosporales) might be older than the land plants. If so, how then could it be explained that land plants presently form symbioses with all of the AMF lineages, including the basal ones? One may, therefore, also speculate about a polyphyletic origin of the AM symbiosis. AMF are flexible and switch between very different photoautotrophic partners such as flowering plants and hornworts. Switches from haploid gametophytes to diploid sporophytes and the colonisation of different organs such as thalli, roots, and adventitious roots all are founded on the same, very basic mechanisms driving all these associations – an interesting point when considering functional and regulatory mechanisms that might have led to AM establishment. Such basic mechanisms could perhaps explain a polyphyletic origin of the AM, as the ‘symbiotic competence’ of AMF would have already existed before their radiation into the recent four main lineages (orders).

C. Glomeromycota Endobacteria Co-Evolved with AMF (and Plants) for at Least 400 MY Another interesting feature of AMF is their widespread association with endobacteria. Two types exist. The better characterised is Candidatus Glomeribacter, representing Burkholderia-related Gram-negative endobacteria. These are enclosed by a host membrane, up to now only found in a relatively young clade of the Glomeromycota, the Gigasporaceae, and they can be lost by repeated AMF sub-culturing (Lumini et al. 2007). Gigaspora rosea is the only member of the Gigasporaceae yet reported to be free of endobacteria. The other type of endobacteria, the so-called bacteria-like organisms (BLOs), appears electron microscopically to be Gram-positive (Schu¨ßler et al. 1994). These BLOs are not enclosed by a host membrane and were detected by electron microscopy in most lineages of Glomeromycota. Recently they were also shown to be present in Gigaspora margarita (Kuga et al. 2008), showing that species in the Gigasporaceae may harbour both types of endobacteria. In contrast to the Glomeribacter endobacteria that represent a more recent association, the Gram-positive BLOs turned out to be very old (>400 MY) endosymbionts occurring in three of the (the Paraglomer-

ales were not investigated) main evolutionary AMF lineages studied (Naumann et al. 2010). On this basis, they are expected to play an important, but yet completely unknown functional role in the AMF and the AM symbiosis. Their phylogenetic affiliation shows that the BLOs most likely belong to a Mycoplasma related clade of bacteria, in the Mollicutes. Surprisingly, very variable bacterial SSU rDNA copies are contained within individual AMF spores, perhaps indicating an endosymbiotically evolving bacterial community (Naumann et al. 2010).

VI. Glomeromycota Asexual Evolution A common question is: how do asexual organisms, lacking meiotic recombination, purge their genomes of deleterious mutations? Since AMF are interpreted to be the oldest extant asexual lineage of eukaryotes, they are well suited for study of this question. However, related questions are if the main purpose of sexual recombination necessarily is to purge a genome from deleterious mutations, or if DNA recombination mainly is a mechanism producing variant recombinant progeny containing fitter individuals under changing conditions?

A. The Oldest Known Eukaryotic Asexuals are Thought to be AMF The main sources of genetic variability in eukaryotes are not random point mutations, but DNA recombination and transposition events. Perhaps stable (relatively invariable) genomes are correlated with adaptations to very stable niches. Efficient DNA repair mechanisms and the lack of exposure to radiation are other conceivable stabilising factors. For large populations, such as those of many AMF, mechanisms like the ‘Mullers ratchet’ may not apply as a driver for the accumulation of deleterious mutations. Similar factors have been discussed as related to the longlasting asexual evolution of bdelloid rotifers (Check-Hayden 2008). Ekelund and Rønn (2008) entitled their discussion of that publication: ‘If you don’t need change, maybe you don’t need sex!’. However life strategies within AMF lineages vary considerably, and the most current opinion regarding their genetics relates to fast growing,


hyphal network forming and massively sporulating species, such as Rh. intraradices-related AMF, not to such species producing fewer offspring, more complex spores and no real hyphal networks, such as many species from the Gigasporacea. The AMF lineages separated many hundreds of MY ago and perhaps germination shields indicate that certain AMF lineages bear some kind of (cryptic) sexuality. Such cryptic sexual reproduction or mitotic recombination could play a role in the purging of genomes. Heitmann (2006) hypothesised that limiting sex enables pathogenic microbes to generate clonal populations after adaptation to host niches. Under this hypothesis, they must however retain their ability to reproduce sexually to enable responses to changes, such as in host defence evolution. This theory is compatible with the ‘red queen’ hypothesis (Hamilton 2001) which relates the costly sexual reproduction with the interaction with parasites. It is considered that the host needs efficient mechanisms to vary its genotype to escape rapidly reproducing parasites that can evolve quickly. Genetic variability (sexual recombination) would mainly be needed as an advantage in antagonistic coevolutionary interactions, whether between predators and prey, competitors within a species, or hosts and their parasites. The rival theory of Kondrashov (1988) suggests that the rapid influx of deleterious mutations creates the need for sex to purge genomes. But there are many protozoan eukaryotes that have lived asexually for a very long time, and there seem to be sophisticated mechanisms to keep asexual genomes functioning (Ekelund and Rønn 2008). May a logical consequence of the Red Queen hypothesis be that sexual recombination is a disadvantage for a mutualistically coevolving obligate symbiotic partner? Wouldn’t a high recombination rate interfere negatively with the tight regulation of the biotrophic symbiont by its host? This might be reasons why AMF are asexuals. Interestingly, second to AMF, the oldest known asexual fungal lineages are insect symbionts, supposed to have evolved asexually for 20–60 MY (Normark et al. 2003). A similar strategy is also hypothesised for the bdelloid rotifers whose genomes are degenerate tetraploids (Check-Hayden 2008). After earlier chromosome duplication, extra copies of many genes are stored on degenerated separate chromosomes, allowing efficient repair of genetic defects including DNA double strand breaks. In asexual bdelloid rotifers this results in genome variability

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without the need for sex and is reflected in high heterozygousity of 3.5–54.0% compared to 1% in sexually reproducing species. In this context it is interesting to note that the Rhizophagus irregularis (often wrongly named Glomus intraradices) DAOM197198 genome sequencing project has been delayed because of high genetic polymorphisms and problems with sequence assembly. The genomic space turned out to be much larger than the previously estimated 15 megabases. Although polymorphic markers based on ESTs are relatively stable in this species (Mathimaran et al. 2008), other unpublished studies showed relatively high variability also on the level of ESTs and it was indicated that segregation can lead to new isolates with altered allele frequencies and distinct effects on plant growth (Angelard et al. 2010). Maybe the assumption that this fungus is haploid was wrong and there is a situation similar to that in asexual bdelloid rotifers?

B. Are AMF Hetero- or Homokaryotic? This basic question is still a matter of debate but its answer is essential to further uncovering why the apparent asexual AMF are so successful. Several publications hypothesise a heterokaryotic nature of AMF (e.g., Hijri and Sanders 2005), whereas others suggest homokaryosis (Pawlowska and Taylor 2004). This controversy is related to the intraspecific genetic variability and putative asexuality. The intraspecific rDNA variability in AMF can be extremely large (Stockinger et al. 2009, 2010) within an individual, polykaryotic spore, and this can also be interpreted to imply that such a spore might represent a heterokaryotic system. For AMF a (recombination driven) concerted evolution of rDNA repeats is evidently relaxed. LSU rDNA regions were shown by in situ hybridisation to be spread over different loci (mostly three or five) within interphase nuclei of several AMF (Trouvelot et al. 1999). In the ascomycetes the rDNA repeats are clustered in one distinct region of a single chromosome, which in the interphase is located at or in the nucleolus (Taga et al. 2003). One may speculate that the different loci in AMF are correlated with the different (main) sequence variants. Between these loci there may be suppressed recombination, for example, by inversion, spatial organisation, or both. Relaxed concerted evolution of rDNA repeats is not too

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unusual. For example, Plasmodium falciparum (the malaria agent) is equipped with ‘mosaic ribosome sets’ and shows different rRNA alleles with sequence identities down to only 80% (MercereauPuijalon et al. 2002). Moreover, pseudogenes as a diversity reservoir and in genome dynamics by promoting ectopic recombination were discussed for Plasmodium. An asexual life strategy may be linked with several different mechanisms leading to genetic recombination. For Giardia intestinales (a dikaryotic parasite) nuclear fusion and genetic exchange including plasmid shuffling between the nuclei was shown to be frequent during encystation (Poxleitner et al. 2008). Similar mechanisms could exist in coenocytic Glomeromycota, keeping their nuclear genomes homogenous. Although much is still based on speculation, AMF clearly offer a fascinating and unique system for the study of asexuality related phenomena. There is some difficulty in defining the terms ‘genetically different’ or ‘genetic variability’, when relating to species or populations. For AMF, it is often not even known whether one deals with one or closely related distinct species. This is exemplified for the model fungus DAOM197198, usually misnamed G.intraradices but in fact not conspecific with Rh. intraradices (synonym G. intraradices) (Stockinger et al. 2009). Probably, AMF are heterokaryotic and the nuclei indeed are genetically diverse, but where is the threshold to being exceptional? Within a human population at least 1% single nucleotide polymorphisms (SNP) are present (would this be what we expect for a population of nuclei in one AMF spore?), but human individuals with ancestry in Europe, Africa or Asia show a ‘genetic difference’ of up to 12% made up by deletions, insertions, duplications and complex multi-site variants (Redon et al. 2006).

VII. Species Concepts, Systematics and Taxonomy Species are a fundamental unit in biology. In general, distinct concepts may be used for species recognition of animals, plants, fungi, and prokaryotes. Different criteria are used among different groups at most, if not all, levels. In sexual species, reproductive isolation is usually believed to play a role and restricted gene flow leads to divergence and then often reproductive incompatibility, or also vice versa. It is difficult to explain speciation in asexual organisms, since here the equilibrating effect of intraspecific interbreeding is presumed to

be lacking. How asexual speciation occurs is debated but it can be a result of adaptation to specific niches (Coyne and Orr 1998). Most of the asexual fungi known seem to have originated recently, for example, asexual Penicillium and Aspergillus species have closely related sexual relatives (LoBuglio et al. 1993) and asexuality can be caused by a recent loss of a mating type (as e.g., in Magnaporte oryza; Couch et al. 2005). However, the bdelloid rotifers are likely to be more than 100 MY old and diversify into entities equivalent to species; sex appears not necessary for speciation (Fontaneto et al. 2007). Operational concepts to recognise evolutionary species (Mayden 1997) are morphological, biological, and phylogenetic or a combination of these. An insightful review of asexual species in the root knot nematode (Heterodera sp.) shows an example of apomictic parasitic species that evolved at least 43 MY ago gaining its nutrition from plant roots (Castagnone-Sereno 2006). Much of that discussion might be applicable to the Glomeromycota. If the Glomeromycota does indeed consist of entirely apomictic lineages, its members would comprise by far the oldest known asexual group of eukaryotes, having separated from their closest related fungal lineages more than 600 MY ago. Thus their asexuality is of ancient origin. It seems that even supra-specific taxa have evolved asexually, implying ancient last meiotic events (Rosendahl 2008). How could such old lineages survive without sex? Did all AMF evolve asexually? What kind of species concept can we use for the Glomeromycota? The only credible biological species concept might be one based on anastomosis compatibility groups. The AMF hyphae are aseptate, with innumerable nuclei sharing the same cytoplasm. Species in some AMF groups anastomose to build up extensive hyphal networks. Nuclei from the same or closely related organisms can thus be exchanged and DNA potentially recombined. However, a degree of vegetative incompatibility was reported among F. mosseae cultures from four different continents (Giovannetti et al. 2003). In contrast, data of Rosendahl et al. (2009) indicate that globally distributed F. mosseae isolates are of very low genetic diversity and perhaps dispersed by human activity. However, separating species from anastomosis compatibility groups has limited usefulness because not all taxa form hyphal networks. For example, fungi in the


Gigasporaceae form anastomoses only with their own self, within an individual hypha (‘wound healing’ or ‘hyphal bridging’), and do not build up hyphal networks. This seems to be related to their different life strategy and functional role in the AM, when compared to fungi in the Glomeraceae (Ijdo et al. 2010). Perhaps for certain groups, such as the genus Rhizophagus (containing the former ‘Glomus intraradices clade’), intrahyphal anastomoses might indicate (parasexual) recombination within a species, whereas in the Gigasporaceae other mechanisms could be in place. Although there is no clear evidence for sexuality, germination shields are complex structures (Walker and Sanders 1986; Fig. 7.3) that involve a large investment and a sophisticated developmental program and they merit further investigation as possible sexual, parasexual, or vestigially sexual structures. If members of the Glomeromycota are solely mitotic, a species concept defining species as evolving through reproductive isolation of sexually produced recombinants cannot be applied. Until recently only a morphological concept could be used for AMF. The definition of morpho-species seems in itself inadequate for some AMF species, but as for bdelloid rotifers many morphological species do exist with a global distribution. These species are supported by molecular phylogenetic analyses. The genetic differentiation in populations observed within fields could be the result of niche differentiation in a heterogeneous soil environment (Rosendahl 2008) and it was reported that haplotypes coexisting in one agricultural field have evolved clonally without recombination (Stukenbrock and Rosendahl 2005). However, the mutation rates suggested that the diversification of the haplotypes exceeds the age of the study country, Denmark, which was under deep ice cover during the Pleistocene, and so however the studied AMF have evolved, they must have invaded post-glaciation. Diversification must therefore have been in an earlier, different environment.

A. Molecular Characterisation of Species In ‘modern’ AMF descriptions, molecular phylogenetic and morphological data were combined to define and recognise taxa (e.g., Gamper et al. 2009). Sequence information is particular valuable for delimiting taxa with few available morphological characters and is necessary for placing them in their appropriate taxonomic hierarchy. For example, many glomoid spore formers previously could not be separated morphologically but such morphotypes occur in all four orders of the phylum. Acaulosporoid spores also occur in species

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Fig. 7.3. Germination shields in Scutellospora and Acaulospora species. These structures are produced as a precursor to germination and are formed from the innermost wall component (or ‘wall layer’). They vary in complexity from simple coil-shaped, through bilobed, to complex shield-like with infolding that may have much thickened edges. Germ tubes emerge as either one from the termination when coil-shaped, or one or more from the lobes when more complex. (A) S. projecturata (coiled): the coil begins centrally and germination takes place at the distal point of the coil (arrowed). In this specimen, the coil has folded back on itself and produced a short branch. (B) A. delicata (coiled): the germ tube can be seen emerging through the spore wall (arrowed). (C) S. heterogama (bilobed): a germ tube initial in the lower lobe of the ‘shield’ (arrowed). (D) S. reticulata (complex, thickened infolds): a germ tube initial can be seen at upper left, near the palisade of thickened folds (arrowed). (E) S. spinosissima (complex, thin infolds): the ‘hole’ through to the spore contents is upper, central, and a germ tube initial can be seen (slightly out of focus, arrowed). F A. capsicula (complex, thin infolds): only part of the large germination shield (bounded by arrows) can be seen and no germ tube initial is evident in this specimen

among different orders. Study of rDNA therefore has fundamentally improved taxonomic concepts in the Glomeromycota. The SSU rDNA dataset, presently the most comprehensive for AMF, also provides a means for distinguishing different genera and higher taxa (Fig. 7.1) in the field. But many phylotypes are not linked with identified organisms, and thus cannot be linked with species. It is not clear at which level any single gene region can be used to define ‘phylogenetic species’. The phylogenetic species concept of Taylor et al. (2000) implies a definition

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based on gene concordances, as different DNA regions may be subjected to distinct mutation rates and selection pressure. There are, however, no such datasets for most AMF species. But, in a multi-gene sequencing approach Sokolski et al. (2010) confirmed the ITS and LSU rDNA based conclusion (Stockinger et al. 2009) that DAOM198197 is conspecific with Rh. irregularis, and not with Rh. intraradices as originally proposed. Moreover, the question of what genes are appropriate is not answered simply. Such a gene must, for example, be one not subject to horizontal gene transfer. Ribosomal DNA gene regions are well defined, conserved in function, and probably do not undergo horizontal gene transfer because they have to interact within too complex a network to allow their replacement. Empirical data (Stockinger et al. 2010) also show that we can apply a ‘molecular species concept’ for AMF based on the rDNA regions. According to the Botanical Code it is legally acceptable to define a new AMF species from just a few dead spores recovered from soil. In some instances, this may actually be useful, where the organisms are so distinctive that they clearly belong to a new taxon, but in many cases where there is little difference between the ‘new species’ and similar, already named specimens, it can be unhelpful and confusing. Usually, a novel species with solely glomoid spores, but lacking any molecular evidence, would have been described in the genus Glomus, pending further study. Hence, this particular genus has become something of a temporary storage for species awaiting correct placement. This strategy may now have to be reconsidered because of the recent (Schu¨ßler and Walker 2010) split of the Glomerales into two families and several genera, based mainly on SSU molecular phylogeny. Combining SSU, ITS and LSU data also is promising for species definition and recognition (Stockinger et al. 2010) because it covers the region(s) likely becoming the DNA barcode for fungi. The present DNA barcoding concept may eventually become part of a more comprehensive phylogenetic species concept, based on several unlinked marker genes.

B. Molecular Phylogeny – from Phylum Definition to Species Recognition in the Field The systematics of AMF was dynamic because unexpected and new phylogenetic relationships were uncovered frequently (Redecker et al. 2007; Walker et al. 2007; Stockinger et al. 2009; Schu¨ßler et al. 2011). There are websites that may help with AMF identification. The INVAM (International Culture Collection of [Vesicular] Arbuscular Mycorrhizal Fungi) and BEG (European Bank of

Glomeromycota) are international sources of well documented germ plasm from pot cultures, and GINCO (Glomeromycota in vitro culture collection) provides some AMF species grown monoxenically in root organ cultures (ROC). One should be aware that cultures may not be correctly identified. This is not necessarily a criticism of the identifications, but may just be a matter of normal progress in the taxonomy of any group of organisms. The use of registered cultures from collections allows comparisons among experiments in which they are used and consequently maintains validity even if nomenclature changes occur. Even for such cultures, vouchers for future verification are essential (Agerer et al. 2000). Although the number of described species in the Glomeromycota is low (~230; see www.amfphylogeny.com) and only a small proportion of the actual species richness, even fewer are represented in the molecular database. At the species level, SSU rDNA exceeds its limits of phylogenetic resolution. In Ambispora (a genus in the Archaeosporales) at least three species (Ambispora leptoticha, Am. callosa, Am. gerdemannii) are ‘hidden’ within the resolution provided even by near full length SSU rDNA phylotypes (Walker et al. 2007). Similarly, environmental sequences determined and annotated in the database as belonging to ‘Gl. versiforme’, are from distinct species (Schu¨ßler et al. 2011). The rDNA regions are also used for molecular detection in ecological studies, and for recognising undescribed or unknown species. But such interpretation often relies on an accurate and sufficiently large dataset of correctly identified species. Molecular characterisation and subsequent detection of AMF species must be founded on knowledge of intraspecific variability and use of sufficiently variable regions, such as the ITS and LSU rDNA region (Stockinger et al. 2010). Other sensitive markers, for example, microsatellite (Mathimaran et al. 2008) or mitochondrial markers (Croll et al. 2008; Bo¨rstler et al. 2010) for species or isolate recognition are being developed, but are presently applicable only for narrow phylogenetic clades such as the one containing Rh. irregulare and close relatives. Many SSU rDNA sequences and phylotypes used in ecological studies are only about 30–50% of full length, so many species may be concealed among such short sequence fragments. Even full-length SSU sequences do

Evolution of the ‘Plant-Symbiotic’ Fungal Phylum, Glomeromycota not resolve species. Therefore, much of the existing diversity is invisible or undefined with the methods in current use. For species identification, there are many reasons why comparison with existing descriptions is difficult or impossible. For some species the herbarium specimens are lost, in other cases the material is in such bad condition that morphological features are no longer recognisable. This is often combined with vague descriptions of key characters. As an information source for taxonomic and phylogenetic data we have established a free and webbased collection of the original species descriptions as a ‘primary data’ reference for taxonomy and descriptions (www.amf-phylogeny.com). On these pages the articles containing the original descriptions (protologues) of almost all current AMF species (including many synonyms) can be downloaded, provided the publishers gave copyright clearance.

C. Biogeography and Geology Only when based on a better understanding of glomeromycotan diversity will biogeography in relation to the evolution of species and higher taxa be understood. The many putative AMF species, as yet known only as phylotypes from environmental samples, will remain hidden until the work of linking organisms with molecular evidence is progressed. Because molecular advances have revealed more complex phylogenetic relationships than anticipated, interpretation of biogeographic patterns and speciation seen in their geological evolutionary context is confounded by the difficulty in defining the boundaries of species or phylotype. SSU rRNA gene sequences do not provide a sufficient resolution and some frequently used primers were shown to be selective for several AMF groups but to exclude others (Schu¨ßler et al. 2001; Kru¨ger et al. 2009) or also to amplify non-glomeromycotan sequences (Douhan et al. 2005). Such problems render subsequent use of DGGE and T-RFLP problematic. Only recently have PCR primers specific for ITS-region and LSU rDNA amplification from AMF been developed (Kru¨ger et al. 2009). Another disadvantage of targeting rDNA regions is that the multiple copies of the repeats may vary considerably. Within one individual spore, ITS region variability can range from 6% for Gi. margarita (Lanfranco et al. 1999) to more than 15% for Rh. intraradices and Rh. irregularis (Jansa et al. 2002; Stockinger et al. 2009). Consequently, cloning or

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deep sequencing approaches are necessary to define the intraspecific variability of AMF species (Scheublin et al. 2004; Stockinger et al. 2009, 2010). Sometimes results from molecular diversity studies in which only the LSU rDNA or ITS region has been analysed are published with unwarranted conclusions. For example, it may be written that in a given ecosystem, ‘only four out of 11 phylotypes belong to known species’. Even with the gene regions with the largest species sampling to date, such conclusions are unfounded because sequence data exist only for about half of the known species. Moreover, the resolution of the mostly used SSU rDNA regions does not allow direct comparison. With the ability to detect AMF at the species level and to sort these species into a phylogenetic ‘backbone’, we should soon be able to tackle the interesting questions of how and when the different lineages and species of AMF evolved in the biogeographical and ecological context. Molecular tools have resulted in a set of AMF community studies by sequencing (van Tuinen et al. 1998) or fingerprinting approaches (de Souza et al. 2004; Lekberg et al. 2007). In molecular ecological studies targeting DNA from roots for PCR and sequencing, rRNA genes were used, corresponding to SSU rDNA (Helgason et al. 2002), LSU rDNA (van Tuinen et al. 1998; Kjøller and Rosendahl 2000), or 5.8S and ITS region rDNA (Redecker 2002). Jansa et al. (2002) performed sequencing and SSCP analyses of the ITS rDNA, based on cultures derived from inoculations with single AMF spores from a field site. The results revealed several ITS sequence variants but no inter-generic genetic exchange. Based on SSU rDNA, Husband et al. (2002) showed a clear temporal succession of dominating AMF over three years in a tropical plant (Tetragastis panamensis), based on 18 phylotypes. In another study, Vandenkoornhuyse et al. (2002) found 24 distinctive AMF phylotypes in two co-existing plant species. About two thirds of the phylotypes found in both these, and in T-RFLP studies (Vandenkoornhuyse et al. 2003) showed distinct AMF communities in co-existing grass species. It is moreover clear, that the AMF diversity is affected by plant communities (Johnson et al. 2003), and ¨ pik et al. (2008) described 34 phylotypes in 5 vice versa. O plant species in a boreal herb-rich coniferous forest. For Glomeraceae (phylogenetic Glomus Group A) only, 19 phylotypes were recorded in trees from an Ecuadorian mountain rainforest site (Kottke et al. 2008), however, for many of the studies it is unclear whether the phylotypes may under- or overestimate species richness. The need to link organisms with phylotypes is increasingly evident. Up to 40 species have been reported in sporebased assays at a certain site.

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VIII. Conclusion Sometimes it is stated that AMF are fundamentally different from other fungi, or even other eukaryotes. Perhaps this will be shown to be correct in the future, but up to now evidence contradicting the present understanding of evolution and speciation has not been verified. The study of AMF evolutionary genetics still is in its infancy and the first genome sequencing, launched in 2004, of an AMF species was still not finished in mid 2011. Many important questions remain to be addressed and even fragmented genome sequence data may answer some. Perhaps the most exceptional character of the Glomeromycota is the putative ancient asexuality and it will be very interesting to discover how this is reflected in genome structure, evolution, ecology, and biology, and distinct evolutionary lineages. Understanding the phylogeny has opened, and will further open new fields for evolutionary interpretations and hypotheses. It may relate the evolution of AM-like symbioses among cyanobacteria and ‘lower’ plants to the understanding of the fundamental mechanisms driving the symbiosis. Theories on early evolution of land plants should be considered with an open mind, as origins for many groups of organisms intermittently are back-dated. The recent fossil record of AMF might indicate that less conservative data published might be closer to reality then the conservative point of view. Unfortunately, despite the impact of AMF on terrestrial ecosystems, their treatment in biological research is still rudimentary. Much more basic research is required to guide and inform investigations into the ecological, economical, and general bio-system relevance of the Glomeromycota. Acknowledgements We thank Manuela Kru¨ger for her help with the artwork and for helpful comments on and proofreading of the manuscript.

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8

Fruiting Body Evolution in the Ascomycota: a Molecular Perspective Integrating Lichenized and Non-Lichenized Groups

IMKE SCHMITT1,2

CONTENTS I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Relevance of Ascomatal Characters in the Pre-Molecular Era . . . . . . . . . . . . . . . . . . . . . . . . . III. Early Molecular Studies . . . . . . . . . . . . . . . . . . . . . . . . . . IV. The Current Molecular Phylogeny of the Ascomycota: Distribution of Fruiting Body Traits Across the Classes. . . . . . . . . . . . . . . . . . . . . . . . . A. Three Subphyla of Ascomycota . . . . . . . . . . . . . . B. The Classes in the Crown Group of Ascomycota (Pezizomycotina) . . . . . . . . . . . . 1. Orbiliomycetes and Pezizomycetes . . . . . . . 2. Laboulbeniomycetes. . . . . . . . . . . . . . . . . . . . . . . 3. Lichinomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4. Eurotiomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5. Lecanoromycetes . . . . . . . . . . . . . . . . . . . . . . . . . . 6. Dothideomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . 7. Arthoniomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . 8. Leotiomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9. Sordariomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Patterns of Morphological Evolution . . . . . . . . . . . VI. Beyond Building the Tree: Statistical Tests of Character Evolution . . . . . . . . . . . . . . . . . . . . VII. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

187 188 189 190 191 192 192 192 192 192 193 193 194 194 194 194 196 198 199

I. Introduction The Ascomycota comprises the largest phylum in the kingdom Fungi and accounts for approximately 75% of all described fungal species (Kirk et al. 2001). Members of this group have evolved a variety of lifestyles, such as saprotrophs, animal and plant pathogens, mycoparasites, mycorrhizae, endophytes, arthropod symbionts, and lichens. Lichenization, that is, the formation of stable associations with phototrophic partners (usually 1

Biodiversity and Climate Research Centre (BiK-F), Senckenberganlage 25, 60325 Frankfurt am Main, Germany; e-mail: imke. [email protected] 2 ¨ kologie, Evolution Goethe Universita¨t Frankfurt, Institut fu¨r O und Diversita¨t, Siesmayerstrasse 70, 60323 Frankfurt am Main, Germany

algae and/or cyanobacteria), is a particularly successful nutritional mode, indicated by the fact that nearly half of the Ascomycota are lichenized (Honegger 1991). Phylogenetic relationships between major groups within the phylum and between lichenized and non-lichenized taxa have been obscure for a very long time in traditional systematics, because of the paucity and homoplasy of morphological and anatomical characters (Cain 1972; Malloch 1981). Phenotypes shared by lichenized and non-lichenized fungi in the crown group of the Ascomycota (Pezizomycotina) include filamentous growth and the formation of fruiting bodies (ascomata). Mainly due to the lack of other complex apomorphies, fruiting body traits have historically been fundamental characters for higher-level classification of the Ascomycota. While nineteenth century classifications divided the phylum according to ascoma macromorphology, twentieth century systems concentrated on fruiting body development and ascus structure. The anatomical studies, however, seemed to create more controversies than they resolved, due in part to the difficulty of identifying homologous versus homoplasious structures, and the one-dimensional application of single characters to taxonomy. Further complications were introduced by the fact that taxonomically artificial boundaries between lichenized and non-lichenized fungi were maintained until the middle of the twentieth century. Consequently, supraordinal classifications were avoided until the early 1990s (Eriksson and Hawksworth 1993). Today, molecular phylogenetics for the first time provides the tools to uncover the evolutionary relationships between the major groups of Ascomycota and to fully integrate the lichenized and non-lichenized taxa. Increasingly robust and detailed multigene phylogenies are becoming available and can be used as a framework on which to trace the evolution of morphological and developmental Evolution of Fungi and Fungal-Like Organisms, The Mycota XIV S. Po¨ggeler and J. Wo¨stemeyer (Eds.) © Springer-Verlag Berlin Heidelberg 2011

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characters. In this chapter, I summarize traditional taxonomical treatments of fruiting body characters in the Ascomycota, give an overview of early and recent developments in molecular phylogenetic classification of the phylum, and discuss the necessity of reanalyzing and reinterpreting traditional ascomatal characters in the light of molecular evidence.

II. Relevance of Ascomatal Characters in the Pre-Molecular Era The fruiting body and related structures are the most complex developmental features in the Ascomycota and have therefore been primary characters in traditional higher-level classification. Traits which have had the strongest influence on taxonomy are ascoma type, ascoma ontogeny, and ascus structure. Early classifications used basic ascoma shapes to define major groups within the Ascomycota: Discomycetes with cupshaped, open apothecia (called hysterothecia when elongated), Pyrenomycetes with pearshaped, apically perforated perithecia, and Plectomycetes with spherical, closed cleistothecia (Fries 1821). These macromorphological distinctions were soon regarded as impractical, because intermediate forms existed (von Ho¨hnel 1907), and additional characters came into use. Nannfeldt (1932) discovered two principal types of ascoma development: ascolocular and ascohymenial. Ascohymenial development begins with fertilization and differentiation of generative hyphae, followed by the development of the ascoma. The result is the formation of true apothecia, perithecia or cleistothecia. Ascolocular development begins with the aggregation of somatic hyphae into an ascostroma, followed by the differentiation of the ascostroma into cavities ‘locules’, which are invaded by generative hyphae that finally form asci. The result is the formation of a pseudothecium, that is, a fruiting body which may look apothecioid or perithecioid, but derived from a different ontogeny. The term pseudothecium was coined by Luttrell (1951), who regarded ascoma ontogeny as well as ascus structure as significant characters. He noticed that fungi classified as Pyrenomycetes can have single-walled (unitunicate) or double-walled (bitunicate) asci and that taxa with ascolocular development always have bitunicate asci (Luttrell 1951). Consequently

he formally described the Loculoascomycetes for bitunicate taxa previously included in Pyrenomycetes (Luttrell 1955). In the following decades the classification of Loculoascomycetes differed greatly between authors (for a review, see Lumbsch and Huhndorf 2007), indicating that homoplasious characters were used to circumscribe what was in fact a heterogeneous assemblage of taxa. The instability of the Lutterellian concept was further enhanced by the discovery of bitunicate asci in taxa with ascohymenial ascoma development (Doppelbaur 1960; Henssen and Jahns 1974; Henssen and Thor 1994; Janex-Favre 1970; Parguey-Leduc and Janex-Favre 1981). Following ontogenetic studies of mainly lichenized taxa, additional ascoma ontogenies beyond the ascohymenial and ascolocular type were described (Henssen 1969a, 1970, 1976; Henssen and Jahns 1974; Henssen et al. 1981). Henssen and Jahns (1974) pointed out alternative versions of ascohymenial development, such as angiocarpous, hemiangiocarpous, and gymnocarpous development. Angiocarpous and hemiangiocarpous development start with the appearance of ascogonia with trichogynes. Ascogonia are often spiral shaped, and trichogyne tips protrude from the thallus surface (Fig. 8.1A). These initial stages are followed by the formation of primordia with a central cavity (Fig. 8.1B). This enclosure later differentiates and forms the margin of the mature ascoma. In angiocarpous development the structure remains closed and forms perithecioid fruiting bodies, while in hemiangiocarpous development the enclosure eventually opens. Depending on the extent of the opening, fruiting bodies with this type of development may appear apothecioid, perithecioid, or have an intermediate morphology (Fig. 8.1D–F). In gymnocarpous development, primordia are formed without a central cavity. Instead, the hymenium of the developing fruiting body is exposed from initiation to maturity, and the ascomatal margin develops during a later ontogenetic stage (Fig. 8.1A, C, F). Fruiting bodies derived from this type of development are typically apothecioid (Fig. 8.1F). Ascomata of lichenized and non-lichenized Ascomycota principally have the same architecture. The majority of lichenized fungi form apothecia, but perithecia and pseudothecia are also common. Ascomatal similarities between lichenized and non-lichenized fungi have led progressive nineteenth century taxonomists to

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primary character to distinguish major groups within the Ascomycota. The main ascus types are unitunicate, bitunicate, and prototunicate. Unitunicate and bitunicate asci typically have mechanisms for forcible ascospore discharge, whereas prototunicate asci passively release the ascospores via decay of the ascus wall. Unitunicate asci can be operculate (opening with a cap) or inoperculate.

Fig. 8.1. Schematic diagram of the main ascoma ontogeny types in higher Ascomycota (Leotiomyceta). Fruiting body development always begins with the formation of ascogonia with trichogynes. Ascogonia are often curled up, and trichogyne tips protrude from the thallus surface (A). Angiocarpous and hemiangiocarpous development (A ! B ! D, E, F) are characterized by the development of ascoma initials (ascogonia with trichogynes; A), followed by the formation of primordia with a central cavity (B). This enclosure eventually opens and forms the ascomatal wall of a perithecium (D), apothecium (F), or an intermediate ascoma type (E). Gymnocarpous development (A ! C ! F) also starts with the development of ascogonia with trichogynes (A), followed by the formation of primordia without central cavity (C). The apothecial margin forms independently at a late developmental stage (F). Drawings by Anna Balla (from: Schmitt et al. 2009)

suggest that lichens should be included in the Fungi (Nannfeldt 1932; Vainio 1890). However, it was not until the middle of the twentieth century that lichens formally became part of fungal taxonomy (Santesson 1953). Even after this merger the separation of lichens into pyrenolichens and discolichens, as with the differentiation of fungi into Pyrenomycetes and Discomycetes, continued to obscure the natural relationships (Richardson 1970). Owing to the historical separation of the disciplines, there are only few hypotheses about the relationships between major groups of lichenized and non-lichenized Ascomycota. Following the ideas of Nannfeldt (1932), Henssen and Jahns (1974) suggested that lichenized and non-lichenized Discomycetes were closely related, in fact that the only difference between the non-lichenized Leotiomycetes and the lichenized Lecanoromycetes was the presence of a phototrophic partner. Similarly, studies of ascoma ontogeny also suggested a close relationship between lichenized and nonlichenized Pyrenomycetes (Henssen and Jahns 1974; Janex-Favre 1970). Despite incongruities in the Lutterellian system, ascus structure continued to be used as a

Asci with a double wall layer can be functionally bitunicate, i.e. they show a clear separation of the two wall layers during ascospore discharge (¼ fissitunicate), or they can be functionally unitunicate, and show no such separation (Reynolds 1989; Trail 2007). A large variety of pore designs from simple to complex have been described (Belleme`re 1977). Several ascus types based on ultrastructure, and iodine staining properties of the ascus wall and the ascus apical structures have been distinguished in the Lecanoromycetes, the largest group of lichenized Ascomycota (Belleme`re and Letrouit-Galinou 1987; Chadefaud 1973; Chadefaud et al. 1963; Honegger 1982).

Despite the presence of a substantial amount of variation in this character (Rambold et al. 1994; Thell et al. 1995), ascus types have had a profound impact on classification of the Lecanoromycetes (Hafellner 1984). Practical problems in studying anatomical features, such as ascus dehiscence and ascoma ontogeny include the fact that the structures of interest are not always clearly discernable (Richardson and Morgan-Jones 1964), and that they can be compromised in herbarium material (Baral 1992). As a result of the difficulties in identifying homologous characters across the Ascomycota, the irresolvable controversies about the significance of individual characters and character combinations for higher-level classification, the problems associated with classifying asexual species, and the artificial separation of mycology and lichenology, the era of strictly morphology-based classification culminated in the observation that natural ascomycete classification was not possible above the ordinal level (Eriksson and Hawksworth 1993). This view was challenged when molecular data became available as an additional taxonomical tool to unravel ascomycete evolution.

III. Early Molecular Studies The first molecular phylogenetic studies in the Ascomycota were based on 18S rDNA sequences

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and focused on the evolution of major lineages. Some of the traditional groupings based on ascoma macromorphology were initially confirmed, for example the monophyly of the classes Plectomycetes and Pyrenomycetes (Berbee and Taylor 1992c, 1995; Gargas and Taylor 1995; Spatafora 1995). However, monophyly of the Discomycetes, and monophyly of the Loculoascomycetes were rejected (Gargas and Taylor 1995; Spatafora 1995). Similarly, bitunicate asci were shown to have multiple origins of evolution (Berbee 1996). The paraphyly of Discomycetes, and the basal position of some Discomycetes in relation to perithecioid and cleistothecioid fungi were soon confirmed by phylogenies based on different molecular markers, such as the RNA polymerase II gene (RPB2), which codes for the second largest subunit of the protein (Liu et al. 1999), and 28S rDNA (Lumbsch et al. 2000). Early literature in euascomycete molecular systematics is summarized elsewhere (Lumbsch 2000; Lumbsch et al. 2005b). The number of lichenized species in molecular phylogenies increased after the development of fungal specific primers for the amplification of mycobiont DNA (Do¨ring et al. 2000; Gargas and DePriest 1996; Liu et al. 1999; Zoller et al. 1999). One repeatedly addressed issue was the phylogenetic origin of lichenization. While Gargas et al. (1995) suggested two separate lichenization events in the Ascomycota (in Arthoniales and Lecanorales) using 18S rDNA, Lutzoni et al. (2001), based on a combined dataset of 18S and 28S rDNA, proposed an early evolution of lichenization within the Ascomycota and its subsequent loss in major non-lichenized lineages (e.g., Eurotiomycetes). The latter view found no support in a study based on RPB1, the gene encoding the largest subunit of RNA polymerase II, which placed the majority of lichenized taxa in a sister group relationship to Eurotiomycetes (Liu and Hall 2004), or in a multi-locus study based on four nuclear and mitochondrial markers (Lumbsch et al. 2005b). It is difficult to assess the number of lichenization and delichenization events in fungal evolution because due to early radiations in the Ascomycota the backbone of the phylogeny is not confidently resolved. Furthermore, taxon sampling is insufficient, and new major independent lineages are still being discovered, for example, the lichenized Lichinomycetes (Reeb et al. 2004). However, it appears that lichenization

occurs in almost all of the major ascomycete lineages, and that the greatest radiations of lichenized taxa have taken place in the Lecanoromycetes, Arthoniomycetes, and Eurotiomycetes. Uncertainties in determining the origins of lichenization and the evolution of fruiting bodies in the Ascomycota using molecular phylogenies have two main causes: unsatisfactory taxon sampling and insufficient resolution of the deeper nodes in the phylogeny. These problems are being tackled by increasing the number of species analyzed and by compiling multi-locus phylogenies. Although the assumption has been made that no amount of molecular data would be sufficient to resolve the rapid radiation of euascomycetes (Berbee et al. 2000), more recent studies suggest that each added gene in concatenated analyses is a stepping stone towards obtaining a better supported backbone of ascomycete phylogeny (Lumbsch et al. 2002, 2005b; Lutzoni et al. 2004; Reeb et al. 2004).

IV. The Current Molecular Phylogeny of the Ascomycota: Distribution of Fruiting Body Traits Across the Classes Several landmark studies on higher-level fungal classification have recently been published as outcomes of major sequencing initiatives (James et al. 2006; Spatafora et al. 2006). The main improvements of these studies in comparison to earlier works are the inclusion of a high number of taxa, as well as a high number of characters in the alignment, and a more balanced taxon sampling of all major groups of fungi known to date. As a result, the first comprehensive phylogenetic classification of the Fungi (Hibbett et al. 2007) is now summarizing the current state of knowledge on molecular fungal classification. This classification, crafted by a historically unprecedented number of mycologists and lichenologists, is implemented in major fungal taxonomy resources, such as GenBank Taxonomy Browser (www.ncbi.nlm. nih.gov), Index Fungorum (www.indexfungorum. org), and Myconet (www.fieldmuseum.org/ myconet) and can be viewed as rigorous working hypothesis. In the following discussion I will use this consensus phylogeny, as well as major


recently published ascomycete phylogenies (Liu and Hall 2004; Lumbsch et al. 2005b; Lutzoni et al. 2004; Spatafora et al. 2006) as a framework to discuss fruiting body evolution in the Ascomycota. A simplified tree modified from some of the above-mentioned papers (Hibbett et al. 2007; James et al. 2006; Fig. 8.2) shows the main classes of fruiting body-forming Ascomycota. I will discuss each class separately with respect to the distribution of fruiting body traits outlined in Section II and cite selected papers relevant to ascomatal character evolution in each respective class.

A. Three Subphyla of Ascomycota

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phyletic subphyla: Taphrinomycotina, Saccharomycotina, and Pezizomycotina, the latter including the majority of filamentous, ascoma-forming species. Basal groups within the Ascomycota are the Taphrinomycotina and Saccharomycotina, which include species that are typically single celled or dimorphic and lack fruiting bodies. An exception is the filamentous ascoma-producing genus Neolecta, which belongs to the Taphrinomycotina (Landvik 1996; Landvik et al. 2001). The phylogenetic placement of this taxon indicates that apothecioid ascomata, as well as unitunicate asci with simple apical apparatus and active ascospore discharge constitute ancestral characters within the Ascomycota (Landvik et al. 2003).

Molecular phylogenetic analyses of nuclear and mitochondrial ribosomal RNA and protein coding genes support a monophyletic Ascomycota phylum, which is a sister group to the Basidiomycota (James et al. 2006). The Ascomycota comprises three mono-

Saccharomycotina are phylogenetically placed between the basal Taphrinomycotina and the derived Pezizomycotina, the crown group of the Ascomycota (James et al. 2006, Liu and Hall 2004, Spatafora et al. 2006). They comprise strictly single-celled, budding yeasts, which produce

Fig. 8.2. Phylogeny of the Ascomycota (cartoon), following Hibbett et al. (2007) and James et al. (2006). Symbols indicate distribution of fruiting body types: open circle apothecium, open triangle perithecium, open triangle with black dot pseudothecium, closed circle cleistothecium,

dash fruiting bodies not known. Large symbols indicate the dominant fruiting body type in each class. Presence of lichenized taxa is indicated in parentheses: (LL) class contains predominantly lichenized species, (L) class contains some lichenized species

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naked asci, suggesting that they may have lost their ascomata as filamentous growth became suppressed in favour of yeast forms.

B. The Classes in the Crown Group of Ascomycota (Pezizomycotina)

(Blackwell 1994). Fruiting bodies occurring in this class are perithecia derived from ascohymenial development. They typically contain unitunicate, clavate, early dissolving asci (Kohlmeyer 1973). Members of the Laboulbeniomycetes are not related to other perithecioid fungi, but constitute an independent lineage in the Ascomycota (Weir and Blackwell 2001).

1. Orbiliomycetes and Pezizomycetes Within the Pezizomycotina, the Orbiliomycetes and Pezizomycetes constitute the earliest diverging lineages. Orbiliomycetes feature apothecioid ascomata and truncate ascus apices, which are sometimes reminiscent of Neolecta spp. (Landvik et al. 2003). Most Pezizomycetes form apothecia containing operculate asci with forcible spore discharge, although other ascoma forms exist (Læssøe and Hansen 2007). An example would be the hypogeous, truffle-shaped ascomata containing prototunicate asci with passive spore dispersal, which have multiple evolutionary origins within the Pezizales (Hansen et al. 2001; Hansen and Pfister 2006; Læssøe and Hansen 2007; O’Donnell et al. 1997; Perry et al. 2007). They presumably constitute a derived fruiting body type, which may have evolved as an adaptation to animal grazing or water stress (Bruns et al. 1989; Thiers 1984). Molecular data have also placed cleistothecial species with prototunicate asci in the Pezizomycetes (Hansen et al. 2005). These taxa are closely related to species which have closed apothecia during early ascoma ontogeny and retain an excipular roof over the hymenium while the asci ripen (van Brummelen 1995). Therefore it has been hypothesized that cleistothecioid ascomata in this lineage are derived from ascomata that open late in development; and that loss of the ascoma opening led to relaxed selection for forcible spore discharge, permitting loss of the accompanying morphological traits, such as an operculum and a distinct hymenial layer (Hansen et al. 2005; Malloch 1981).

Sequence data of the group are still rare, and restricted to a single locus (18s rDNA). Thus, the phylogenetic placement of Laboulbeniomycetes within the Ascomycota, and the classification among members of the group await further molecular studies.

3. Lichinomycetes Lichinomycetes only includes lichenized fungi associated with cyanobacteria as primary photobionts. Members of the Lichinomycetes typically have apothecioid, or more rarely perithecioid ascomata derived from ascohymenial, hemiangiocarpous development that can show considerable variation (Henssen 1963; Schultz and Bu¨del 2002). Ascus types in the Lichinomycetes include the prototunicate ascus and the functionally unitunicate lecanoralean (rostrate) ascus. Based on these morphological traits traditional studies suggested a close relationship to the lichenized Lecanoromycetes (Henssen and Jahns 1974). However, molecular studies suggest that Lichinomycetes forms an isolated, possibly early diverging, and lichenized lineage in the Ascomycota (Reeb et al. 2004). Apothecioid, hemiangiocarpous ascomata with unitunicate asci must therefore be the result of convergent evolution in Lecanoromycetes (see Section IV.B.5) and Lichinomycetes. The precise phylogenetic position of Lichinomycetes in the Ascomycota will hopefully be established with improved taxon sampling. 4. Eurotiomycetes

2. Laboulbeniomycetes Most species in Laboulbeniomycetes are obligate arthropod parasites, which have evolved numerous unique morphological features in adaptation to their lifestyle. This is one reason the classification of the group and its phylogenetic affiliations have been obscure for a long period of time

Eurotiomycetes comprises a diverse set of fungi, displaying various lifestyles (saprotrophs, parasites, pathogens, lichens) and ascoma types (apothecia, cleistothecia, perithecia, pseudothecia). Three subclasses are currently distinguished: Chaetothyriomycetidae, Eurotiomycetidae, Mycocaliciomycetidae. The Chaetothyriomycetidae includes lichenized, parasitic, and saprobic


ascomycetes. Members of this group typically have perithecioid ascomata with bitunicate asci and ascolocular development. However, the lichenized orders Pyrenulales and Verrucariales with ascohymenial ascoma ontogeny also belong in Chaetothyriomycetidae (del Prado et al. 2006; Gueidan et al. 2007; Lutzoni et al. 2001, 2004; Miadlikowska and Lutzoni 2004; Reeb et al. 2004; Schmitt et al. 2005), as well as some nonlichenized unitunicate species (Winka 2000). The second subclass Eurotiomycetidae includes most fungi previously recognized as Plectomycetes because of their cleistothecial ascomata and prototunicate asci. The third subclass Mycocaliciomycetidae consists of parasites or commensals on lichens or saprobes. Members of this group have unitunicate asci with active spore dispersal or, more rarely, passive spore dispersal (Geiser et al. 2006; Tibell and Wedin 2000). Eurotiomycetes appears to be a prime example of a monophyletic group containing species with homoplasious morphological and anatomical traits. It is unclear whether there are plesiomorphic phenotypic characters that can be used to circumscribe this group.

5. Lecanoromycetes Lecanoromycetes comprises the bulk of lichenforming fungi. Three subclasses are distinguished: Acarosporomycetidae, Lecanoromycetidae, Ostropomycetidae. The Acarosporomycetidae (Reeb et al. 2004) consists of lichenized taxa, typically having immersed or sessile apothecia formed by ascohymenial, gymnocarpous development. The asci are bitunicate, functionally unitunicate, and generally contain more than 100 simple and colorless ascospores. The Lecanoromycetidae contains strictly lichenized taxa, which typically have apothecia derived from ascohymenial, gymnocarpous development. Exceptions include the genera Lobaria and Peltigera, which have a hemiangiocarpous ascoma ontogeny (Miadlikowska and Lutzoni 2004). The Ostropomycetidae includes some nonlichenized species, which are considered secondarily de-lichenized, as well as the Conotrema–Stictis species complex, a unique ecological adaptation, and the only example of ‘optional lichenization’ known to date (Wedin et al. 2004). Fruiting body types in the Ostropomycetidae include apothecia and perithecia.

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Perithecia in some genera (e.g., Coccotrema, Pertusaria, Thelotrema) were typically interpreted as perithecioid apothecia, because they lack true periphyses, and they are found in species closely related to apothecioid taxa (Henssen and Jahns 1974; Lumbsch et al. 2001b). However, even families which were traditionally not classified in Lecanoromycetes because their fruiting bodies were thought to be ‘true perithecia’, such as Porinaceae, Thelenellaceae, Protothelenellaceae, and Thrombiaceae were placed in Ostropomycetidae by molecular data (Grube et al. 2004; Schmitt et al. 2005). Moreover, Arctomiaceae, which have ascohymenial, gymnocarpous ascoma development (Henssen 1969b; Lumbsch et al. 2005a) that produces apothecioid ascomata are also placed in this subclass (Lumbsch et al. 2005a; Wedin et al. 2005). In conclusion, it is difficult to provide a morphological circumscription of each of the two subclasses in Lecanoromycetes. Ostropomycetidae appears to have predominantly hemiangiocarpous ascoma development resulting in apothecioid to perithecioid fruiting bodies with all stages of intermediates, while Lecanoromycetidae is predominantly gymnocarpous, rarely hemiangiocarpous, and usually produces apothecia.

Asci in the Lecanoromycetes are functionally unitunicate, and exhibit a large number of unique apical structures. These ‘ascus types’ have had a major impact on the classification of the group (Hafellner 1984). They are important clues to understanding the systematic affiliation of taxa, if they are correlated with other traits. However, classification based on ascus type alone is rejected by molecular data (Buschbom and Mueller 2004; Lumbsch et al. 2001a, 2007; Reeb et al. 2004; Wedin et al. 2005). Analyses of ancestral ascus types in the Lecanoromycetes have revealed a large number of state transformations during the evolution of this character (Ekman et al. 2008). 6. Dothideomycetes Dothideomycetes includes mostly non-lichenized fungi characterized by ascolocular ascoma development and bitunicate asci. It accommodates the bulk of taxa formerly classified as Loculoascomycetes (Lumbsch and Huhndorf 2007). During ascoma development, the ascostroma usually becomes a perithecioid pseudothecium. A few lichenized taxa have recently been classified in the Dothideomycetes based on molecular evidence. They include the Arthopyreniaceae (Lumbsch et al. 2005b), the Trypetheliaceae (del Prado et al. 2006), and the vegetatively reproducing species Cystocoleus ebeneus and Racodium rupestre (Muggia et al. 2008). Ascoma ontogeny has not been studied in

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lichenized members of the Dothideomycetes, except in Arthopyrenia, which has been shown to have ascolocular development (Janex-Favre 1970).

7. Arthoniomycetes The mostly lichenized Arthoniomycetes is the sister group to the primarily non-lichenized Dothideomycetes. Members of this lineage exhibit the character combination bitunicate asci and ascohymenial ascoma development and have therefore been suggested to be an intermediate form between ascohymenial and ascolocular fungi (Henssen and Jahns 1974; Henssen and Thor 1994). Molecular data support monophyly of Arthoniomycetes (Lumbsch et al. 2005b), reinforcing the notion that the influential classification systems proposed by Nannfeldt and Luttrell are incongruent. Common ascoma shapes in the Arthoniomycetes include apothecia or hysterothecia, which are elongated, string-shaped apothecia. However, taxa without real fruiting bodies, but only loosely distributed asci are also suggested to belong to the class (Grube 1998). A greater taxon sampling and more molecular data are needed to improve our understanding of phenotypic evolution this group (Tehler and Irestedt 2007).

8. Leotiomycetes Members of this class comprise the majority of non-lichenized apothecioid Ascomycota with inoperculate, unitunicate asci. The close relationship to lichenized apothecioid, inoperculate, unitunicate species (Lecanoromycetes) suggested by (Henssen and Jahns 1974) is not supported by molecular data (Lumbsch et al. 2005b; Spatafora et al. 2006). Instead, several molecular analyses suggest that Leotiomycetes is the sister group of the primarily perithecioid Sordariomycetes (Liu and Hall 2004; Lumbsch et al. 2005b; Spatafora et al. 2006). The circumscription of Leotiomycetes is still in flux, since the lineage is not monophyletic in some analyses (James et al. 2006; Lutzoni et al. 2004; Spatafora et al. 2006), and taxon sampling is still insufficient (Wang et al. 2006). The placement of the powdery mildews (Erysiphales), a group of cleistothecioid plant pathogens in the Leotiomycetes, must therefore await further molecular studies.

9. Sordariomycetes This class contains fungi with diverse ecological adaptations, such as saprobes, coprophiles, endophytes, mycoparasites, pathogens, and insect mutualists (Spatafora et al. 2006). However, lichenized forms are unknown. Most taxa are perithecioid, but cleistothecia have been derived through loss of the ostiolar canal several times (Samuels and Blackwell 2001; Suh and Blackwell 1999). Asci are typically unitunicate with a variety of different apices. Some species have prototunicate asci, which are interpreted as derived characters that have evolved as adaptation to insect and water dispersal, or in connection with evolution of cleistothecia (Blackwell 1994; Spatafora et al. 1998).

V. Patterns of Morphological Evolution Recent phylogenetic studies on higher-level fungal classification provide an emerging picture of monophyletic classes in the Ascomycota (Hibbett et al. 2007). Eleven of the 15 currently accepted classes are fruiting body-forming (Fig. 8.2). Mapping of ascomatal traits onto these molecularly inferred clades reveals a high degree of variability in fruiting body characters. Each class typically has a dominant fruiting body type, ascoma ontogeny, or ascus type, but alternative morphologies often exist (Table 8.1, Fig. 8.2). The distribution of characters over the molecular phylogeny reveals at least two recurring evolutionary themes: multiple derived origins of the prototunicate ascus, and the evolution of closed fruiting body types from more open types. Closed fruiting body types derived from apothecioid ancestors are found in two unrelated euascomycete classes: perithecia in the Lecanoromycetes (Grube et al. 2004; Schmitt et al. 2005), and cleistothecia in the Pezizomycetes (Hansen et al. 2005). In both cases ascoma ontogeny is invoked to explain the unexpected presence of a closed fruiting body within an apothecial lineage. In the lichenized Lecanoromycetes, a strongly hemiangiocarpous development characterized by initially closed ascomata (Fig. 8.1), is typical for species closely related to the perithecioid taxa. Therefore Grube et al. (2004) and Schmitt et al. (2005) suggest that perithecia in this lineage resemble pedomorphic apothecia, that is, fruiting


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Table 8.1. Distribution of ascomatal characters across the classes of fruiting body-forming Ascomycota (classification following Hibbett et al. 2007). Characters in parentheses indicate character states that deviate from the predominant form in the class Class

Ascoma type

Ascoma ontogeny

Ascus type

Neolectomycetes Pezizomycetes

Apothecia Apothecia (cleistothecia)

? Ascohymenial

Orbiliomycetes Laboulbeniomycetes

Apothecia Perithecia

? Ascohymenial

Lichinomycetes

Apothecia (perithecia)

Ascohymenial

Unitunicate/clavate Unitunicate–operculate (prototunicate) Unitunicate/clavate Unitunicate to prototunicate Unitunicate to prototunicate

Eurotiomycetes: Eurotiomycetidae Chaetothyriomycetidae Mycocaliciomycetidae Dothideomycetes Arthoniomycetes Leotiomycetes Sordariomycetes Lecanoromycetes

Cleistothecia Pseudothecia Apothecia Pseudothecia Apothecia Apothecia (cleistothecia) Perithecia (cleistothecia) Apothecia (perithecia)

Ascohymenial Ascolocular, ascohymenial Ascohymenial Ascolocular Ascohymenial Ascohymenial Ascohymenial Ascohymenial

bodies that reach maturity without forming a typical, wide apothecial opening. Similarly, cleistothecia in the non-lichenized Pezizomycota are interpreted as being derived from apothecioid forms and having a special type of development. These species retain an excipular roof over the hymenium until a very late stage in ascoma ontogeny that eventually tears open (Hansen, et al. 2005).

A repeated pattern within the perithecioid Sordariomycetes is the independent evolution of cleistothecial ascomata. Based on morphological and ecological evidence, some pre-molecular studies suggested that certain perithecioid taxa are closely related to cleistothecioid taxa (von Arx 1973). Molecular data have corroborated this hypothesis, showing that cleistothecia originated several times within the perithecioid Sordariomycetes through the loss of the ostiolar canal (Berbee and Taylor 1992a; Rehner and Samuels 1995; Suh and Blackwell 1999). Prototunicate asci with passive spore dispersal occur in several unrelated lineages, such as the lichenized Lichinomycetes, and Lecanoromycetes, as well as the non-lichenized Eurotiomycetidae and Mycocaliciomycetidae (Eurotiomycetes), and some of the non-lichenized Sordariomycetes and Pezizomycetes. In the pre-molecular literature there is some discussion whether the prototuni-

Prototunicate Bitunicate–fissitunicate Prototunicate Bitunicate–fissitunicate Bitunicate–fissitunicate Unitunicate Functionally unitunicate Functionally unitunicate (prototunicate)

cate ascus constitutes an ancestral or a derived character state. Asci with forcible spore discharge in the non-lichenized Pyrenomycetes were either regarded as advanced (Luttrell 1955), or as ancestral (Ingold 1971; Malloch 1981). Similarly, the lichenized prototunicate Lichinaceae (Lichinomycetes) were regarded as an ancestral (Tehler 1995), or derived lineage (Henssen 1994). In both cases molecular data support the view that there have been multiple losses of the active spore dispersal trait, and that prototunicate asci constitute a derived character state (Berbee and Taylor 1992a; Schultz et al. 2001; Schultz and Bu¨del 2003). Prototunicate asci in both subclasses of the Lecanoromycetes (e.g., Calicium, Lecanoromycetidae; Nadvornikia, Ostropomycetidae) also appear to be derived (Lumbsch et al. 2004; Wedin and Tibell 1997). Prototunicate asci can be an adaptation to the evolution of cleistothecia, or particular dispersal ecologies. The evolution of closed ascomata from ancestors with open fruiting bodies is often accompanied by the loss of active spore dispersal and the emergence of prototunicate asci. In these cases prototunicate asci have evolved due to relaxed selection for ascus structures required for active spore dispersal. Examples include the repeated evolution of hypogeous or semi-hypogeous closed ascomata from epigeous apothecial ancestors in the Pezizomycetes (Hansen and Pfister 2006), the evolution of epigeous cleistothecial

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species in the Pezizomycetes (Hansen et al. 2005), and the evolution of cleistothecial species in the perithecioid Sordariomycetes (Berbee and Taylor 1992b). However, prototunicate asci can also be adaptations to particular dispersal ecologies. Perithecioid fruiting bodies with prototunicate asci in the Sordariomycetes, which release ascospores by oozing rather than shooting out, have presumably evolved under selection for a special dispersal agent, such as insects (e.g., Ophiostoma) and/or water (e.g., Corollospora; Blackwell 1994; Spatafora et al. 1998). Lichenized taxa often produce prototunicate asci in stalked or sessile apothecia (e.g., Calicium). In these cases it is less clear which dispersal ecology is associated with prototunicate asci and the passively protruding ascospore masses.

VI. Beyond Building the Tree: Statistical Tests of Character Evolution Mapping non-molecular characters on a molecular phylogeny provides a wealth of information on trait evolution, such as the presence of synapomorphic or homoplasious characters, or the evolutionary origin of a particular trait. However, simply inspecting the phylogenetic tree and the character states of its terminal taxa is not sufficient, if we want to assess character transformations in the course of evolution, infer ancestral fruiting body traits, and determine causal factors in the evolution of ascomata. Such hypotheses about processes of morphological evolution can only be tested using phylogenetic comparative methods, for example ancestral character state reconstruction. This method uses the distribution of traits observed in extant organisms to infer character states at ancestral nodes in the phylogeny (Huelsenbeck and Bollback 2001; Maddison 1995; Pagel 1999). It is becoming increasingly popular to analyze morphological evolution, and lifestyle switches in the Fungi (Binder and Hibbett 2006; Ekman et al. 2008; Gueidan et al. 2007; Hibbett 2004; Hibbett and Binder 2002; Lumbsch et al. 2006; Lutzoni et al. 2001; Miadlikowska and Lutzoni 2004; Reeb et al. 2004; Spatafora et al. 2007). Ancestral character state reconstruction, like all comparative methods, relies on the estimation of models of evolutionary processes. Maximum

parsimony, maximum likelihood, and Bayesian approaches are available to reconstruct ancestral states. Maximum parsimony, for example, assumes equal probabilities of character gains and losses, and minimizes the number of evolutionary events, which lead to the observed character distribution (Maddison and Maddison 2000; Maddison 1995). This approach can be misleading, however, when the probabilities of gains and losses of a character are unequal (Cunningham et al. 1998; Schultz et al. 1996). Maximum likelihood methods circumvent this problem by estimating a model of evolution for a character, and then calculating support for alternative ancestral states under that model (Pagel 1997; Schluter et al. 1997). Each of the above methods can be used on a single tree, or on a Bayesian tree sample. Combining the maximum parsimony or maximum likelihood approach for character tracing with a Bayesian approach for phylogenetic reconstruction adds the advantage of accommodating phylogenetic uncertainty (i.e. uncertainty in tree topology and branch lengths) in the estimate (Huelsenbeck et al. 2000). This method renders results independent of any one particular phylogenetic tree (Lutzoni et al. 2001; Pagel 1997, 1999). However, ancestral state reconstruction itself on each of the trees in the Bayesian tree sample can also be performed using a Bayesian approach. These fully Bayesian approaches take into account not only phylogenetic uncertainty, but also uncertainty in the state reconstruction (Huelsenbeck and Bollback 2001; Pagel and Meade 2006; Pagel et al. 2004). It should be noted that the probabilistic models underlying these methods certainly oversimplify actual morphological processes, such as fruiting body evolution (Hibbett 2007), and that the various methods of state reconstruction potentially produce conflicting or highly unlikely results (Binder and Hibbett 2006; Ekman et al. 2008). However, as pointed out by Hibbett (2007) they are currently the only approaches available that permit statistical tests of hypotheses of morphological evolution, and we should use them – cautiously – as they emerge. We have recently used likelihood and fully Bayesian approaches to study the evolution of fruiting body macromorphology and ascoma ontogeny in the Lecanoromycetes (Fig. 8.3; Schmitt et al. 2009). Our results suggest multiple statistically supported switches between apothecia and perithecia, and between gymnocarpous and


Fig. 8.3. Evolutionary transitions of ascomatal characters in the Lecanoromycetes (lichenized Ascomycota). The apothecioid, gymnocarpous ancestor of Lecanoromycetes (node 20) gave rise to at least two perithecioid groups (nodes 3 and 7). Open circle Apothecium, closed circle perithecium, empty square gymnocarpous ascoma ontogeny,

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filled square angiocarpous ascoma ontogeny, ? insignificant result. Icons behind species names indicate observed characters in extant species. Symbols at nodes in the phylogeny indicate Bayesian ancestral character state reconstructions (from: Schmitt et al. 2009)

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angiocarpous development during the evolution of the group. The ancestor of Lecanoromycetes had apothecia with gymnocarpous development (Fig. 8.3, node 20). Within the Ostropomycetidae there were at least two transitions to perithecia (Fig. 8.3, nodes 3, 7). Interestingly, correlation analyses showed that the evolution of perithecia is linked to the presence of angiocarpous ascoma development (Schmitt et al. 2009). This finding is consistent with the hypothesis of a neotenic origin of perithecia in the Ostropomycetidae (Grube et al. 2004). Clearly, the transition from open to closed fruiting bodies most likely also involves radical changes internal ascoma morphology, such as ascus-type. Since ascus-types were heavily relied on in the taxonomy of lichenized fungi (Hafellner 1984), it is now obvious why certain lichenized clades with perithecioid ascomata (e.g., ‘Microglaena’) were poorly classified in the past. This study emphasizes that understanding the phenotypic evolution of fruiting bodies can only be achieved by considering the functional correlation of characters (Schmitt et al. 2009).

VII. Conclusion The phylogenetic distribution of ascomatal traits across the Ascomycota shows that fruiting body evolution follows a complex pattern, which we are only beginning to understand. Fruiting body types within classes are variable, and we have to look beyond the apothecium, perithecium, cleistothecium, pseudothecium paradigm of ascomycete classification. Evidently, molecular data reject the circumscription of natural groups based on a priori recognition of cardinal characters, such as ascoma type, ascoma ontogeny and ascus structure. This indicates that there is an urgent need for systematists to revisit the classical character sets and study them in the light of new information gained from molecular phylogenetics. We can now invert traditional practices and use phylogenies to pinpoint cases where character transformations have occurred, and to detect and reexamine previously misinterpreted morphological and ontogenetic characters. While fruiting body type and ascus structure apparently show an enormous level of homoplasy at the class level within the Ascomycota, ascoma ontogeny may be a character that can provide

clues to understanding the evolution of fruiting bodies in the Ascomycota. Historically, ascoma ontogeny was recognized as a fundamental character for ascomycete classification (Nannfeldt 1932). However, it plays a surprisingly little role in modern taxonomy. One reason for this probably is the fact that much of the early original work on ascoma development was published in French (Belleme`re 1967; Chadefaud et al. 1968; JanexFavre 1970; Roux et al. 1986), or in German (Esser et al. 1958; Henssen 1963, 1969a; Henssen and Jahns 1974), and was therefore never widely propagated in the mycological community. In contrast, early landmark studies on classification based on ascus structure were published in English (Luttrell 1951, 1955), and found a much broader audience. Also, the classification system based on ascus structure/dehiscence was probably preferred, because it is less technically challenging and time-consuming to study ascus structure than ascoma development. Although the value of ontogenetic characters for higher-level classification has been recognized (Do¨ring and Lumbsch 1998) and confirmed by molecular phylogenies (Lumbsch et al. 2001a, 2005a), only a few recent ontogenetic studies involving ascomycetes exist (Do¨ring et al. 1999; Do¨ring and Wedin 2000; Greif et al. 2004, 2007; Henssen and Lu¨cking 2002; Lumbsch 1999; Lumbsch et al. 2005a). This is unfortunate, because ontogenetic analyses in connection with molecular phylogenies are a powerful tool for interpreting developmental evolution. Furthermore, thorough characterization of the developmental phenotypes is a necessary precursor to genetic studies on ascoma ontogeny. Developmental observations on ascomata have not been accompanied by extensive genetic analyses in most ascomycetes. The molecular mechanisms controlling fruiting body formation have only been studied in a few model organisms, such as Aspergillus nidulans, Neurospora crassa, and Sordaria macrospora, and are still poorly understood (Busch and Braus 2007; Po¨ggeler et al. 2006; see The Mycota, vols I, XV). During ascoma development simple vegetative hyphae differentiate into up to 15 cell types that form the complex structure of the fruiting body (Bistis et al. 2003). This process is influenced by external parameters, such as light, temperature, and nutrient supply, as well as internal factors, such as speciesspecific signalling molecules (Po¨ggeler et al. 2006).


A multitude of functional genes are involved in controlling the regulatory network underlying ascoma development (Engh et al. 2007a, b; Nolting and Po¨ggeler 2006; Nowrousian and Ku¨ck 2006; Nowrousian et al. 2005; Po¨ggeler et al. 2006). To date this information has only partially been analyzed in a phylogenetic framework. For example, sequence analysis of mating type (MAT) loci showed that MAT genes have higher evolutionary rates than non-reproductive genes (Po¨ggeler 1999, Turgeon 1998, Wik et al. 2008; see Chapter 12 in this volume). Estimations of synonymous versus non-synonymous substitutions indicate that the rapid evolution of MAT-genes in heterothallic Neurospora taxa is caused by positive selection, whereas in homothallic taxa it is caused by relaxed selective constraints, suggesting that the evolutionary mechanisms acting on reproductive genes can change after a switch in the reproductive strategy of a fungus occurs (Wik et al. 2008). Phylogenies based on mating type genes also allow speculation about the evolutionary origins of heterothallism and homothallism in the Ascomycota (O’Donnell et al. 2004; Po¨ggeler 1999). It can be expected that additional phylogenetic analyses of genes involved in ascoma development from a variety of diverse and unrelated fungi (beyond the current model organisms and beyond MAT genes) will provide new insights into how developmental mechanisms become modified through evolution (see Chapter 12 in this volume). The current molecular phylogeny of the Ascomycota corroborates multiple independent origins of the lichen symbiotic lifestyle. Due to lacking backbone support it is not possible to pinpoint exactly where gains and losses of this mutualism occurred. However, the closely intertwined evolutionary history of lichenized and non-lichenized taxa shows that we can only comprehend the evolution of shared morphological characters, such as ascomatal traits, if we consider the entirety of the phylum. Perithecioid lichens, for example, are key players in our understanding of fruiting body evolution in the Ascomycota. While true perithecia were once regarded as restricted to the non-lichenized Sordariomycetes, the phylogenetic distribution of perithecioid lichens shows that this fruiting body type has also evolved in the Eurotiomycetes and Lecanoromycetes. Although the integration of lichenized and non-lichenized taxa in phylogenies has recently improved, taxon sampling must still be extended. Furthermore, many interesting life-

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styles with potentially revealing fruiting body morphologies are not yet represented in molecular phylogenies, or not even described or discovered (Hawksworth 1991, 2001). Examples would be ecologically highly specialized species, such as lichenicolous fungi, ‘borderline lichens’ (species having loose affiliations with a phototrophic partner), and possibly endophytes. The phylogenetic placement of these taxa may evidence that we are currently missing a number of fruiting body phenotypes. Acknowledgements The author thanks Thorsten Lumbsch (Chicago) for initially introducing me to this interesting topic and for continuously supporting and encouraging me in my research. I am indebted to Thorsten, Mariette Cole and Steffen Pauls (both St. Paul) for helpful comments on the manuscript. Anna Balla (Chicago) kindly provided the drawings in Fig. 8.1. My studies on fruiting body evolution in lichenized fungi were partly supported by a fellowship of the Deutscher Akademischer Austauschdienst (DAAD).

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Fruiting Body Evolution in the Ascomycota: a Molecular Perspective Perry BA, Hansen K, Pfister DH (2007) A phylogenetic overview of the family Pyronemataceae (Ascomycota, Pezizales). Mycol Res 111:549–571 Po¨ggeler S, Nowrousian M, Ku¨ck U (2006) Fruiting-body development in ascomycetes. In: Kues U, Fischer R (eds) The Mycota. Springer, Berlin Heidelberg New York, pp 325–355 Rambold G, Mayrhofer H, Matzer M (1994) On the ascus types in the Physciaceae (Lecanorales). Plant Syst Evol 192:31–40 Reeb V, Lutzoni F, Roux C (2004) Contribution of RPB2 to multilocus phylogenetic studies of the euascomycetes (Pezizomycotina, Fungi) with special emphasis on the lichen-forming Acarosporaceae and evolution of polyspory. Mol Phylogenet Evol 32:1036–1060 Rehner SA, Samuels GJ (1995) Molecular systematics of the Hypocreales – a teleomorph gene phylogeny and the status of their anamorphs. Can J Bot 73:S816–S823 Reynolds DR (1989) The bitunicate ascus paradigm. Bot Rev 55:1–52 Richardson DHS (1970) Ascus and ascocarp structure in lichens. Lichenologist 4:350–361 Richardson DHS, Morgan-Jones G (1964) Studies on lichen asci. I. The bitunicate type. Lichenologist 2:205–224 Roux C, Belleme`re A, Boissiere JC, Esnault J, Janex-Favre MC, Letrouit-Galinou MA, Wagner J (1986) Les bases de la systematique moderne des lichens. Bull Soc Bot Fr 133:7–40 Samuels GJ, Blackwell M (2001) Pyrenomycetes – fungi with perithecia. In: McLaughlin DJ, McLaughlin EG, Lemke PA (eds) The Mycota, vol 7A. Systematics and evolution. Springer, Berlin Heidelberg New York, pp 221–255 Santesson R (1953) The new systematics of lichenized fungi. In: Osvald H, Aberg E (eds) Proceedings of the seventh botanical congress, Stockholm 1950. Almquist and Wiksell, Stockholm, pp 809–810 Schluter DT, Price T, Mooers Ø, Ludwig D (1997) Likelihood of ancestor states in adaptive radiation. Evolution 51:1699–1711 Schmitt I, Mueller G, Lumbsch HT (2005) Ascoma morphology is homoplaseous and phylogenetically misleading in some pyrenocarpous lichens. Mycologia 97: 362–374 Schmitt I, del Prado R, Grube M, Lumbsch HT (2009) Repeated evolution of closed fruiting bodies is linked to ascoma development in the largest group of lichenized fungi (Lecanoromycetes, Ascomycota). Mol Phylogenet Evol 52:34–44 Schultz M, Arendholz WR, Bu¨del B (2001) Origin and evolution of the lichenized ascomycete order Lichinales: monophyly and systematic relationships inferred from ascus, fruiting body and SSU rDNA evolution. Plant Biol 3:116–123 Schultz M, Bu¨del B (2002) Key to the genera of the Lichinaceae. Lichenologist 34:39–62 Schultz M, Bu¨del B (2003) On the systematic position of the lichen genus Heppia. Lichenologist 35:151–156 Schultz TR, Cocroft RB, Churchill GA (1996) The reconstruction of ancestral character states. Evolution 50: 504–511

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9

Genomic and Comparative Analysis of the Class Dothideomycetes

JAMES K. HANE1,2, ANGELA H. WILLIAMS1, RICHARD P. OLIVER3

CONTENTS I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Significant Phytopathogenic Species . . . . . . . . . . . . . A. Order Pleosporales. . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Order Capnodiales . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Dothideomycetidae of Uncertain Phylogenetic Placement. . . . . . . . . . . . . . . . . . . . . . . . III. Genome Sequencing in the Dothideomycetes . . . A. Phaeosphaeria nodorum . . . . . . . . . . . . . . . . . . . . . . . B. Mycosphaerella graminicola . . . . . . . . . . . . . . . . . . . C. Leptosphaeria maculans . . . . . . . . . . . . . . . . . . . . . . . IV. Comparative Genomics . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Synteny . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Completion of Draft Genome Assemblies Using Mesosyntenic Predictions. . . . . . . . . . . B. Mitochondrial Genomes . . . . . . . . . . . . . . . . . . . . . . . C. Repetitive DNA. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Repeat-Induced Point Mutation . . . . . . . . . . . V. Pathogenicity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Mechanisms of Pathogenicity . . . . . . . . . . . . . . . . . 1. Lateral Gene Transfer . . . . . . . . . . . . . . . . . . . . . . 2. Generation of Diversity via RIP . . . . . . . . . . . VI. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

205 205 205 207 207 207 207 213 213 214 214 217 219 221 222 223 223 224 225 225 226

I. Introduction The class Dothideomycetes (Schoch et al. 2009) is a recently defined taxon within the phylum Ascomycota. It is one of the largest classes within the Ascomycota, with approximately 20000 member species. Its members span a wide spectrum of host interactions and lifestyles, which include pathogens of plants, animals and other fungi, epiphytes, saprobes and lichens. Of these, the group which have the greatest human impact and which have received the vast majority of scientific attention

are phytopathogenic species. These cause major losses to the agriculture and forestry industries. The class Dothideomycetes has replaced the class Loculascomycetes (Luttrell 1955), in which species were grouped based on the morphology of their sexual fruiting bodies. Ascus morphology in the Dothideomycetes is distinct from other fungal taxons in that the asci are double-walled (bitunicate). The fruiting bodies (pseudothecia) also form distinctive cavities or ‘locules’ in which sexual spores (ascospores) are subsequently formed (ascolocular [syn. ascostromatic] development). Since the creation of this class, the taxonomic placements of Dothideomycete genera and species have been refined several times based on molecular phylogenetic data (Schoch et al. 2006). Recent molecular phylogenies predict two major sub-classes within the Dothideomycetes (Schoch et al. 2009; Fig. 9.1). The Pleosporomycetidae is the larger of the two sub-classes and contains species possessing pseudoparaphyses in contrast to Dothideomycetidae species which do not. The Pleosporomycetidae and Dothideomycetidae are each divided again into four orders. The sub-class Pleosporomycetidae contains the orders Pleosporales, Hysteriales, Mytilinidiales and Jahnulales (Boehm et al. 2009). The Pleosporales is the largest order within the Pleosporomycetidae (Zhang et al. 2009). The sub-class Dothideomycetidae contains the orders Dothideales, Capnodiales, Myriangiales and Trypetheliales (Aptroot et al. 2008). Two remaining orders, the Patellariales and Botryosphaeriales, have not been assigned to either sub-class.

II. Significant Phytopathogenic Species 1

Faculty of Health Sciences, Murdoch University, Perth, WA 6150, Australia 2 CSIRO Plant Industry, CELS Floreat, Perth, WA 6014, Australia 3 Department of Environment and Agriculture, Curtin University, Perth, WA 6102, Australia; e-mail: [email protected]

A. Order Pleosporales Phaeosphaeria nodorum (anamorph Stagonospora nodorum, syn. Leptosphaeria nodorum, syn. Evolution of Fungi and Fungal-Like Organisms, The Mycota XIV S. Po¨ggeler and J. Wo¨stemeyer (Eds.) © Springer-Verlag Berlin Heidelberg 2011

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Fig. 9.1. Cladogram of significant phytopathogens within the class Dothideomycetes and their phylogenetic relationships with various classes of the Ascomycota (adapted from Schoch et al. 2009)

Septoria nodorum) is the causal agent of Stagonospora nodorum blotch (SNB) and glume blotch in wheat. P. nodorum is widely distributed (Solomon et al. 2006) and can cause up to 31% yield losses (Bhathal et al. 2003). Within Western Australia it is the primary wheat pathogen and is estimated to cause AUD 108 million in annual yield losses (Murray and Brennan 2009). Pyrenophora tritici-repentis (anamorph Drechslera tritici-repentis, syn. Helminthosporium tritici-repentis) is the causal agent of tan spot (syn. yellow spot) and is a globally spread wheat pathogen. P. tritici-repentis emerged relatively recently as a major pathogen of wheat and has since increased rapidly in severity, incidence and distribution. The sudden increase in virulence of P. tritici-repentis is believed to have occurred after a lateral gene transfer event with S. nodorum (Friesen et al. 2006).

Leptosphaeria maculans (anamorph Phoma lingam) is the causal agent of blackleg (syn. stem canker) of oilseed rape (syn. canola) and other crucifers. L. maculans is rapidly adaptable to fungicide treatment and has recently overcome Rlm1 resistance (using ‘Surpass 400’) in Australia (Li and Cowling 2003; Rouxel et al. 2003) and also Rlm6 resistance under experimental conditions in France (Brun et al. 2000). Like P. tritici-repentis, the incidence and severity of L. maculans infection has increased in recent years, particularly in Eastern Europe, replacing the endemic but less virulent L. biglobosa (Fitt et al. 2006). Cochliobolus heterostrophus (anamorph Bipolaris maydis), the causal agent of southern corn leaf blight (SCLB) of maize, is widespread in warm and humid areas (Smith and White 1988). C. heterostrophus has three distinct races (Hooker et al. 1970): the endemic race O, race C which


is currently restricted to China and infects C male-sterile cytoplasm cultivars (Wei et al. 1988), and the highly virulent race T which infects Texas male-sterile cytoplasm cultivars (Levings and Siedow 1992). An epidemic of race T during the 1970s in North America caused a 15% reduction in yield (Ullstrup 1972). The sister species C. victoriae, an oat pathogen, was also responsible for an epidemic of Victoria oat blight during the 1940s in the United States (Curtis and Wolpert 2004). The genus Alternaria contains several important phytopathogenic species (Andrew et al. 2009) which also impact upon human health due to mycotoxin contamination of food supplies and the allergenic properties of their airborne spores (Thomma 2003). A. alternata causes leaf spot on a wide range of plant species and produces a vast array of mycotoxins and phytotoxins (Thomma 2003). A. brassicicola and A. brassicae infect most Brassicae species (cabbages syn. mustards), causing brassica dark leaf spot and grey leaf spot respectively. Corynespora cassiicoli is the causal agent of target spot (syn. leaf fall disease). It is a cosmopolitan pathogen with a wide host range including tomato, cucumber, cotton, soybean, tobacco, cocoa and cowpea (Silva et al. 2000). C. cassiicoli most significantly impacts upon rubber tree plantations in the south-east Asian tropics (Hashim 1998). B. Order Capnodiales Mycosphaerella graminicola (anamorph, Septoria tritici) is the most economically significant pathogen of wheat in Western Europe (Palmer and Skinner 2002), causing Septoria tritici blotch (STB, syn. Septoria leaf blotch, syn. speckled blotch, syn. leaf spot). In the last century, STB has replaced SNB as the major pathogen of wheat in this region due to changes in agricultural practices, air pollution, pathogen control strategies and climactic factors (Hardwick et al. 2001; Bearchell et al. 2005). The genus Mycosphaerella also contains M. fijiensis (anamorph Pseudocercospora fijiensis), which causes black leaf streak disease (syn. black sigatoka) in banana and is widespread in all banana growing regions of the world (Ploetz 2001). The genus Cercospora contains several pathogenic species infecting a wide variety of fruits, vegetables and ornamentals. Cercospora beticola, the causal agent of Cercospora leaf spot, is the

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most economically significant pathogen of sugar beet worldwide (Groenewald et al. 2005). Passalora fulva (syn. Cladosporium fulvum) causes tomato leaf mould. P. fulva is a highly refined model for gene for gene host–pathogen interactions in which several pathogenicity effectors have been characterised (Oliver 1992; Wulff et al. 2009). C. Dothideomycetidae of Uncertain Phylogenetic Placement Venturia inaequalis is the causal agent of apple scab and is geographically widespread in applegrowing regions (Gladieux et al. 2008) and is rapidly adaptable to resistant cultivars and fungicide treatments.

III. Genome Sequencing in the Dothideomycetes The class Dothideomycetes contains many agriculturally and economically significant species, yet whole genome sequencing within this class was undertaken relatively late compared with other classes of fungi (Fig. 9.2). At the time of writing, only three Dothideomycete genomes have been sequenced and have had a genome analysis published. The first publically released Dothideomycete genome was that of P. nodorum in 2005 (Table 9.1), which was followed by the publication of its genome analysis in 2007 (Hane et al. 2007). The genomes of M. graminicola and L. maculans were released into the public domain in 2007 and 2010 respectively and genome analyses of both were published in 2011 (Table 9.1; Rouxel et al. 2011; Goodwin et al. 2011). The genomes of C. heterostrophus, A. brassicicola and M. fijiensis are also available although publication of their respective genome analyses is still pending (Table 9.1). A. Phaeosphaeria nodorum The P. nodorum genome (Tables 9.1, 9.2) was thefirst species in the class Dothideomycete to be publicly released, analysed and published (Hane et al. 2007). It was sequenced and assembled in 2005 by the Broad Institute (www.broad.mit.edu) in conjunction with the Australian Centre for Necrotrophic Fungal Pathogens (ACNFP), producing 109 scaffold sequences.

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Fig. 9.2. The number of fungal species with publically available whole genome sequences, grouped by class, submitted to the NCBI Genome database over time (http://www.ncbi.nlm.nih.gov/genomes). The column >2010 indicates species for which genome projects

are scheduled for upcoming release or proposed for sequencing. The class Dothideomycetes (black), although containing several significant species, is relatively underrepresented by whole-genome sequencing compared to most classes of Fungi

The Broad also performed an in silico gene prediction which produced 16597 hypothetical gene models (Table 9.3; version 1). This gene dataset has been substantially revised and is now obsolete (see below).

sion 2 dataset. The version 2 assembly has 107 scaffolds instead of the 109 in version 1. Two scaffolds were removed from the nuclear assembly and combined to form a circular mitochondrial genome of 49.8 kb (Hane et al. 2007). Version 2 also included minor assembly revisions based on the discovery of sequencing and assembly errors after the re-sequencing of various genomic regions. Fourteen to 19 chromosomes had been predicted in P. nodorum by pulsed-field gel electrophoresis (PFGE; Cooley and Caten 1991). Between 19 and 38 sub-telomeric regions were predicted in the version 2 assembly, which was consistent with the PFGE chromosome estimate (Hane et al. 2007); and 98% of ESTs aligned to the genome assembly, indicating that the gene-rich regions of these chromosomes were well represented by these 107 scaffolds (Hane et al. 2007). In a subsequent publication, the gene annotations from version 2 were curated using supporting evidence from both proteomic and proteogenomic experiments (Bringans et al. 2009).

The P. nodorum genome assembly and gene annotations have since been considerably improved by the ACNFP, combining multiple layers of supporting evidence (Table 9.3). The analysis in Hane et al. (2007) defined a revised gene list (version 2) supported by expressed sequence tags (ESTs), manual curation and EST-trained gene prediction. Expressed sequence tags from oleate-grown mycelium (10752 ESTs) and infected wheat libraries (10751 ESTs) were aligned to the genome assembly, supporting 2695 genes. We manually curated genes with EST support and used 795 fully EST-supported genes to train a second round of gene prediction. This produced a reliable set of 10762 predicted genes (Table 9.3; version 2). A total of 5354 gene models from version 1 were unsupported in version 2 and were retained, but did not form part of the ver-

Strain/isolate

ATCC 96836

C5

v23.1.3

CIRAD86

IPO323

Species

Alternaria brassicicola

Cochliobolus heterostrophus

Leptosphaeria maculans

Mycosphaerella fijiensis

M. graminicola

JGI

JGI

JGI

JGI

JGI

COGEME

Genoscope/INRA INRA

Genoscope/INRA

Genoscope/INRA

Genoscope/INRA

JGI JGI

JGI

JGI

JGI

COGEME JGI WUGC

WUGC

WUGC

Organisationa

EST sequencing Miscellaneous sequences Draft whole genome assembly Finished whole genome assembly Whole genome nonassembled reads EST sequencing Miscellaneous sequences Draft whole-genome assembly

Draft whole genome assembly Whole genome nonassembled reads EST sequencing Gene annotation Genome finishing Miscellaneous sequences Draft whole genome assembly Whole genome nonassembled reads Transcriptome sequencing EST sequencing Gene annotation Miscellaneous sequences Draft whole genome assembly Finished whole genome assembly Whole genome nonassembled reads Genome finishing Genome analysis

Project/data type

Sanger sequencing

Sanger sequencing

Sanger sequencing

Genetic mapping

Sanger sequencing

Genetic mapping

Sanger sequencing

Sanger sequencing

Sanger sequencing

Sanger sequencing

Roche 454 GS-FLX

Sanger sequencing

Sanger sequencing

BAC physical mapping

Sanger sequencing

Sanger sequencing

Sanger sequencing

Method

Completed 2007 Ongoing Complete

Completed 2007

Completed 2010

Ongoing Completed 2007

Completed 2010 Published 2011

Completed 2007

Completed 2010

Completed 2008 Ongoing Completed 2007

Completed 2007 Completed 2008 Ongoing Completed 2007

Completed 2006

Completed 2006

Project status

Table 9.1. Online bioinformatic resources for currently available dothideomycete species and future dothideomycete genome projects

continued

NCBI [EST] NCBI [NUC,PRO] JGI

NCBI [Trace Archive]

JGI

N/A Rouxel et al. (2011) NCBI [Genome Project: 63129] COGEME NCBI [NUC/PRO] JGI, NCBI


N/A

NCBI [SRA:SRX014521, SRX014522] NCBI [EST] JGI NCBI [NUC/PRO] N/A

COGEME JGI WUGC NCBI [NUC/PRO] JGI, NCBI [WGS: ACIW00000000] NCBI [Trace Archive]

JGI, NCBI [WGS: ACIW00000000] NCBI [Trace Archive]

Availability

Genomic and Comparative Analysis of the Class Dothideomycetes 209

Genome analysis EST sequencing Transciptome analysis Miscellaneous sequences Draft whole-genome assembly Finished whole-genome assembly Whole-genome nonassembled reads Gene annotation EST sequencing Miscellaneous sequences Genome finishing Draft genome

ACNFP ACNFP/Broad ACNFP

Draft genome Transcriptome

JGI JGI

Broad

Broad

ACNFP

Broad

Genome analysis EST sequencing Miscellaneous sequences Draft whole-genome assembly Mitochondrial genome assembly Whole-genome nonassembled reads Gene annotation

JGI/USDA JGI

JGI

Baudoinia compinacensis Cercospora zeae-maydis

Pt-1C-BFP

Pyrenophora tritici-repentis

Mitochondrial genome assembly Whole genome nonassembled reads Gene annotation

JGI

JGI

SN15

Phaeosphaeria nodorum

Finished whole-genome assembly

JGI

JGI

Project/data type

Organisationa

Aigialus grandis

Strain/isolate

Species

Table 9.1. continued

Completed 2005

Sanger sequencing

Completed 2007 Completed 2007 Sanger sequencing

Complete

Awaiting material

Completed 2007 Awaiting material

JGI, BROAD NCBI [EST] NCBI [NUC/PRO/GSS] Broad

Completed 2007

Sanger sequencing

Optical mapping 454 Titanium and Illumina GAII 454 Titanium and Illumina GAII 454 Titanium and Illumina GAII


Completed 2007

Completed 2007 Completed 2006 Completed 2010 Ongoing Completed 2007

JGI, NCBI [Genome Project: 19047] Hane et al. (2007) NCBI [EST] Ip-cho et al. (2010) NCBI [NUC/PRO] Broad JGI, NCBI [WGS: AAXI00000000] Broad, JGI


JGI NCBI [Genome Project: 19047] Goodwin et al. (2011) JGI NCBI [NUC,PRO] JGI, NCBI [WGS: AAGI00000000] NCBI [NUC:EU053989]


NCBI [NUC:EU090238]

Availability

Optical mapping

Sanger sequencing

Sanger sequencing Microarray

Completed 2005

Sanger sequencing

Completed 2007

Completed 2005

Published 2011 Completed 2007

Completed 2007

Completed 2007

Completed 2007

Project status

Sanger sequencing

Sanger sequencing

Sanger sequencing

Genetic mapping, re-sequencing, mesosynteny Sanger sequencing

Method

210 James K. Hane et al.

JGI JGI BGI-Shenzhen JGI JGI JGI

SCOH1-5

CBS116301

NZE10

STIR04 3.11.1

CBS121621

race 4

DW7 race 5

SO2202

Et28A

C. zeae-maydis

Cryomyces antarticus Dothistroma septosporum Mycosphaerella graminicola Passalora (syn. Cladosporium) herbarum Pyrenophora tritici-repentis P. tritici-repentis

Septoria musiva

Setosphaeria turcica

JGI

Zasmidium cellare

Draft genome

Draft genome

Draft genome

Draft genome

Draft genome

Resequencing

Draft genome

Resequencing

Draft genome

Draft genome

Draft genome

Transcriptome

Draft genome

454 Titanium and Illumina GAII 454 Titanium and Illumina GAII 454 Titanium and Illumina GAII 454 Titanium and Illumina GAII 454 Titanium and Illumina GAII 454 Titanium and Illumina GAII

454 Titanium and Illumina GAII

454 Titanium and Illumina GAII 454 Titanium and Illumina GAII 454 Titanium and Illumina GAII 454 Titanium and Illumina GAII 454 Titanium and Illumina GAII Illumina HiSeq

Awaiting material

Library construction Library construction Awaiting material

Complete

Complete

Awaiting material

Library construction Awaiting material

Library construction Awaiting material

Complete

Awaiting material

a

ACFNP ¼ www.envbio.curtin.edu.au; BGI-Shenzhen ¼ www.genomics.cn; Broad ¼ www.broad.mit.edu; COGEME ¼ cogeme.ex.ac.uk; Genoscope ¼ www.cns.fr/spip; INRA ¼ www. international.inra.fr; JGI ¼ www.jgi.doe.gov; NCBI ¼ www.ncbi.nlm.nih.gov/genomes; USDA ¼ www.usda.gov; WUGC ¼ genome.wustl.edu

JGI

Trypethelium spp.

JGI

JGI

JGI

JGI

303B

JGI

Cenococcum geophilum Cercospora beticola

Genomic and Comparative Analysis of the Class Dothideomycetes 211

212


Table 9.2. Comparison of sequenced dothideomycete genome characteristicsa (adapted from Rouxel et al. 2011). Genome statistics were correct at the time of printing Species

P. nodorum

L. maculans

M. graminicola

M. fijiensis P. triticirepentis

C. A. heterostrophus brassisicola

Strain/isolate

SN15

v23.1.3

IPO323

C5

ATCC 96866

Assembly version Nuclear genome size (Mbp) Assembly coverage Sequencing institution Assembler algorithm Number of scaffoldsb Number of chromosomes or linkage groupsc Number of scaffolds not contained in chromosomes/ linkage groups Assembly N50 Assembly L50 (Mbp) Assembly Lmax (Mbp) Assembly Lmin (Kbp) Number of annotated genes Annotation version Gene density (genes per 10 kb) Average gene length (bp) % repetitive DNA

2 37.21

2 45.1

2 39.7

CIRAD 86 Pt-1C (subculture BFP, race 1) 2 1 74.1 37.8

1 34.9

1 32.0

10 Broad Arachne 107 14–19 (PFGE)

9 Genoscope Arachne 72 17–18 (WGF)

8.9 JGI Jazz 21 21 (WGF)

7.1 JGI Jazz 56 10–23 (GM)

6.9 Broad Arachne 47 11 (OM)

9.9 JGI Arachne 89 15–16 (GM)

6.4 WUGC PCAP 838 9–11 (PFGE) 172 (PM)

107

45

0

Unknown

22

11

Unknown

13 1.0 2.5 2 12194

10 1.8 4.3 0.5 12469

6 2.7 6.1 409.2 10952

5 5.9 8.6 1.01 13107

4d 3.1d 9.5d 6.0d 12169

11 1.3 2.7 1.3 9633

6 2.5 4.2 0.1 10688

3 3.1

2 4.2a

2 3.4

2 1.4

1 3.8

1 3

1 3.9

1326

1323a

1600

1599

1618

1836

1523

6.2

34.2

18.0

Unknown

16.0

7.0

9.0

a

Excludes genes within A:T-rich isochores (see Section II.C). Excludes scaffolds identified to be mitochondrial in origin. c OM ¼ optical mapping, PFGE ¼ pulse field gel electrophoresis, PM ¼ physical mapping, WGF ¼ whole genome finishing (Cooley and Caten 1991; Tzeng et al. 1992; Malkus et al. 2009; Rouxel et al. 2011). d Calculated using artificial chromosome sequences consisting of scaffolds joined by mapping and unmapped scaffolds. b

Table 9.3. Summary of scaffolds, genes and evidence supporting the three versions of the P. nodorum genome assembly Version

1

2

3

Status Availability Number of scaffolds Number of genes Supporting evidence Re-sequencing Transcriptomics: EST Transcriptomics: microarray Proteomics Orthology

Static Broad 109 16597

Static NCBI/JGI 107 (þ1 mitochondrial) 10762

Revision in progress [email protected] 107 (þ1 mitochondrial) 12194

No No No No No

Yes Yes No No No

Yes Yes Yes Yes Yes

Proteomics involves matching the masses and fractionation patterns of semi-purified proteins which have been trypsin-digested and analysed via mass-spectrometry (MS) against a peptide database generated in silico from

a set of known or predicted protein sequences. Proteogenomics uses complex mixtures of proteins, which are then trypsin-digested, separated by two-dimensional liquid chromatography and analysed by tandem mass spectrometry

Genomic and Comparative Analysis of the Class Dothideomycetes (MS/MS). The MS data are then compared to both the predicted proteome and a larger database generated by the wholesale translation of open reading frames from both strands of the entire genome assembly. Both proteogenomic and conventional proteomics techniques can improve gene/protein identification. As draft genome assemblies rely heavily on gene prediction algorithms to determine the proteome, there is a high risk of incorrectly identifying proteins or not detecting them at all if a protein dataset is incorrect. As proteogenomics maps peptides directly to the genome sequence it can bypass any errors introduced by inefficient gene prediction algorithms. This ‘direct to genome’ peptide mapping also makes proteogenomics a powerful tool for genome curation. Proteogenomics can identify errors in start and stop codons and exon–intron boundaries in hypothetical gene models. It can also identify incorrect gene translations caused by frame shifts that have been introduced by either misannotation, genome sequencing errors or assembly errors. Additionally, proteogenomics is capable of detecting protein coding regions of a genome not previously identified by gene prediction software, thus providing data for the annotation of new genes.

Proteomic and proteogenomic analysis of intracellular proteins of P. nodorum supported 1946 version 2 gene models and 188 version 1 gene models (1946 + 188 ¼ 2134; Bringans et al. 2009), which was comparable to that supported by EST sequencing (2695). Of the genes detected by proteogenomics, 62% had not previously been detected by EST sequencing. Several gene annotations were revised after consideration of proteogenomic data, with frame-shift errors detected in 144 genes and exon boundary errors detected in 604 genes. Three new genes were also predicted. The curation of the latest version of the P. nodorum genome, version 3, is still in progress and is currently comprised of 12194 gene models. Version 3 improves upon version 2, incorporating the proteogenomic data summarised in Bringans et al. (2009) as well as additional proteomic, microarray (IpCho 2010) and orthology data (Hane 2011). It is available on request from Professor Richard Oliver ([email protected]) and its use over previous versions is strongly encouraged.

B. Mycosphaerella graminicola The genome of M. graminicola (Tables 9.1, 9.2) was publically released in 2006 (version 1) and revised in 2008 (version 2). The version 2 genome sequence has been curated to a high degree of accuracy and completeness, containing nearly the

213

entire genome from one telomere to the other on all chromosomes (Goodwin et al. 2011). The M. graminicola genome contains 10952 annotated genes on a total of 21 chromosomes. Thirteen of these chromosomes are essential or ‘core’ chromosomes and 8 dispensable chromosomes are non-essential and can be absent in some isolates. These dispensable chromosomes make up 12% of the total genome size, and are distinct from the core chromosomes in being far smaller in size (0.42–0.77 Mb) and having higher repeat contents, lower gene densities and abnormal structure. Over half (54%) of core genes were assigned a functional prediction by Goodwin et al. (2011), compared to 2% of genes in dispensable chromosomes. The majority of dispensable chromosome genes were divergent paralogs of core genes and were highly represented by transcription factors and genes relating to regulation of expression and signal transduction. Dispensable chromosomes were also enriched for microRNAs, containing 21% of predicted microRNAs within the whole genome. Some regions of dispensable chromosomes corresponded to fragments of core chromosome sequences. Goodwin et al. (2011) proposed that the dispensable chromosomes are byproducts of meiotic recombination. However, no significant synteny (see Section III.A) between dispensable chromosomes and the core chromosomes or chromosomes of other species was detected.

C. Leptosphaeria maculans The genome assembly of Leptospaeria maculans v23.1.3 (Tables 9.1, 9.2) was completed in 2007 (Rouxel et al. 2011). The genome comprises 45.1 Mb in 77 scaffolds. Genetic mapping, CHEF hybridisation and bioinformatic predictions (refer to section III.A.1) determined that these scaffolds are contained on 17–18 chromosomes. A total of 42222 ESTs across 15 different libraries were sequenced in three isolates of L. maculans (IBCN18, PL86, v23.1.3). Rouxel et al. (2011) predicted 12 469 genes with coding sequences greater than 100 bp based on the genomic alignment of ESTs and in silico predictions. Genes with coding sequences greater than 300 bp and with supporting evidence from EST alignment or association with conserved domains, a total of 11561 genes, were considered to be reliable gene models.

214


The remaining 908 less reliable gene models were manually revised. The genome is divided into two types of isochores (regions of distinct G:C content), G:Cequilibrated (51% G:C content) and A:T rich isochores which have low (34%) G:C content. The gene density in G:C-equilibrated isochores is high, compared to the heterochromatin-like A:T-rich isochores which have low gene densities and high repetitive DNA content. Putative pathogenicity effectors make up a large proportion (20%) of predicted genes within the A:T-rich isochores. Many of these putative effectors are cysteinerich, small secreted proteins (SSPs) with plant translocation signals. The whole-genome microarray was designed with probes specific to 12396 Eugene-predicted gene models as well as 63 nonpredicted, putative SSP genes and 1316 consensus sequences of EST clusters that did not align to the genome. Gene expression was analysed in 1-weekold culture grown on media and infected oilseed rape sampled at 7 and 14 days post-infection (dpi). Most of the EuGene predictions (84.4%) and unmapped EST clusters (90.8%) were expressed in at least one of these conditions, whereas this applied to only half (51.0%) of the unpredicted SSP genes. Microarray analysis found the expression of SSP genes residing within AT-rich isochores to be up-regulated during infection (at 7 dpi). Gene predictions were also validated by proteomic analysis of mycelial and culture filtrate samples. The majority (83%) of detected proteins in these samples contained a predicted signal peptide sequence. However, only 39 SSPs were detected, of which none resided within AT-rich isochores.

IV. Comparative Genomics A. Synteny Comparing whole genomes can reveal the extent to which species have diverged over time and also highlights regions of synteny. In the context of genomics, synteny is a term used to describe the preservation of the physical order of orthologous genes between two species. Synteny between two genomes can be comprehensibly visualised in the form of a ‘dot-plot’. A dot-plot is a

two-dimensional graph representing matching sequences between two genomes. The sequences belonging to either species are represented by the lengths of the x- and y-axes. Sequence matches are represented by lines drawn between the x- and y-coordinates corresponding to their genomic location in either species. At a whole genome resolution, these lines are usually visualised as dots. Syntenic regions appear as many dots in a linear arrangement hence synteny is also commonly referred to as ‘co-linearity’.

Synteny can be qualitatively distinguished based on the length of the matching region. ‘Macrosynteny’ refers to synteny across large regions which are observable at a whole chromosome scale (Fig. 9.3A). ‘Microsynteny’ refers to synteny at the scale of a handful of genes (Fig. 9.3B). Several cases of macrosynteny have been observed between the genomes of animals (McLysaght et al. 2000; Pennacchio 2003; Kohn et al. 2004) and between those of plants (Cannon et al. 2006; Phan et al. 2007; Shultz et al. 2007). However within Fungi, macrosynteny has only been reported between species within the genus Aspergillus (Galagan et al. 2005; Machida et al. 2005; Pel et al. 2007). Synteny between two species implies either a relatively short length of time since speciation or a selective pressure to retain the physical arrangement of genes. Within the Dothideomycetes, most reported examples of microsyntenic gene clusters are involved in the biosynthesis of secondary metabolites. Leptospaeria maculans has a gene cluster for the synthesis of the polyketide sirodesmin which is syntenic with Aspergillus fumigatus, Chaetomium globosum, Magnaporthe grisea and Fusarium graminearum (Gardiner et al. 2004). L. maculans also has a gliotoxin biosynthesis gene cluster which is syntenic with A. fumigatus (Gardiner and Howlett 2005). Phaeosphaeria nodorum has a putative polyketide biosynthesis cluster that is partially syntenic with Sordaria macrospora (Nowrousian et al. 2010) and a quinate biosynthesis cluster that is well conserved between a wide range of fungi including A. nidulans, M. grisea, Neurospora crassa, Saccharomyces cerevisiae and Schizosaccharomyces pombe (Hane et al. 2007). Nonetheless, in all of the cases above, co-linearity is never precisely conserved and a certain number of genes are rearranged in order or orientation (see Chapter 10 in this volume). Whole genome comparisons of L. maculans (Rouxel et al. 2011) and M. graminicola (Goodwin et al. 2011) with P. nodorum (Hane et al. 2007)


Fig. 9.3. Different types of synteny relationships. (A) Macrosynteny is characterised by co-linear conservation of sequence that can be observed at a chromosomal scale. In the diagram provided, chromosomes A1 and A2 of species A are respectively equivalent to chromosome B1 and B2 of species B. This can be represented in a dot-plot, where the axes represent the length of the sequence and lines within the boxes represent matching sequences. (B) Microsynteny is characterised by co-linear conservation of sequence and order observable at gene level. In the diagram provided, genes A1 and A2 of species A correspond respectively to genes B1 and B2 of species B. This type of relationship is usually too small to be visualised in a dotplot at chromosomal level. (C) Mesosynteny is charac-

215

terised by conservation of sequence between equivalent chromosomes without the conservation of sequence order. In the dot-plot provided, chromosomes A1 and A2 of species A correspond respectively to chromosomes B1 and B2 of species B. The mesosyntenic pattern appears as a scattered ‘block’ of ‘dots’, which represent multiple regions of microsynteny rearranged in order. (D) Mesosynteny can be used to complete genomes which are comprised of partial chromosome sequences, or scaffolds. In the diagram, scaffolds A1 and A2 are incomplete scaffolds which correspond to the complete B1 sequence. By observing the mesosyntenic conservation with B1, shown on the dot-plot, A1 and A2 can be hypothesised to be physically co-located to the same chromosome in species A

216


Fig. 9.4. Mesosyntenic genome finishing in L. maculans (adapted from Rouxel et al. 2011). (A) Mesosyntenic conservation between two scaffolds. Scaffold 4 of L. maculans and scaffold 11 of P. nodorum share multiple regions of homology. Lines drawn between the sequence pair represent regions matched by tblastx with >75% identity over >500 bp length. (B) The six-frame translations of both genomes were compared via MUMMER 3.0 (Kurtz et al. 2004). Scaffolds 0–40 from L. maculans are aligned

along the x-axis and scaffolds 1–109 from P. nodorum are aligned along the y-axis. Dots represent matching regions, similar to the lines in A, between translated scaffold sequences. Presented as a dot-plot, mesosyntenic regions appear as rectangular ‘blocks’ comprised of many dots. Mesosyntenic conservation between scaffold 4 of L. maculans and scaffold 11 of P. nodorum A is retained between this sequence pair almost exclusively, suggesting that these scaffolds correspond to equivalent chromosomes

both exhibit a novel pattern of sequence conservation (Figs. 9.3, 9.4). This conservation pattern is characterised by a distinct lack of co-linearity and extensive rearrangement of short sections of a sequence relative to the compared species

(Fig. 9.4A). This pattern is represented on a dotplot as ‘blocks’ of many ‘dots’ whereby regions homologous to one scaffold are predominantly found in one or a few scaffolds of the other fungus (Fig. 9.4B). We also note that the dots are not


found in diagonal lines, characteristic of macrosynteny. As the dots are mainly genes, the pattern means that genes are conserved on homologous chromosomes but their order and orientation is not conserved. We have coined the term ‘mesosynteny’ to refer to this novel conservation pattern (Rouxel et al. 2011; Goodwin et al. 2011; Hane et al 2011). This pattern degrades with increasing evolutionary distance as indicated by the level of sequence similarity between P. nodorum and L. maculans (Fig. 9.4, Pleosporales vs Pleosporales, 80–100%) compared to that between P. nodorum and M. graminicola (Fig. 9.5; Pleosporales vs. Capnodiales, 60–80%). Similarly, the density of dots within blocks was also observed to become sparser with increasing evolutionary distance. This type of conservation is observed beyond the previously mentioned species and is common between the genomes of Dothideomycete fungi. A whole-genome comparison between the Sordariomycetes Podospora anserina and Neurospora crassa performed by Espange et al. (2008) also reported a mesosyntenic-like pattern. The order of microsyntenic regions was extensively rearranged with respect to the alternate genome (as in Fig. 9.3C). As the genomes of both species had been assembled into whole chromosomes (Galagan et al. 2003; Espagne et al. 2008), Espange et al. (2008) observed that these microsyntenic regions were almost exclusively co-located on ‘equivalent’ chromosomes (i.e., chromosomes derived from the same chromosome of their last common ancestor) and intra-chromosomal rearrangements occurred at a much higher rate than that of translocations between non-sister chromosomes.

1. Completion of Draft Genome Assemblies Using Mesosyntenic Predictions As chromosomal content is retained (albeit rearranged in order and orientation) on equivalent chromosomes between species exhibiting mesosynteny, it can be applied to the process of genome ‘finishing’ (Rouxel et al. 2011; Goodwin et al. 2011). Whole genome sequencing involves the sequencing of many short DNA fragments which are bioinformatically assembled into longer sequences. Unfortunately, repetitive sequences often cannot be assembled unambiguously. Therefore, draft assemblies of fungal genomes are not

217

composed of complete chromosome sequences but of contigs or scaffold sequences which represent partially sequenced regions of chromosomes. Conventional finishing methods include genetic mapping (Rouxel et al. 2011), optical mapping (Schwartz et al. 1993), HAPPY mapping (Dear and Cook 1993) and re-sequencing. Using mesosyntenic predictions to aid genome finishing has certain advantages over these methods. It is applicable to fungal species in which a sexual phase is absent or cannot be induced under laboratory conditions, making genetic mapping impossible. It also has the potential to achieve the same end result as these traditional techniques at lower expense and throughput. ‘Mesosyntenic finishing’ (Fig. 9.3C) of a species of interest (species A) relies on the availability of a whole or draft genome of a related species (species B). All sequence combinations between two genomes are tested for significant proportions of matching sequence. A bioinformatic method which automates this process has been developed by the authors of this article (unpublished data). If two scaffold sequences (A1 and A2) from species A both exhibit mesosyntenic conservation with a single sequence (B1) of species B, then scaffolds A1 and A2 can be hypothesised to be co-located on the same chromosome. The hypothesis can be tested in a number of ways. Scaffold-specific primers can be designed leading outwards from the scaffold termini. PCR amplification can only occur between such primers if the two scaffolds are physically joined thus the presence of an amplicon validates the hypothesis. In addition, sequencing the amplicon can improve the genome assembly by filling in gaps in the sequence. Alternatively, hybridisation techniques such as Southern blotting, binding scaffold-specific probes to fragmented genomic DNA can be employed on a much smaller scale than would be feasible by a ‘blind’ approach. Chromosomal co-location, order and orientation of scaffolds can be determined by hybridization of multiple probes to the same DNA fragment. These techniques contributed to finalising the draft genomes of L. maculans and M. graminicola. Mesosyntenic analysis predicted joins between L. maculans scaffolds, based on comparisons with P. nodorum (Fig. 9.4): 8 and 10, also 20, 21 and 23. It also predicted joins based on comparisons with other Dothideomycetes (data not shown): 2 and 19, 3 and 31, also 6, 11 and 29, also 12, 15 and 32. Mesosyntenic predictions for

218


Fig. 9.5. Mesosyntenic genome finishing in M. graminicola (Goodwin et al. 2011; Hane 2011). Comparisons of M. graminicola genome assembly (y-axis) versions 1 (top) and 2 (bottom) against the genome assembly of P. nodorum (x-axis). Scaffolds/chromosomes are ordered along their respective axes according to both decreasing length and increasing number. The six-frame translations of both genomes were compared via MUMMER 3.0 (Kurtz et al. 2004).

Homologous regions are plotted as dots, which are shaded for percent similarity (right). Mesosyntenic predictions were used to finalise the assembly of scaffolds (Version 1) into whole chromosomes (Version 2) as indicated by the numbered circles. Groups I–III demonstrate instances where M. graminicola scaffolds were joined to form a chromosome and those in group IV and V illustrate where a scaffold was cleaved to form two separate chromosomes

Genomic and Comparative Analysis of the Class Dothideomycetes Table 9.4. Features of dothideomycete mitochondrial genome sequences (adapted from Hane et al. 2007; Torriani et al. 2008; Rouxel et al. 2011) P. nodorum Reference Strain/isolate Size (bp) G:C content Genetic codea tRNA rRNAs Large ribosomal RNA subunit (rnl) Small ribosomal RNA subunit (rns) Mitochondrial proteinencoding genes ATP synthase subunits atp6 atp8 atp9 Cytochrome oxidase subunits cox1 cox2 cox3 Cytochrome b (cytb) Nicotinamide adenine dinucleotide ubiquinone oxidoreductase subunits nad1 nad2 nad3 nad4 nad4L nad5 nad6 5S ribosomal protein (rps5) Unknown ORFs Intronic endonucleases

M. L. graminicola maculans

Hane et al. Torriani (2007) et al. (2008) SN15 IPO323, STBB1 49761 43960 29.4 32 4 4 27 27 2 2 Yes Yes

154863 30 – – – –

Yes

Yes

–

12

14

–

1

3

–

Yes No No 3

Yes Yes Yes 3

– – – –

Yes Yes Yes Yes 7

Yes Yes Yes Yes 7

– – – – –

Yes Yes Yes Yes Yes Yes Yes Yes

Yes Yes Yes Yes Yes Yes Yes No

– – – – – – – –

3 4

8 0

– –

Rouxel et al. (2011) v23.1.3

a

4 indicates the mold, protozoan, and coelenterate mitochondrial code and the Mycoplasma/Spiroplasma code, http://www.ncbi. nlm.nih.gov/taxonomy/utils/wprintgc.cgi

M. graminicola in comparison with P. nodorum predicted all scaffold joins between version 1 scaffolds (scaffolds: 10 and 14, 7 and 17, 12 and 22)

219

and scaffold breaks (scaffolds 4 and 9) which were updated in version 2 (Fig. 9.5). These predictions were verified by genetic mapping, hybridisation and re-sequencing in L.maculans and genetic mapping and re-sequencing in M. graminicola (Rouxel et al. 2011; Goodwin et al. 2011).

B. Mitochondrial Genomes Mitochondrial genome sequences (mtDNAs) are currently available for all three published Dothideomycete genomes (P. nodorum, M. graminicola, L. maculans) however only those of P. nodorum and M. graminicola have been annotated and studied in detail (Hane et al. 2007; Torriani et al. 2008; Rouxel et al. 2011). All three mtDNAs have similarly low G:C contents, ranging from 29.4 to 32.0% (Table 9.4) which are typical of the mitochondrial genomes of filamentous fungi (Torriani et al. 2008). The mitochondrial genomes of P. nodorum and M. graminicola are similar in size but the mtDNA of L. maculans is approximately three times larger (Table 9.4). The L. maculans assembly also has four mitochondrial linear plasmids ranging in size from 4.4 to 7.6 kb which contain polymerase genes (Rouxel et al. 2011). Both P. nodorum and M. graminicola mtDNA encode the large (rnl) and small (rns) subunits of the mitochondrial ribosomal RNA complex and 27 transfer RNAs (tRNAs) which can attach to all 20 amino acids (Hane et al. 2007; Torriani et al. 2008). These tRNAs are contained within five separate clusters in both mtDNAs (Fig. 9.6, Table 9.5). The two largest of these clusters in both species are those flanking the rnl gene. The 5’ flanking tRNA cluster is well conserved, however the 3’ cluster is variable between Dothideomycetes species and also across the Ascomycetes. Some tRNAs in P. nodorum and M. graminicola were atypical. Both had mitochondrial tRNAs for threonine and phenylalanine with nine nucleotides instead of the typical seven in the their anti-codon loops and tRNA-Arg2 of P. nodorum had 11 nucleotides in its anticodon loop.

Fungal mtDNAs typically encode for 12 mitochondrial proteins, which are hydrophobic subunits of respiratory chain complexes (Table 9.4). P. nodorum lacks the atp8 and atp9 genes whereas M. graminicola lacks the rps5 gene. M. graminicola mitochondrial genes differ from those of most fungi in that they lack introns

Fig. 9.6. Published and annotated mitochondrial genomes of the Dothideomycetes Phaeosphaeria nodorum and Mycosphaerella graminicola (adapted from Hane et al. 2007 and Torriani et al. 2008)

220 James K. Hane et al.


221

Table 9.5. Comparison of tRNAa gene clusters flanking the rnl gene in dothideomycete and ascomycete species (adapted from Hane et al. 2007; Torriani et al. 2008) Species

Class

5’ Upstreama,b,c

rnl

3’ Downstreamb,c

Genbank accession

Mycosphaerella graminicola Phaeosphaeria nodorum Aspergillus niger A. tubingensis Epidermophyton floccosum Penicillium marneffei Verticillium dahliae Metarhizium anisopliae Lecanicillium muscarium Fusarium oxysporum Trichoderma reesii Podospora anserina

Dothideomycetes Dothideomycetes Eurotiomycetes Eurotiomycetes Eurotiomycetes Eurotiomycetes Sordariomycetes Sordariomycetes Sordariomycetes Sordariomycetes Sordariomycetes Sordariomycetes

GDS1WIS2P VKGDS1WIRS2P KGDS1WIS2P KGDS1WIS2P KGDS1IWS2P RKG1G2DS1WIS2P KGDS*VW*R*P1*P2 YDS1N*G*LIS2W GVISW*P VISWP ISWP ISP

rnl rnl rnl rnl rnl rnl rnl rnl rnl rnl rnl rnl

M1L1EAFL2YQM2HRM3 TM1M2EAFLQHM3 TEVM1M2L1AFL2QM1H TEVM1M2L1AF2QLM3H TEVM1M2L1AFL2QM3H TEVM1M2L1AFL2QM3H TE1M1M2L1AFL2QHM3 TEM1M2L1AFKL2QHM3 TE1M1M2L1E2FKL2QHM3 TEM1M2L1AFKL2QHM3 TEM1M2L1AFKL2QHM3 TEIM1L1AFL2QHM2

EU090238 EU053989 DQ207726 DQ217399 AY916130 AY347307 DQ351941 AY884128 AF487277 AY945289 AF447590 X55026

a

Asterisks indicate where functional genes interrupt the tRNA gene sequence. The numbers 1, 2, 3 indicate the presence of more tRNA genes for the same amino acid in the consensus sequences. c Capital letters refer to tRNA genes for: R, arginine; K, lysine; G, glycine; D, aspartic acid; S, serine; W, tryptophan; I, isoleucine; P, proline; T, threonine; E, glutamic acid; V, valine; L, leucine; A, alanine; F, phenylalanine; Q, glutamine; H, histidine; Y, tyrosine; N, asparagine. b

and therefore have no intron-encoded proteins. P. nodorum has 4 such intronic genes which encode for GIY-YIG and LAGLIDADG-type endonucleases. Both P. nodorum and M. graminicola have additional open-reading frames of unknown function (3 and 8 respectively). The expression of the ORFs orf5, orf6 and orf8 of M. graminicola has been confirmed by EST sequencing (Torriani et al. 2008). C. Repetitive DNA P. nodorum was initially predicted to have 26 different families of long interspersed repeats, which make up 6.2% of the nuclear genome (Table 9.2; Hane et al. 2007). The number of repeat families has since been revised to 25, of which 17 have been characterised (Hane and Oliver 2008, 2010). A significant proportion of these are non-transposon-derived, instead being highly replicated copies of endogenous genes and gene clusters. These genes include a telomere associated RecQ helicase, ubiquitin conjugating enzyme, Rad5 and Rad6 homologues and several genes of unknown function. These endogenous gene-rich repeats were among the largest and most abundant types of repeats in P. nodorum, on a scale eclipsing most transposable element repeat families. The M. graminicola genome is comparable in size to P. nodorum, (39.7 and 37.1 Mb, respectively) but its repetitive content is much larger at

18% (Table 9.2). The repetitive content is almost doubled within the dispensable chromosomes (30%; see Section II.B) compared to its core chromosomes (15.9%). Details of the types of repeats present in the M. graminicola genome have not been published. The genome of L. maculans is significantly larger (45 Mb) than P. nodorum and M. graminicola (Rouxel et al. 2011). This is due to the higher content of repetitive DNA making up 34% of the whole genome (Table 9.2). Unlike P. nodorum, the most abundant repeat families are class II retroelements, with nine families making up 27.26% of the genome. The next most abundant is class I DNA transposons, with nine repeat families making up 2.64% of the genome. There are 11 other repeat families, including rDNA and short telomeric repeats, which together make up the remaining 4.12%. Rouxel et al. (2011) propose that genome invasion by retrotransposons on such a vast scale caused the formation of distinct regions of consistent G:C content (syn. isochores; see Section II.C). While this data indicates vast diversity in the repeat content of each Dothideomycete genome, each analysis has used different software and detection criteria for the identification of repetitive DNA. P. nodorum repeats were predicted de novo via RepeatScout (Price et al. 2005), requiring each repeat family to have a minimum full length of 200 bp, at least 75% identity and a minimum

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of 10 full-length copies. L. maculans repeats were predicted via the REPET pipeline (Bao and Eddy 2002; Edgar and Myers 2005; Quesneville et al. 2005; Rouxel et al. 2011). Details of the method and criteria for prediction of repeats in M. graminicola have not been published. Due to these differences in methodology these results are not directly comparable. 1. Repeat-Induced Point Mutation The repetitive DNA of many fungal genomes is subject to a genome defence mechanism specific to fungi called a repeat-induced point mutation (RIP). RIP was first observed in Neurospora crassa (Selker et al. 1987) and has since been experimentally demonstrated to occur in many other filamentous fungi (Hane and Oliver 2008). It is likely that RIP protects the genome against transposon replication by introducing mutations in repetitive DNA that are likely to introduce stop codons in transposon genes. Prior to meiosis, RIP converts cytosine bases to thymine within similar regions of DNA of a sufficient length. In N. crassa the requirements for RIP are a shared sequence identity of at least 80% over at least 400 bp (Watters et al. 1999). The base adjacent to the mutated cytosine is biased towards adenine (i.e., CpA dinucleotides become TpA) in most species. The mutation of CpA to TpA has a high probability of introducing TAG or TAA stop codons within the repetitive region on both strands of the affected DNA molecule. RIP can be detected by comparing the transition to transversion ratio (Tn:Tv) of retroelement sequences. Transitions are mutations between two purines bases (i.e., A!G and G!A) or two pyrimidines (i.e., C!T and T!C) whereas transversions are mutations from a purine to pyrimidine, or vice versa (i.e., C!A, A!C, G!T or T!G).

RIP has been observed in L. maculans. After the introduction of multiple copies of an exogenous hygromycin resistance gene and subsequent sexual crossing, RIP-like sequence mutations were detected within resistance gene regions in hygromycinsensitive progeny (Idnurm and Howlett 2003). Whole genome analysis indicated that RIP is highly active in L. maculans, with only 19 out of 42222 EST sequences aligning to repetitive regions and no detectable expression of transposon openreading frames by microarray (Rouxel et al. 2011). RIP was bioinformatically predicted in the whole genome of M. graminicola (Goodwin et al. 2011).

As RIP mutations involve C!T transitions, the Tn:Tv ratio of RIP-affected repeats was expected to be higher than non-RIP-affected regions. The Tn:Tv of selected repetitive regions were reported by Goodwin et al. (2011) to be significantly higher (ranging from 25.3 to 42.5) than that of a non-repetitive control set of mutated sequences (1.0). Furthermore, coding regions from transposons present in high copy number were found to contain numerous stop codons as would be expected after RIP. The repeats of P. nodorum were analysed using recently developed software for the analysis of RIP mutation: RIPCAL (Hane and Oliver 2008). Traditional methods of predicting and quantifying RIP have relied on the use of ratios of di-nucleotide frequencies, or RIP indices. While indices are useful when searching for RIP within single-copy sequences, with the availability of whole-genome sequences it is now more appropriate to use alignment-based methods on all repeats within a repeat family. RIPCAL extracts repeat sequences from a whole genome and uses CLUSTALW to generate multiple alignments of each repeat family. Mutations were compared within each repeat family, quantified and their RIP-like identity determined. As in most species, the dominant form of RIP mutation in P. nodorum was predicted to be CpA!TpA. The high-copy number repeats of P. nodorum were predicted to be affected by RIP to varying degrees. The RIPCAL package also includes a tool to reverse the effects of RIP: deRIP (Hane and Oliver 2010). The deRIP tool identifies RIP-like mutation sites within a repeat family alignment and generates an alignment consensus with modifications that reverse RIP-affected regions to their presumed pre-RIP states. This allows for accurate characterisation of the origin of a RIP-affected sequence via similarity searches. For several repeat families of P. nodorum, BLAST searches did not retrieve hits which could be used to identify the role or origin of that repeat. However after deRIP was applied, the origins of 5 previously uncharacterised repeat families were identified. This increased the proportion of characterised repeats in P. nodorum from 65% to 88%. Intriguingly, while some of these were heavily RIP-degenerated transposable elements, others were endogenous P. nodorum genes or gene clusters (Hane and Oliver 2008; see Section III.C).


V. Pathogenicity A. Mechanisms of Pathogenicity The three published genome analyses of the Dothideomycetes P. nodorum, L. maculans and M. graminicola each describe the mechanisms of pathogenicity in each species (Hane et al. 2007; Rouxel et al. 2011; Goodwin et al. 2011). These closely related fungi have evolved remarkably diverse strategies for survival within their respective hosts. Indeed, a recent Ascomycete-wide genome comparison reported that a significant proportion (8–11%) of the gene content of Dothideomycetes is specific to class or genera (Schoch et al. 2009). While no published work has yet compared the gene contents of these three species in detail, individually each genome analysis highlights the major differences between these species. During wheat infection, P. nodorum expresses genes encoding for a battery of cell wall-degrading enzymes (CWDEs) which act predominantly upon xylan and cellulose and also include enzymes degrading carbohydrates and proteins (Hane et al. 2007; IpCho 2010). Membrane transporter proteins, particularly those of carbohydrates, are also important during infection. During the initial stages of infection (3 dpi), P. nodorum highly expresses genes involved in ribogenesis and protein localisation. This is followed (5 dpi) by high expression of genes involved in nutrient assimilation and catabolism which continues into the late stages of infection (7–10 dpi). P. nodorum infection is inferred by the authors to involve the secretion of pathogenicity effectors and cell wall degrading enzymes into the extracellular space. These induce necrosis and disrupt the cell walls of neighbouring host cells, either causing nutrient leakage or facilitating fungal penetration. Enzymes degrading plant carbohydrates, sugars and proteins are also secreted, producing simple metabolites which are imported into the fungal cell by membrane transport proteins. Studies of the genome content in L. maculans have chiefly focussed on small secreted proteins (SSPs), in an effort to discover and characterise avirulence (AVR) effector genes (Rouxel et al. 2011). AVR genes of L. maculans are located within or adjacent to A:T-rich isochores and some of these are purported to generate sequence

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mutations by exploiting ‘RIP leakage’ (see Section III.D.3). Among the SSPs, 60% located in A:T-rich isochores and 73% in GC-equilibrated isochores were found to contain RxLR amino acid motifs, which facilitate translocation into the host cell (Kale et al. 2010). Gene functions of genes within A:T-rich isochores were also compared by Rouxel et al. (2011) against those of genes within G:Cequilibrated isochores. A:T-rich isochore genes were comparatively deficient in genes involved in cell communication, transmembrane and vesicle-mediated transport, assembly of cellular components, gene regulation, translation, carbon metabolism, cell growth, sporulation and sexual reproduction. Significant enrichment was observed within A:T-rich isochore genes for functions relating to catabolic processes, response to chemical and biotic stimuli, cell wall metabolic processes, cell cycle processes and microtubulebased processes (Rouxel et al. 2011). L. maculans and M. graminicola both undergo a latent biotrophic phase prior to their necrotrophic phase (Howlett et al. 2001; Palmer and Skinner 2002), in contrast to P. nodorum which lacks a biotrophic phase (Oliver 2009). This significant difference in pathogenic lifestyle may be reflected in their respective genome contents. Although this difference has not been addressed in the genome analysis of L. maculans, it has been examined in detail for M. graminicola. The genome of M. graminicola is significantly depleted in CWDE genes targeting cellulose and xylan, carbohydrate binding genes and carbohydrate metabolism genes relative to other phytopathogenic fungi and there are significantly more genes involved in protein metabolism (Goodwin et al. 2011). This lack of CWDE and carbohydrate metabolism is unusual in a cereal pathogen and Goodwin et al. (2011) suggest that this assists M. graminicola to evade detection by host defences during its biotrophic phase. Furthermore, Goodwin et al. (2011) proposed that M. graminicola does not access nutrients from the plant cytoplasm during its biotrophic phase, instead metabolising proteins within the apoplastic fluid and intracellular space. However, after switching to its necrotrophic phase, M. graminicola may express pathogenicity effectors and CWDEs in a similar manner to P. nodorum. This ‘stealth biotrophy’, as coined by Goodwin et al. (2011), suggests that

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Fig. 9.7. The laterally transferred region containing ToxA loci in P. nodorum and P. tritici-repentis. The sequence of P. nodorum scaffold 55 compared with the P. tritici-repentis scaffold 4 contains regions of high sequence similarity flanking the ToxA and transposase genes. The P. nodorum

sequence is flanked by AT-rich sequence that is multiply repeated in the P. nodorum genome. The percent similarities between the P. nodorum and P. tritici-repentis sequences are shown below (adapted from Friesen et al. 2006)

M. graminicola may have originally evolved from an endophytic species.

ToxA was flanked by highly repetitive regions (Fig. 9.7). PCR screening for ToxA in globally representative samples revealed a lower incidence of ToxA (40%; Stukenbrock and McDonald 2007) in P. nodorum isolates compared to 80% in P. tritici-repentis (Friesen et al. 2006). Furthermore, P. nodorum had been described as a pathogen since at least the year 1889, whereas P. tritici repentis was initially identified in 1902 as a saprophyte of grass and in 1928 of wheat. It was only later described as the causal agent of tan spot in 1941. The recent emergence of P. tritici repentis as a plant pathogen and rise to prominence as the most important pathogen of crops in Australia, the low level of diversity in isolates of P. tritici repentis relative to P. nodorum, the presence of a proximal transposase (Fig. 9.7) and flanking repetitive DNA (Fig. 9.7), high sequence conservation at the nucleotide level (Fig. 9.7) and the general lack of sequence conservation between species for other proteinaceous pathogenicity effectors, all suggested that a LGT has occurred between these two species (Friesen et al. 2006). This was the first published evidence of a LGT that was linked to evolution of virulence in a eukaryote. Besides ToxA, several other pathogenicity effector genes of Dothideomycetes have been predicted to have been acquired via LGT events (Oliver and

1. Lateral Gene Transfer Lateral gene transfer (LGT, syn. horizontal gene transfer, HGT) involves the incorporation of the genetic material of one species into the genome of another. There are a number of reported cases of LGTs in the Dothideomycetes involving pathogenicity-related genes (see Chapters 10, 13 in this volume). The best documented among these is the LGT of the necrotrophic effector gene ToxA between P. nodorum and P. tritici-repentis (Friesen et al. 2006). ToxA encodes a 13.2-kDa protein which induces necrosis in Triticum aestivum (wheat) cultivars possessing the dominant allele of the susceptibility gene Tsn1 (Faris et al. 1996). After the sequencing of the P. nodorum genome, it was observed that the sequence of the gene SNOG_16571.1 was 99.7% similar to the previously characterised ToxA gene in P. tritici-repentis. Friesen et al. (2006) used sequences flanking the P. nodorum ToxA sequence to design primer pairs for the purpose of sequencing the corresponding region in P. tritici-repentis. An 11-kb region containing the ToxA locus was conserved between the two species and contained an hAT family transposase gene adjacent to ToxA, which together with


Solomon 2010). The Ace1 polyketide synthase/ non-ribosomal peptide synthase hybrid gene of M. grisea has homologues in several Pezizomycotina species, including P. nodorum (Khaldi et al. 2008). Phylogenetic evidence suggests that LGT is the probable explanation for the acquisition of Ace1 in P. nodorum. The T-toxin biosynthesis gene cluster of C. heterostrophus is another LGT candidate, residing within 1.2 megabases of sequence which is present in race T but absent in race O (Turgeon and Baker 2007). The 6-methyl-salicylic acid biosynthesis gene clusters of the Dothideomycetes, Eurotiomycetes, Lecanoromycetes and Sordariomycetes are also purported to have been acquired via LGT from the Actinobacteria (Kroken et al. 2003; Schmitt and Lumbsch 2009). A recent study predicting probable LGTs of prokaryotic genes into fungal genomes found that LGTs were much more prevalent in the Pezizomycotina (filamentous Ascomycetes, Fig. 9.1), relative to non-filamentous Ascomycetes and other fungal phyla (Marcet-Houben and Gabaldon 2010). They speculated that the differences in genome size and gene density between filamentous Pezizomycotina species (larger genomes, lower gene densities) and the non-filamentous Saccharyomycotina (smaller genomes, higher gene densities) may account for their different rates of LGT. However, due to the diversity of lifestyles and genome compositions between Pezizomycotina species, Marcet-Houben and Gabaldon (2010) were unable to determine any further shared characteristics which may predispose Pezizomycotina species towards LGT. P. nodorum and M. fijiensis represented the Dothideomycetes used in this study, both having 23 predicted prokaryotic LGT events. One notable example of prokaryotic LGT to these Dothideomycetes mentioned by the authors was the transfer of a bacterial catalase from the bacterium Psuedomonas syringae. This catalase was predicted to decompose reactive oxygen species and potentially plays a role in the fungal evasion of host plant defences. This gene was also detected in the Leotiomycete Botrytis cinerea, and phylogenetic considerations suggest that B.cinerea obtained its copy of the gene from the Dothideomycetes. 2. Generation of Diversity via RIP Avirulence (AVR) effectors are gene products which play a role in the pathogenesis of some Dothideomycete species, most notably L. macu-

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lans (Fudal et al. 2009; Rouxel et al. 2011; Van de Wouw et al. 2010) and P. fulvum (Wulff et al. 2009). AVR effectors are disadvantageous to a pathogen if the host plant possesses resistance (R) genes which are capable of recognising them thereby activating the host plant’s defences. Therefore fungi which employ AVR effectors are under selective pressure to mutate and adapt. The AVR genes AvrLm6 and LymCys1 of L. maculans have been observed to exhibit RIPlike polymorphism between isolates (Fudal et al. 2009; Van de Wouw et al. 2010). This is unusual as only one copy of both of these genes is found in the L. maculans genome and RIP is triggered by the alignment of repetitive DNA (2 or more copies). However RIP acting upon repetitive DNA has been reported to ‘leak’ into neighbouring nonrepetitive regions within a limited range (Irelan et al. 1994). Fudal et al. (2009) proposed that certain AVR genes of L. maculans have accumulated RIP mutations from their neighbouring repetitive DNA regions. Thus RIP could be being exploited by the fungus as a means of accelerating the rate of mutation in certain genes (Irelan et al. 1994; Fudal et al. 2009). This was supported by Van de Wouw et al. (2010), who observed a gradient in the frequency of RIP-like mutations between AVR genes of different isolates which is dependent upon the distance from the neighbouring repeat. Other pathogenicity effectors of the Dothideomycetes are also frequently associated with neighbouring repetitive DNA or A:T-rich regions. These include the effector genes PtrToxA of P. tritici-repentis and SnToxA and SnTox3 of P. nodorum which are next to transposable elements (Friesen et al. 2006; Liu et al. 2009). The dispensable chromosomes of M. graminicola (see Section II.B) contain clusters of genes encoding secreted proteins, which are also frequently transposon-associated (Goodwin et al. 2011). Whether association with repetitive DNA also generates effector diversity in these species remains to be determined.

VI. Conclusions The Dothideomycetes are an ecologically diverse class of Fungi of which many species are pathogens of economically important crops. Significant phytopathogenic species share the ability to rapidly

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adapt to host plant defences and agricultural practices by mutation or lateral gene transfer. Highly virulent isolates of several species have rapidly emerged and spread over vast distances. While the attributing factor for this remarkable adaptability is currently unknown, available genome data has revealed several complex mechanisms geared towards generating diversity at the genomic level. Dothideomycete genome structure is remarkably plastic, undergoing major rearrangement whilst retaining gene content within the same chromosomes. They are believed to readily acquire new genes via lateral gene transfer, many of which are involved in pathogenicity. Fungal genome defences against transposons may have also been hijacked in order to accelerate mutation rates in pathogenicity effector genes. Significant proportions of Dothideomycete genes are unique to the class or respective genera, indicating adaptations to highly specialised ecological niches. It is therefore unsurprising that each of the Dothideomycetes outlined here employ diverse pathogenicity strategies specifically tailored to their host and lifestyle. Acknowledgements This work was supported by the Grains Research and Development Corporation, Barton, ACT, Australia.

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10

Evolution of Genes for Secondary Metabolism in Fungi

INES TEICHERT1, MINOU NOWROUSIAN1

CONTENTS I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Polyketides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Polyketide Synthases in Fungi. . . . . . . . . . . . . . . . B. Clustering and Genomic Distribution of Polyketide Biosynthesis Genes in Ascomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Biological Functions of Fungal Polyketides. . III. Nonribosomal Peptides . . . . . . . . . . . . . . . . . . . . . . . . . . A. Nonribosomal Peptide Synthetases in Fungi B. Biological Functions of Fungal Nonribosomal Peptides . . . . . . . . . . . . . . . . . . . . . . . IV. Alkaloids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Terpenes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI. Melanins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. DOPA Melanin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. DHN Melanin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Biological Functions of Fungal Melanins . . . . VII. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

231 232 232 234 235 237 237 238 239 239 240 242 242 243 244 245

I. Introduction Fungi are a group of organisms well noted for the production of so-called secondary metabolites. These are mostly small molecules that are not produced constantly, but are often associated with distinct morphological phases of the life cycle and/or produced only under specific environmental conditions (Keller et al. 2005). Thus, in contrast to the products of primary metabolism, they are usually not essential for the organism, at least not under laboratory conditions. However, several metabolites produced by enzymes that are characteristic for secondary metabolism are involved in core cellular processes that are usually regarded as part of primary metabolism. Therefore, the term “natural products” is sometimes used to describe these molecules. Secondary metabolites comprise a wide variety of chemical structures, 1

Lehrstuhl fu¨r Allgemeine und Molekulare Botanik, RuhrUniversita¨t Bochum, 44780, Bochum, Germany; e-mail: minou. [email protected]

and they are commonly classified according to which precursor molecules from primary metabolism are used for their biosynthesis, and which specific enzyme class synthesizes their basic chemical structure (Hoffmeister and Keller 2007; see chapter 13 in this volume). The best studied to date are polyketides, nonribosomal peptides, terpenes and alkaloids. A common feature of secondary metabolites is that most are bioactive, with many of them being studied for their pharmacological or toxic effects on humans, livestock, and crops (Bennett and Klich 2003; Misiek and Hoffmeister 2007). There exists a huge number of chemically different secondary metabolites, with each metabolite often being taxonomically restricted to few or one fungal species. However, despite numerous studies on the biosynthesis of secondary metabolites and their impact on human society, their role in the biology of the fungi producing them remains largely unknown (Fox and Howlett 2008). During the past 30 years, genetic and molecular studies have identified many genes that encode enzymes or regulators involved in the biosynthesis of fungal secondary metabolites. One characteristic feature that emerged early on is that genes encoding enzymes for a specific biosynthetic pathway are often physically clustered within the genome (Keller and Hohn 1997). The sequencing in recent years of more than 100 fungal genomes underscored this common theme, and at the same time uncovered many more genes/ clusters involved in secondary metabolism in each sequenced species than the number of known metabolites produced by this species. Many of these clusters are transcriptionally silent under laboratory conditions, which explains the lack of identified metabolites from these clusters (Brakhage and Schroeckh 2010); however, genome sequence data suggest that the biochemical potential of most fungi far exceeds their currently known metabolic spectrum. Another finding that became Evolution of Fungi and Fungal-Like Organisms, The Mycota XIV S. Po¨ggeler and J. Wo¨stemeyer (Eds.) © Springer‐Verlag Berlin Heidelberg 2011

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Ines Teichert and Minou Nowrousian

apparent through an increasing amount of sequence data is that genes for secondary metabolism are not infrequently subject to horizontal gene transfer among fungi or between fungi and other taxonomic groups (Rosewich and Kistler 2000; Khaldi et al. 2008). In this review, we will therefore focus on: (i) the distribution and phylogenetic origins of secondary metabolites and their biosynthetic and regulatory genes in fungi and (ii) the known biological functions of these molecules. In the following sections, we will cover the four main classes of fungal secondary metabolites, namely polyketides, nonribosomal peptides, alkaloids, and terpenes. In addition, we will describe in more detail the melanins, a group of secondary metabolites that is not characterized by a common biochemistry but rather by common properties. Melanins have been studied intensively, and therefore much is known about their biosynthesis, genetic regulation, and in some cases biological function.

II. Polyketides Polyketides are the largest group of fungal secondary metabolites. They comprise structurally very different metabolites ranging from small molecules like the mycotoxin aflatoxin to large, complex structures like melanins. However, a common denominator is that their basic structures are synthethized from small carboxylic acids (e.g., acetate or propionate) by enzymes called polyketide synthases (PKSs; Staunton and Weissman 2001; Keller et al. 2005; Cox 2007). PKSs can be divided into types I–III. While type II PKSs are multimeric proteins that are restricted to bacteria and therefore will not be discussed here, type I and type III PKSs can be found in fungi among other organisms. These PKSs and their products will be described in the following sections.

A. Polyketide Synthases in Fungi The majority of fungal PKSs are type I PKSs. These are large multidomain proteins of several thousand amino acids with a domain structure that is closely related to that of type I fatty acid synthases (FASs) found in animals and fungi (Cox 2007). Type I PKSs have been found in bacteria, fungi, and protozoa (Staunton and Weissman

2001; Kroken et al. 2003; Eichinger et al. 2005). Interestingly, fungal type I PKS are more closely related to animal FASs than to fungal FASs (Kroken et al. 2003). Among fungi, type I PKSs are widely distributed in ascomycetes, the exception being the groups of saccharomycetes and schizosaccharomycetes with mostly unicellular members (Kroken et al. 2003). They can also be found in basidiomycetes, but are absent from the sequenced genomes of zygomycetes and chytridiomycetes (Table 10.1). At present, it is difficult to determine whether their absence in these more basal groups of eumycota is due to secondary loss or whether this indicates that fungal PKSs were an “invention” in the ancestor of the dikarya (asco-, basidiomycetes). However, it is noteworthy that the social amoeba Dictyostelium discoideum contains 43 predicted pks genes in its genome (Eichinger et al. 2005), and BLAST searches in sequenced genomes of other protozoa retrieve several putative pks genes in other species not closely related to D. dictyostelium. At present, the somewhat patchy picture that the distribution of PKSs among eukaryotes presents is not easily explained. One reason for this might be that pks genes are fast-evolving genes with gene duplication events and subsequenct differential loss as well as horizontal gene transfer contributing to their phylogenetic distribution (see below). In fungi, the highest number of pks genes per genome can be found in filamentous ascomycetes (Table 10.1). When the first genomes of filamentous fungi were sequenced, it was hypothesized that plant pathogenic fungi might have undergone a life style-specific expansion of pks genes, because the genomes of plant pathogens like Gibberella zeae or Magnaporthe grisea contain significantly more pks genes than the genome of the first sequenced filamentous fungus, Neurospora crassa (Galagan et al. 2003, Dean et al. 2005, Cuomo et al. 2007). However, with more fungal genomes available now, this hypothesis no longer seems to be the most likely, because saprobic ascomycetes harbor similar numbers of pks genes in their genomes compared to plant pathogenic fungi (Table 10.1). Apart from N. crassa, which generally contains few gene families due to an active genome defense mechanism that prevents gene duplications (Borkovich et al. 2004), the lowest number of pks genes among filamentous ascomycetes is present in the genome of the black truffle Tuber melanosporum (Martin et al.


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Table 10.1. Polyketide synthase (pks) genes (including pks-nrps hybrid genes) and nonribosomal peptide synthetase (nrps) genes in sequenced fungal genomes. References are given as footnote. The single nrps gene present in Zygomycetes and S. cerevisiae represents the aminoadipate reductase that is involved in fungal lysin biosynthesis and is present in all fungal genomes investigated (Bushley and Turgeon 2010) Species Chytridiomycetes Allomyces macrogynus Batrachochytrium dendrobatidis Zygomycetes Mucor circinelloides Rhizopus oryzae Ascomycetes Saccharomycetes Saccharomyces cerevisiae Schizosaccharomycetes Schizosaccharomyces pombe Pezizomycetes Tuber melanosporum Leotiomycetes Botrytis cinerea Sclerotinia sclerotiorum Sordariomycetes Gibberella zeae Magnaporthe grisea Nectria haematococca Neurospora crassa Podospora anserina Sordaria macrospora Dothideomycetes Pyrenophora tritici-repentis Stagonospora nodorum Eurotiomycetes Aspergillus fumigatus Aspergillus nidulans Aspergillus niger Aspergillus oryzae Coccidioides immitis Penicillium chrysogenum Basidiomycetes Coprinus cinereus Cryptococcus neoformans Laccaria bicolor Malassezia globosa Postia placenta Ustilago maydis

Life style

Number of pks genes

Number of nrps genes

References

Saprobe Amphibian pathogen

– –

2 4

1 2, 3

Saprobe Saprobe, opportunistic human pathogen

– –

1 1

4 5

Saprobe

–

1

6, 31

Saprobe

–

2

6, 32

Mycorrhizal symbiont

2

2

7

Plant pathogen Plant pathogen

17 16

5 5

8 9

Plant pathogen Plant pathogen Plant pathogen Saprobe Saprobe Saprobe

15 31 13 8 20 11

20 6 12 3 5 3

10, 11 12 13 6, 14 15, 33 16

Plant pathogen Plant pathogen

16 20

6 8

17 18

Saprobe, opportunistic human pathogen Saprobe Saprobe Saprobe Human pathogen Saprobe

14

13

19

28 41 30 10 22

13 17 18 5 11

20 21 19, 20 22 23

Saprobe Human pathogen Mycorrhizal symbiont Human pathogen Saprobe Plant pathogen

2 – 1 1 1 3 (partial)

4 1 1 1 2 3

24 25 26, 34 27, 35 28, 35 29,30

References: 1 http://www.broadinstitute.org/annotation/genome/multicellularity_project/MultiHome.html 2 http://genome.jgi-psf.org/Batde5/Batde5.home.html 3 http://www.broadinstitute.org/annotation/genome/batrachochytrium_dendrobatidis/MultiHome.html 4 http://genome.jgi-psf.org/Mucci2/Mucci2.home.html 5 http://www.broadinstitute.org/annotation/genome/rhizopus_oryzae/MultiHome.html 6 Kroken et al. (2003) 7 Martin et al. (2010) 8 http://www.broadinstitute.org/annotation/genome/botrytis_cinerea/Home.html 9 http://www.broadinstitute.org/annotation/genome/sclerotinia_sclerotiorum/MultiHome.html 10 Gaffoor et al. (2005)

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11 Cuomo et al. (2007) 12 Dean et al. (2005) 13 Coleman et al. (2009) 14 Galagan et al. (2003) 15 Espagne et al. (2008) 16 Nowrousian et al. (2010) 17 http://www.broadinstitute.org/annotation/genome/pyrenophora_tritici_repentis/Home.html 18 Hane et al. (2007) 19 Nierman et al. (2005) 20 Machida et al. (2005) 21 Pel et al. (2007) 22 http://www.broadinstitute.org/annotation/genome/coccidioides_group/MultiHome.html 23 van den Berg et al. (2008) 24 http://www.broadinstitute.org/annotation/genome/coprinus_cinereus/MultiHome.html 25 http://www.broadinstitute.org/annotation/genome/cryptococcus_neoformans/MultiHome.html 26 Martin et al. (2008) 27 Xu et al. (2007) 28 Martinez et al. (2009) 29 Ka¨mper et al. (2006) 30 Bo¨lker et al. (2008) 31 http://www.yeastgenome.org 32 Schrettl et al. (2004) 33 http://podospora.igmors.u-psud.fr/ 34 http://genome.jgi-psf.org/Lacbi1/Lacbi1.home.html 35 http://www.ncbi.nlm.nih.gov/sutils/genom_table.cgi

2010; Table 10.1). T. melanosporum is currently the only member of the basal ascomycete group of pezizomycetes with a sequenced genome, which is also the only sequenced genome from a mycorrhizal ascomycete. Therefore, it is difficult to determine whether its low number of only two pks genes represents an ancestral state in filamentous ascomycetes or is an adaptation to the mycorrhizal life style. Generally, a connection between the number of pks genes and different life styles in filamentous ascomycetes is not at present apparent, but this might change when more fungal genomes become available in the future. In ascomycetes, fusion genes encoding a combined PKS (type I) and nonribosomal peptide synthase (NRPS, see below) have been discovered in a number of species. The corresponding PKSNRPS enzymes synthesize polyketides fused to amino acids, thereby further increasing the biochemical variety of secondary metabolites these fungi can produce (Cox 2007). Examples for pksnrps genes are the pathogenicity factor ace1 in Magnaporthe grisea and the fusA gene from Fusarium moniliforme and F. venenatum that is involved in the biosynthesis of the secondary metabolite fusarin (Bo¨hnert et al. 2004; Song et al. 2004). Type III PKSs, also known as chalcone/ stilbene synthases, were originally identified in plants, but have since then been found in bacteria,

fungi, and protozoa (Seshime et al. 2005; Austin et al. 2006; Funa et al. 2007; Flores-Sanchez and Verpoorte 2009). The plant and fungal enzymes are much smaller proteins than type I PKSs with just one multifunctional protein domain, while in D. discoideum, a type III PKS domain is fused to a type I fatty acid synthase to create a large novel enzyme responsible for the biosynthesis of the differentiation-inducing factor DIF1 (Austin et al. 2006). The number of type III pks genes in fungal genomes is lower than those encoding type I PKSs with usually only one putative type III pks gene per genome in filamentous ascomycetes, and none in ascomycetous yeasts, basidiomycetes, zygomycetes, and chytridiomycetes.

B. Clustering and Genomic Distribution of Polyketide Biosynthesis Genes in Ascomycetes The basic structures of polyketides that are synthesized by PKSs are usually further modified by additional enzymes that act downstream of the PKS in a biosynthesis pathway and contribute to the high structural diversity of these molecules. In filamentous fungi, the pks genes and other genes of such pathways are often physically clustered as well as transcriptionally co-regulated (Hoffmeister


and Keller 2007). This transcriptional regulation is sometimes mediated by a transcription factor that is part of the cluster itself. This is, for example, the case for AflR, the transcriptional regulator of the clustered genes for aflatoxin and sterigmatocystin biosynthesis in several Aspergilli (Woloshuk et al. 1994; Fernandes et al. 1998). An additional level of transcriptional regulation can be provided by the chromation structure of the genomic locus containing the clustered genes. This was first discovered through studies of the laeA gene in several Aspergilli (Bok and Keller 2004; Bok et al. 2006a, b). An laeA mutant strain no longer expresses many gene clusters for secondary metabolites of Aspergillus nidulans and other Aspergilli, whereas clusters of genes for primary metabolites are unaffected. LaeA has homologies to methyltransferases, and it has been suggested that it is involved in methylation changes in histones or other chromatin-associated proteins, thereby regulating the chromatin conformation and thus the transcriptional accessibility of gene clusters for secondary metabolism. In Aspergilli, LaeA acts in a complex with the Velvet and Velvet-like proteins, regulatory proteins that mediate the balance between development and secondary metabolism in response to extracellular stimuli like light (Bayram et al. 2008).

Since the discovery of laeA, several more factors involved in the regulation of chromatin structure have been shown to be necessary for the correct expression of polyketide and other secondary metabolite gene clusters (Shwab et al. 2007; Bok et al. 2009; Reyes-Dominguez et al. 2010). Among these is the histone deacetylase HdaA that is necessary for the transcriptional repression of telomereproximal but not telomere-distal gene clusters in A. nidulans (Shwab et al. 2007). In connection with this, it is interesting to note that secondary metabolite gene clusters in filamentous ascomycetes are often concentrated towards telomere ends (Galagan et al. 2005; Nierman et al. 2005; Perrin et al. 2007). This nonrandom genomic distribution may be one reason for the high diversity and species-specific distribution of clusters for polyketide biosynthesis genes and other genes for secondary metabolites, because genomic regions close to chromosome ends are more prone to recombination than telomere-distal regions. This might lead to recombination within genes and clusters and resulting in the

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observed lack of orthology of many secondary metabolite clusters within fungal genomes. Another reason for the high diversity might be that telomereproximal clusters are often surrounded by repeat regions and/or transposons that might facilitate recombination events including duplications and deletions. In addition, transposon sequences adjacent to clusters were recently shown to be involved in the regulation of cluster gene expression (Shaaban et al. 2010). The chromatin structure-dependent regulation of secondary metabolite genes as mentioned above may be one reason for the physical clustering of these genes. However, another hypothesis that has been proposed is based on the fact that clustered genes can be used to move complete biochemical pathways between species by means of horizontal gene transfer (HGT) (Walton 2000) (see chapter 9 this volume). This hypothesis states that clustering of genes is not generally necessary for the coregulation of genes for a pathway, and that therefore, for clustering to be maintained, other selective pressures must act under which clustering confers an advantage compared to dispersed genomic locations. One such selective pressure would be in place if the clustered genes were propagated not only through vertical transmission (i.e. from one generation to the next) but through HGT to the same or different species. Since this hypothesis was put forward, there has been increasing evidence, e.g., through comparative analysis of sequenced fungal genomes, that HGT in general and specifically HGT of secondary metabolite clusters is indeed more common than previously thought and might be an evolutionary mechanism for fungi to increase their biochemical potential (Garcia-Vallve´ et al. 2000, Rosewich and Kistler 2000, Friesen et al. 2006, Patron et al. 2007, Khaldi et al. 2008, Khaldi and Wolfe 2008, Coleman et al. 2009, Nowrousian et al. 2010) (see chapter 9 and 13 this volume). Thus, both their propagation by HGT as well as their regulation at the chromatin level might be reasons for the evolutionary conservation of the clustered organization of genes for the biosynthesis of polyketides and other secondary metabolites. C. Biological Functions of Fungal Polyketides Although through genome sequencing projects much has been learned about the number and

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genomic organization of polyketide biosynthesis genes in fungi, there is still little knowledge about what products the majority of the corresponding enzymes might synthesize and what the biological functions of these products might be (Fox and Howlett 2008). A role in fungal biology has been established for only a few polyketides; however, these examples already indicate that polyketides influence a broad variety of manifestations of fungal life. Known functions include protection against stress, plant pathogenicity, or effects on sexual development, and some of these examples will be discussed in this section. One of the best-studied classes of polyketides are the melanins, complex macromolecules that can be synthesized via a polyketide pathway and which serve many functions ranging from pathogenicity factors to protection from environmental stresses. As they can also be derived from other biosynthesis pathways and because of their important role in fungal biology, they will be described in a special section (see Section VI). Polyketides may act as pathogenicity or virulence factors of plant-pathogenic fungi (van der Does and Rep 2007). One example is T-toxin from Cochliobolus heterostrophus, the causal agent of southern corn leaf blight (Kono et al. 1980). T-toxin is produced only by race T of C. heterostrophus. This race is highly virulent against maize containing Texas male sterile (T) cytoplasm, whereas race O does not produce T-toxin and is weakly virulent on maize in general. Detailed studies of the genetic mechanisms behind this race-specific toxin production showed that the ability to synthesize T-toxin is conferred by two genomic loci, Tox1A and Tox1B, that are present in race T, but not in race O. These loci reside at the breakpoints of two chromosomes that have undergone reciprocal translocations in race T compared to race O. During the past 15 years, nine genes from these loci were shown to be involved in the production of T-toxin; among these are two polyketide synthase genes, one decarboxylase, and five dehydrogenases (Yang et al. 1996; Kodama et al. 1999, Rose et al. 2002; Baker et al. 2006; Inderbitzin et al. 2010). Another example where a polyketide synthase is necessary for host-specific interactions is studied in Magnaporthe grisea. In this rice pathogen, strains carrying the avirulence gene ACE1 are unable to infect rice cultivars carrying the resistance gene Pi33. ACE1 is a hybrid pks-nrps gene, and

even though the corresponding metabolite has not been identified yet, it is likely that a polyketide-based metabolite is involved in this host-specific recognition event (Bo¨hnert et al. 2004, Collemare et al. 2008). However, polyketides can also act as nonhost-specific pathogenicity factors or toxins, for example, melanins (see Section VI) or cercosporin, a toxin produced by many Cercospora species that are pathogenic to a wide range of crops (Choquer et al. 2005).

Polyketides are produced not only by plantpathogenic fungi, but also by a large number of filamentous ascomycetes, many of which live in the soil. It has been suggested that polyketides and other secondary metabolites produced by soilliving fungi protect the fungus from predators like nematodes or arthropods or prevent growth of competing fungi and other microorganisms (Vining 1990; Fox and Howlett 2008). There is little direct evidence for this hypothesis; however, the increasing number of fungal mutants with defects in polyketide biosynthesis now allows comparative studies where the fitness of mutant strains is compared to that of wild-type strains. Such a study recently showed that LaeA mutants of A. nidulans are preferentially consumed by the fungivorous springtail Folsomia candida (Rohlfs et al. 2007). These mutants have an overall reduced production of secondary metabolites compared to the wild type (see Section II.B), therefore it is not yet clear whether one specific metabolite or a combination of different factors might be responsible for the effect on springtail feeding. Another possibly protective function has recently been hypothesized for fungal polyketides and other secondary metabolites, namely a role in the reaction of fungi to oxidative stress (Reverberi et al. 2010). Polyketides have also been implied as factors involved in fungal development. However, while a close correlation between asexual sporulation and the production of polyketides and other secondary metabolites has been observed in many cases, usually there is no direct requirement of the metabolite for the sporulation process, at least under laboratory conditions (Calvo et al. 2002). With regard to sexual development, several studies suggest a role for polyketides in the production of fruiting bodies and sexual spores in filamentous ascomycetes. Early studies on zearalenone, a polyketide from Gibberella zeae, indicated that zearalenone influences fruiting body production in a concentration-dependent manner (Wolf and Mirocha 1973). The fact that the effect of


zearalenone is concentration-dependent with low concentrations enhancing and high concentrations inhibiting fruiting body morphogenesis has led to the hypothesis that the metabolite might act as a signaling molecule. Genetic analyses of predicted polyketide biosynthesis genes in Neurospora crassa and Sordaria macrospora showed that in N. crassa, one pks gene is essential for fruiting body development, whereas in both fungi, a putative dehydrogenase from a novel polyketide biosynthesis gene cluster is involved in fruiting body maturation (Nowrousian 2009). However, the metabolites produced by the corresponding enzymes have not been identified yet. Generally, several biological roles that have been identified or suggested for fungal polyketides, namely host–pathogen interactions, protection in soil environments, and signaling during sexual development, would also offer an explanation for the high diversity of these metabolites: Virulence factors as well as protecting molecules may be part of an arms race between pathogen and host or predator and prey (van der Does and Rep 2007); whereas signaling molecules involved in sexual reproduction are often species-specific and may constitute a mechanism to ensure reproductive isolation by preventing mating of individuals from different species. A trend for genes involved in stress-related functions to be subject to more gene deletion and loss events than genes involved in more basic cellular functions was observed across fungal genomes in general (Wapinski et al. 2007).

III. Nonribosomal Peptides The second major group of fungal secondary metabolites are nonribosomal peptides. Similar to polyketides, they are synthesized by large multidomain enzymes called nonribosomal peptide synthetases (NRPS; Keller et al. 2005). In contrast to genetically encoded peptides that are synthesized at ribosomes, nonribosomal peptides can contain not only proteinogenic, but also nonproteinogenic amino acids and this, in addition to other chemical modifications either by NRPSs or downstream enzymes, is one reason for their great structural diversity. Nonribosomal peptides comprise a number of widely used antibiotics and immunosuppressive drugs (Hoffmeister and Keller 2007); however, a number of studies have

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also highlighted their diverse roles in fungal biology ranging from functions in cellular metabolism and development to pathogenicity. These functions as well as insights in the evolution of the nonribosomal peptide biosynthesis genes will be described in the next sections.

A. Nonribosomal Peptide Synthetases in Fungi Genes encoding NRPSs can be found in fungi and bacteria, but not in animals and plants (Bushley and Turgeon 2010). They comprise one or more modules each of which usually consists of three domains for adenylation (A domain), thiolation (T domain or peptidyl carrier protein) and condensation (C domain; Hoffmeister and Keller 2007). In recent years, genome sequencing projects have made genome-wide surveys of nrps genes across many fungal groups possible, and a recent phylogenomic study revealed that fungal NRPSs fall into two major groups that can further be subdivided into distinct subfamilies (Bushley and Turgeon 2010). The first group comprises mono/bi-modular NRPSs that have more ancient origins and a more conserved domain architecture than the mostly multimodular NRPSs of the second group. Bacterial NRPSs cluster with fungal NRPSs of the first group, whereas the second group contains only fungal NRPSs. Among fungi, most filamentous ascomycetes contain many nrps genes, basidiomycetes and chytrids contain few, whereas zygomycetes and the yeast S. cerevisiae contain only one gene that belongs to the nrps group, namely an aminoadipate reductase that is involved in fungal lysin biosynthesis (Bushley and Turgeon 2010; Table 10.1). Thus, nrps genes seem to be an ancient invention with a history of more recent expansion and loss in filamentous ascomycetes. Additionally, similar to the case for some pks genes, there is evidence for horizontal gene transfer of several nrps genes, in this case from bacteria to fungi. A well-studied example are the genes encoding the ACV synthetases involved in the biosynthesis of beta-lactam antibiotics in Penicillium chrysogenum, A. nidulans, and Acremonium chrysogenum (Landan et al. 1990; Penalva et al. 1990; Buades and Moya 1996; Brakhage et al. 2009; Bushley and Turgeon 2010). Another similarity of pks and nrps genes is that both are often found in clusters with other

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genes that are involved in the same biochemical pathway (Keller and Hohn 1997). Studies of the genomes of several members of the Eurotiomycetes have shown that, similar to the case of other secondary metabolite clusters, nrps clusters are often located close to telomers on chromosomes and are regulated by the major regulator LaeA (Galagan et al. 2005; Nierman et al. 2005; Perrin et al. 2007; Fedorova et al. 2008). The subtelomeric location may contribute to the high diversity of nrps clusters due to increased recombination rates at these chromosomal locations (see Section II.B).

iron homeostasis (Eisendle et al. 2003), they play specific roles in the biology of several fungi.

B. Biological Functions of Fungal Nonribosomal Peptides

In N. crassa, siderophores were shown to be essential for the germination of conidia (Horowitz et al. 1976, Matzanke et al. 1987). In A. nidulans, deletion of sidC, the gene encoding the NRPS responsible for the biosynthesis of the siderophore ferricrocin, results in a pleiotropic phenotype with defects in oxidative stress resistance, conidial germination and fruiting body development (Eisendle et al. 2006). Requirement of siderophores for sexual development was also observed in C. heterostrophus and Fusarium graminearum (Oide et al. 2007). In the human pathogen A. fumigatus as well as in several plantpathogenic fungi, siderophores were shown to be required for pathogenicity; and it was hypothesized that they play essential roles in supplying fungal pathogens with iron, which is a limiting factor within host environments (Lee et al. 2005; Oide et al. 2006; Schrettl et al. 2007).

Several nonribosomal peptides have received much attention due to their pharmaceutical value as antibiotic or immunosuppressive drugs (Hoffmeister and Keller 2007); however, a number of studies have also shed light on the biological functions of this diverse class of molecules. These range from functions in cellular metabolism to roles in development, pathogenicity, and stress response. Recent phylogenomics studies have revealed that the more conserved NRPSs that comprise mostly mono/bi-modular enzymes are often involved in more basic cellular processes whereas the multimodular NRPSs from the second group of fungal NRPSs play a role in processes that are often species-specific or even strain-specific and are more consistent with the classical definition of secondary metabolites (Bushley and Turgeon 2010). Among the NRPSs that synthesize products for core cellular processes are the aminoadipate reductases that are involved in fungal (and bacterial) lysin biosynthesis and are present in all fungal genomes analyzed so far (Velasco et al. 2002; Bushley and Turgeon 2010). A second group of NRPSs that play a role in basic metabolism are enzymes involved in siderophore biosynthesis. Siderophores are low molecular mass iron chelators that can either be secreted for acquisition of extracellular iron or can be used for intracellular iron storage (Haas et al. 2008). They are present in the genomes of filamentous ascomycetes, a number of basidiomycetes, and in the yeast Schizosaccharomyces pombe (Schrettl et al. 2004; Bushley et al. 2008). Apart from their general function in

A number of other nonribosomal peptides have also been identified as pathogenicity factors in several plant pathogenic fungi. In contrast to siderophores, these nonribosomal peptides are mostly produced by multimodular NRPSs with restricted phylogenetic distribution and often act as host-specific toxins. Examples are HC-toxin from Cochliobolus carbonum (Panaccione et al. 1992), AM-toxin from Alternaria alternata (Johnson et al. 2000), enniatin from Fusarium avenaceum (Herrmann et al. 1996), and sirodesmin from Leptosphaeria maculans (Gardiner et al. 2004; Elliott et al. 2007). NRPSs also participate in the biosynthesis of ergot alkaloids, which are discussed in the next section (see Section IV). Nonribosomal peptides may also act as protective agents for the producing fungi. It has long been hypothesized that, for example, beta-lactam antibiotics are produced by soil-living fungi to reduce bacterial competition. However, definite proof for a precise biological role of nonribosomal peptide metabolites is missing in many cases. One exception is the case of peramine, a nonribosomal peptide synthesized by the endophyte Epichloe¨ festucae. Mutants lacking the corresponding NRPS do not produce peramine; and associations of the mutant strain and perennial ryegrass are as susceptible to weevil feeding damage as plants without the endophytic symbiont, indicating that peramine protects the plant–fungus association from herbivore damage (Tanaka et al. 2005). Peptides produced by multimodular NRPSs may also play a role in fungal development, as was shown in a gene disruption mutant of the Alternaria


brassicicola AbNPS2 gene which displays severe defects in conidial cell wall morphology (Kim et al. 2007).

IV. Alkaloids Alkaloids are secondary metabolites that contain nitrogen in a heterocycle that usually originates from amino acids. In contrast to polyketides or nonribosomal peptides, their biosynthesis is not dependent on a specific enzyme class like PKSs or NRPSs (although e.g., NRPSs can be involved in alkaloid biosynthesis, see below). Alkaloids are produced by many organisms and show a great chemical diversity. In fungi, the best studied alkaloids are the ergot and loline alkaloids (Hoffmeister and Keller 2007). They are produced by Claviceps, Epichloe¨, and Neotyphodium species, grass pathogens and endophytes of the family Clavicipitaceae. Ergot alkaloids are well known for their pharmacological properties, which have been review extensively (Schardl et al. 2006; Haarmann et al. 2009). Elucidation of the biosynthetic pathway leading to ergotamine and other ergot alkaloids was possible through studies of Claviceps purpurea and other Claviceps species (Haarmann et al. 2009). The first step of the pathway is the isoprenylation of tryptophan to 4-dimethylallyltryptophan by dimethylallyl tryptophan synthase (DMATS; Tsai et al. 1995; Tudzynski et al. 1999). Later steps involve several genes encoding NRPSs that convert the intermediate lysergic acid to the ergopeptines ergotamine in C. purpurea and ergovaline in Neotyphodium species (Panaccione et al. 2001; Haarmann et al. 2005, 2008). The genes encoding these NRPSs as well as the DMATS and other enzymes of the pathway are clustered in the genome of C. purpurea and other Claviceps species (Haarmann et al. 2005; Lorenz et al. 2009). Data mining of fungal genome sequences revealed that DMATS genes are present in many fungi, and the enzymatic functionality of several of the encoded proteins was demonstrated for genes from Aspergillus fumigatus and Malbranchea aurantiaca (Unso¨ld and Li 2005; Ding et al. 2008; Metzger et al. 2009; Steffan et al. 2009). Thus, it is likely that the ability to produce ergot alkaloids or similar structures is widespread in fungi and that further genome mining will reveal more about the evolution of the corresponding genes and enzymes.

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The second group of fungal alkaloids that have been studied in detail are the loline alkaloids. They are produced by endophytic Epichloe¨ and Neotyphodium species and have insecticidal properties that protect the fungal–grass symbiosis from herbivores (Schardl 2001). Similar to the case for other secondary metabolites, genes for loline alkaloids are clustered (Wilkinson et al. 2000; Spiering et al. 2005); however, the exact biochemical pathway of loline biosynthesis has not yet been determined and it is not yet clear whether other fungi outside of the Clavicipitaceae have the ability to synthesize these compounds.

V. Terpenes Terpenes are metabolites that are derived from activated forms of isoprene, a hydrocarbon molecule with five carbon atoms; therefore the number of carbon atoms in terpenes is usually a multiple of five. Terpenes can be further modified, for example, by oxidation, and are then called terpenoids or isoprenoids, although “terpenes” and “terpenoids” are often used interchangeably. Terpenes are widespread metabolites present in many groups of organisms. They include not only classical secondary metabolites, but also molecules like sterols or carotenoids that are important for basic cellular functions in many organisms (Sandmann 1994; Vershinin 1999; Sturley 2000). Carotene derivatives also serve as communication molecules in fungi as reviewed elsewhere (Schimek and Wo¨stermeyer 2009; see Chapter 13 in this volume). However, a number of typical fungal secondary metabolites are terpenoids, e.g gibberellins, indole-diterpenes, and trichothecenes (Hoffmeister and Keller 2007), and these will be described in this section. Gibberellins are phytohormones that regulate various developmental processes in plants. However, gibberellins were first discovered as secondary metabolites of the plant pathogenic ascomycete Gibberella fujikuroi. They cause, among other symptoms, hyperelongation of internodes in infected plants (Bo¨mke and Tudzynski 2009). The elucidation of gibberellin biosynthesis pathways in plants and fungi has uncovered fundamental differences, therefore it is now assumed that these pathways evolved independently (Hedden and Kamiya 1997; Tudzynski 2005; Yamaguchi 2008). In fungi, genes for gibberellin biosynthesis have

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been identified in G. fujikuroi, Sphaceloma manihoticola, and a Phaeosphaeria species; and in those species where it was investigated, the genes are physically clustered (Kawaide et al. 1997; Tudzynski and Ho¨lter 1998; Tudzynski et al. 1998; Bo¨mke et al. 2008). Interestingly, Gibberella belongs to the Sordariomycetes whereas Sphaceloma and Phaeosphaeria belong to the only distantly related Dothideomycetes. However, within the genome sequence of the Sordariomycetes Fusarium graminearum (a close relative of G. fujikuori) and Neurospora crassa, no gibberellin biosynthesis cluster was found (Bo¨mke and Tudzynski 2009). This raises the question about the origin of this somewhat patchy distribution of gibberellin biosynthesis clusters in fungal genomes. One possibility is horizontal gene transfer, but this has not been investigated in detail, and a history of differential duplication and loss for this cluster and genes for the biosynthesis of related terpenoids cannot be excluded (Bo¨mke and Tudzynski 2009). Further analysis of sequenced fungal genomes might shed light on the distribution and evolutionary origin of this class of secondary metabolites. Fungal secondary metabolites belonging to the group of indole-diterpenes have been isolated from a number of species among the Eurotiomycetes and Sordariomycetes (Saika et al. 2008). Well studied indole-diterpenes include, for example, paxilline from Penicillium paxilli, aflatrem from Aspergillus flavus, and lolitrems from Neotyphodium lolii, all of which act as tremorgens in mammals. Lolitrems from endophytes like Neotyphodium help to protect the fungal–grass symbiosis from grazing animals (Gallagher et al. 1981). Genes encoding enzymes for the biosynthesis of lolitrems, paxilline, and aflatrem have been isolated and, similar to other secondary metabolite genes, were found to be clustered in the genomes of the corresponding fungi (Young et al. 2001, 2005, 2009; Zhang et al. 2004; Young et al. 2006; Nicholson et al. 2009). Analysis of the gene content of the clusters as well as analysis of mutants in several biosynthesis genes led to the identification of four genes that are necessary for the production of paspaline, a common intermediate of indole-diterpene biosynthesis in fungi (Saikia et al. 2006, 2008). Downstream steps of the biosynthesis vary in different fungal species, and so does the genetic organization of the indole-terpene gene clusters. Recombination events, possibly aided by transposon activity, are hypothesized to

have caused gene duplication and loss, similar to events leading to high interspecies diversity of other fungal secondary metabolite gene clusters (Saika et al. 2008).

Another important group of terpenoid fungal secondary metabolites are the trichothecenes (Kimura et al. 2007). They are produced by several Fusarium species and a number of other fungi from the order of Hypocreales. Some trichothecenes, for example, deoxynivalenol, act as phytotoxins towards the host plants of phytopathogenic Fusarium species (Proctor et al. 1995). In addition, many of them are potent mycotoxins that affect animals through consumption of contaminated agricultural products. Studies of mutant strains of F. sporotrichioides that are no longer able to produce the trichothecene T-2 toxin led to the identification of several clustered genes that are required for the biosynthesis of T-2 toxin and other trichothecenes (Hohn and Beremand 1989; Hohn et al. 1993; Kimura et al. 2003; Brown et al. 2004; Kimura et al. 2007). In addition to this large cluster of trichothecene biosynthesis genes, two other genetic loci containing one or two genes each contribute to this biosynthetic pathway. Phylogenomics analyses indicate that genes were relocated from the two smaller loci to the larger trichothecene gene cluster during evolution (Proctor et al. 2009). As in clusters of other secondary metabolite genes, gene duplications as well as loss of function mutations are hypothesized to play a role in the evolution of the trichothecene clusters, whereas so far there is no evidence for horizontal gene transfer among different members of the Hypocreales (Kimura et al. 2007; Proctor et al. 2009). Whether there are trichothecene biosynthesis genes present in other fungal groups remains to be elucidated.

VI. Melanins Melanins, as mentioned above, can be synthesized via various pathways and are described as a large group of diverse substances having similar properties (Wheeler and Bell 1988; Butler and Day 1998; Henson et al. 1999; Jacobson 2000; Butler et al. 2001). They are mostly black or brown macromolecules that are insoluble in aqueous and organic solvents, resistant to concentrated acids and oxidizing agents, hydrophobic, and negatively charged. Generally, melanins are formed


by oxidative polymerization of phenolic or indolic compounds (Langfelder et al. 2003). Different functions have been described for fungal melanin, for example, protection from various environmental stresses and a role in pathogenicity (see Section VI.C). In fungi, at least four different melanin biosynthesis pathways can be found. Melanin is synthesized either from tyrosine via L-3,4-dihydroxyphenylalanine (DOPA), from g-glutaminyl-3,

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4-dihydroxybenzene (GDHB), from catechol, or from 1,8-dihydroxynaphthalene (DHN; Bell and Wheeler 1986). GDHB- and catechol-derived melanin has been described to occur in several basidiomycetes (Stu¨ssi and Rast 1981; Wang et al. 1996). However, most fungi produce either DOPA or DHN melanin (Table 10.2) and thus, these two will be described in the next two sections, followed by an overview of melanin functions.

Table 10.2. Melanins in fungi. With the exception of the basidiomycete Cryptococcus neoformans, all fungi in this table are ascomycetes Species

Type of melanin

Reference

Alternaria alternata Alternaria brassicicola Ascochyta rabiei Aspergillus fumigatus Blastomyces dermatitidis Candida albicans Cladosporium carrionii Coccidioides posadasii Cochliobolus carbonum C. miyabeanus Colletotrichum lagenarium Cryptococcus neoformans Curvularia protuberata Diplocarpon rosae Diplodia gossypina Fonsecaea pedrosoi Gaeumannomyces graminis Glarea lozoyensis Guignardia mangiferae Histoplasma capsulatum Macrophomina phaseolina Magnaporthe grisea M. oryzae Nodulisporium sp. Ophiostoma floccosum O. piceae Paracoccidioides brasiliensis Penicillium marneffei Pleospora infectoria Podospora anserina Sclerotinia minor S. sclerotiorum S. trifoliorum Sclerotium cepivorum Scytalidium dimidiatum Sordaria macrospora Sporothrix schenckii Verticillium albo-atrum V. dahliae V. nigrescens V. tricorpus

DHN DHN DHN DHN DHN (DOPA)a DOPA DHN

Wheeler (1983), Kimura and Tsuge (1993) Wheeler (1983) Akamatsu et al. (2010) Langfelder et al. (1998), Tsai et al. (1998, 1999) Nosanchuk et al. (2004) Morris-Jones et al. (2005) Taylor et al. (1987) Nosanchuk et al. (2007) Wheeler (1983) Wheeler (1983) Kubo et al. (1991, 1996), Takano et al. (1995), Perpetua et al. (1996) Wang et al. (1996) Wheeler (1983) Gachomo et al. (2010) Wheeler (1983) Alviano et al. (1991), Cunha et al. (2005), Franzen et al. (2008) Caesar-Tonthat et al. (1995) Zhang et al. (2003) Suryanarayanan et al. (2004) Nosanchuk et al. (2002) Wheeler (1983) Chumley and Valent (1990), Vidal-Cros et al. (1994) Motoyama et al. (1998) Fulton et al. (1999) Tanguay et al. (2006) Tanguay et al. (2006) Gomez et al. (2001), da Silva et al. (2006) Youngchim et al. (2005) Wheeler (1983) Coppin and Silar (2007) Wheeler (1983) Wheeler (1983), Starratt et al. (2002), Butler et al. (2009) Wheeler (1983) Wheeler (1983) Morris-Jones et al. (2004) Engh et al. (2007) Romero-Martinez et al. (2000) Wheeler (1983) Bell et al. (1976), Stipanovic and Bell (1976) Wheeler (1983) Wheeler (1983)

b

DHN DHN DHN DOPA DHN DHN DHN DHN DHN DHN DHN DHN (DOPA)a DHN DHN DHN DHN DHN DHN DHN (DOPA)a DOPA DHN DHN DHN DHN DHN DHN b

DHN DHN DHN DHN DHN DHN

a DOPA melanin was experimentally confirmed, but additional evidence hints at the formation of DHN melanin under normal growth conditions. b The type of melanin produced by this fungus has yet to be determined.

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A. DOPA Melanin DOPA melanin is synthesized by phenoloxidases, which fall into two groups: tyrosinases (EC 1.10.3.2) and laccases (catechol oxidases, EC 1.14.18.1; Bell and Wheeler 1986; Langfelder et al. 2003). Both are type-3 copper proteins that require bound copper ions to be active, but they differ in substrate specificity and overall structure (Langfelder et al. 2003; Baldrian 2006; Halaouli et al. 2006). Tyrosinases catalyze the o-hydroxylation of monophenols and the subsequent oxidation of the resulting o-diphenols into reactive o-quinones using molecular oxygen (Halaouli et al. 2006). Subsequently, o-quinones undergo non-enzymatic reactions that lead to intermediates that autooxidize spontaneously (Soler-Rivas et al. 1999). In fungi, L-tyrosine is oxidized to L-dopaquinone in a two-step reaction mechanism and subsequently oxidized to L-dopachrome (Sanchez-Ferrer et al. 1995; Langfelder et al. 2003). Additional hydroxylation and carboxylation reactions lead to dihydroxyindoles that polymerize to DOPA melanin (Ozeki et al. 1997a, b; Williamson et al. 1998). The production of DOPA melanin via tyrosinases in fungi is very similar to the production of melanin in mammals. In contrast to mammals, however, fungal DOPA melanin does not contain any nitrogen (Bell and Wheeler 1986; Riley 1997; Langfelder et al. 2003). Laccases represent the second group of phenoloxidases involved in DOPA melanin synthesis. They typically catalyze the oxidation of p-hydroxyphenols, which in fungal melanin production is the one-step oxidation of L-DOPA to dopaquinone (Langfelder et al. 2003; Baldrian 2006). Fungal laccases are involved not only in pigmentation, but also in morphogenesis, pathogenic interactions, stress defense, and lignin degradation in wood-rotting fungi (Thurston 1994). Phenoloxidases occur in all phyla, indicating their early origin (Jaenicke and Decker 2004). Tyrosinases are widely distributed and, besides fungi, occur in mammals, insects, plants, and bacteria and are involved in, for example, wound healing and pigmentation (Claus and Decker 2006; Halaouli et al. 2006; Dittmer and Kanost 2010). Fungal tyrosinases cluster in groups for basidiomycetes and ascomycetes, and comparison of amino acid sequences highlights the highly conserved copper-binding domains (Halaouli et al. 2006). Laccases occur in filamentous ascomycetes

and in basidiomycetes, but have never been described for zygomycetes or chytridiomycetes (Baldrian 2006). Besides, they have been found in plants, where they are involved in lignin formation, in some bacteria, archea, and, recently, in insects (Ranocha et al. 2002; Koschorreck et al. 2009; Dittmer and Kanost 2010; Uthandi et al. 2010). Caution is reasonable when testing a fungus for the production of DOPA melanin, because it can occur extracellularly if adequate substrates are present in the environment that can be oxidized by secreted phenoloxidases and can then autooxidize (Butler and Day 1998). Several fungi have been described to form DOPA melanin when supplied with L-DOPA or tyrosine, but have been shown to synthesize DHN melanin under normal conditions, for example, the black yeast Phaeococcomyces and the filamentous ascomycete Verticillium dahliae (Wheeler et al. 1978; Bell and Wheeler 1986; Butler et al. 1989). There is one model system for which DOPA melanin synthesis has been described in detail, and that is the basidiomycete Cryptococcus neoformans (Polacheck and Kwon-Chung 1988; Williamson et al. 1998; Casadevall et al. 2000; Karkowska-Kuleta et al. 2009). Two paralogous cell wall-anchored laccases, CNLAC1 and CNLAC2, are crucial for melanin production which is only possible if exogenous substrate is supplied, leading to different types of DOPA melanin (Zhu and Williamson 2004). Interestingly, melanin precursors can obviously be supplied by bacteria that grow in close proximity to the fungus (Frases et al. 2006, 2007). This dependence on exogenous substrate is clearly different from DHN melanin synthesis, which is described in the next section.

B. DHN Melanin DHN melanin is widely distributed in the class of ascomycetes and its production requires at least four enzymatic steps (Bell and Wheeler 1986; Langfelder et al. 2003). First, a type I polyketide synthase (see Section II.A) catalyzes the formation of 1,3,6,8-tetrahydroxynaphthalene (1,3,6,8-THN), a pentaketide, from acetyl-CoA or malonyl-CoA precursors. 1,3,6,8-THN is subsequently reduced to scytalone which is dehydrated by scytalone dehydratase to 1,3,8-trihydroxynaphtalene (1,3,8-THN). This again is reduced to vermelone which is then


dehydrated, possibly again by scytalone dehydratase, to 1,8-dihydroxynaphthalene (DHN) for which the pathway is named. Subsequent polymerization of DHN is hypothesized to occur spontaneously in the presence of molecular oxygen, or, alternatively, to be catalyzed by a phenoloxidase (Bell and Wheeler 1986; Butler and Day 1998). It is supposed that two different enzymes carry out the reduction from 1,3,6,8-THN to scytalone and from 1,3,8-THN to vermelone. Indeed, two hydroxynaphtalene reductases were identified in many fungi, for example, in Sordaria macrospora and Magnaporthe grisea (Vidal-Cros et al. 1994; Thompson et al. 2000; Engh et al. 2007). However, for other fungi like Bipolaris oryzae, only one hydroxynaphtalene reductase gene has been found so far (Kihara et al. 2004). In any case, the reduction steps of the DHN pathway can be specifically inhibited by tricyclazole, leading to the accumulation of reddish intermediates, and this inhibition is very often used as a first test for DHN melanin production (Tokousbalides and Sisler 1979; Bell and Wheeler 1986; Romero-Martinez et al. 2000; Engh et al. 2007). DHN melanin biosynthesis (DMB) genes occur distributed throughout the genome in some species and clustered or partially clustered in others. Two prominent examples for DMB gene clusters are those of Alternaria alternata and Aspergillus fumigatus (Kimura and Tsuge 1993; Langfelder et al. 1998; Tsai et al. 1998, 1999; Brakhage et al. 1999). The Alternaria alternata cluster contains three genes, alm, brm1 and brm2, encoding a polyketide synthase, a scytalone dehydratase and a reductase, respectively (Kimura and Tsuge 1993). The three genes are located within a 30-kb genomic region and are expressed synchronously with mycelial melanization. In Aspergillus fumigatus, DMB is rather more complex than in other fungi and this can already be deduced from the gene cluster which here consists of six genes (Brakhage et al. 1999; Tsai et al. 1999). While three genes, pksP/alb1, arp2 and arp1 code for polyketide synthase, 1,3,6,8-THN reductase and scytalone dehydratase, respectively (Langfelder et al. 1998; Tsai et al. 1998, 1999), ayg1, abr1 and abr2 serve novel functions. It was demonstrated that the polyketide synthase PksP produces an unusual heptaketide which is then shortened to 1,3,6,8-THN by Ayg1 (Tsai et al. 2001, Fujii et al. 2004). Abr1 and Abr2 are a multicopper oxidase and a laccase that probably perform additional

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oxidation steps in DMB, leading to the typical blue-green pigment found in Aspergillus conidia (Tsai et al. 1999; Sugareva et al. 2006). Recently, the DMB gene cluster of Penicillium marneffei has been described which is highly similar to that of A. fumigatus and other related fungi. Several gene rearrangements indicate that the cluster was acquired by a common ancestor of Aspergilli and Penicilli (Woo et al. 2010). The kind of pigment in these fungi differs clearly from the melanin found in more distantly related fungi and this may be why these other fungi have less DMB genes than A. fumigatus and its close relatives.

Interestingly, it was found that A. alternata DMB genes, which are clustered, can restore melanin-deficient mutants of Colletotrichum lagenarium and M. grisea, species with unlinked DMB genes (Chumley and Valent 1990; Kubo et al. 1991; Kawamura et al. 1997; Takano et al. 1997). These results demonstrate that the linkage relationship and arrangement of melanin biosynthesis genes can be quite different among fungi, although the biosynthetic pathway of DHN melanin is almost identical. An intriguing aspect, however, is that melanin is deposited into different structures in A. alternata (conidia) versus C. lagenarium and M. grisea (appressoria). Thus, it may serve different functions and its biosynthesis may be differently regulated and this in turn could be related to the different arrangement of the genes (Kawamura et al. 1997; Takano et al. 1997; see Section VI.C).

C. Biological Functions of Fungal Melanins Melanin is thought to protect fungi from various environmental stresses such as UV radiation, desiccation, the host immune system, enzymatic lysis, predators or even radioactivity (Bell and Wheeler 1986; Butler and Day 1998; Jacobson 2000; Gomez and Nosanchuk 2003; Nosanchuk and Casadevall 2003, 2006; Karkowska-Kuleta et al. 2009). Melanin is one of the traits supposed to be responsible for the high diversity of fungi and their ability to exploit extreme environments (Gostincar et al. 2010). For example, conidia and pycnidia of melanin-deficient A. alternata and A. rabiei strains, respectively, were more sensitive to UV light than wild-type structures (Kawamura et al. 1999; Akamatsu et al. 2010). Singaravelan and colleagues (2008) showed a direct connection

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between the amount of solar UV radiation and conidial melanin concentration in A. niger. Intriguingly, melanized fungi are able to grow in the presence of radionuclides and have been described to grow in the highly contaminated Chernobyl reactor 4 and to increase in the surroundings of the explosion site (Zhdanova et al. 1991; Mironenko et al. 2000). It has been found that melanized fungi are highly radiation-resistant under experimental conditions, and some fungi actually display radiotropism, that is, the positive growth response of fungi towards a radiation source (Dadachova and Casadevall 2008; Dighton et al. 2008). Apparently, not only does melanin shield fungal cells from radiation damage, but ionizing radiation changes the electronic properties of melanin and leads to enhanced growth, indicating that melanin can function in energy capture and utilization (Dadachova et al. 2007). Much work has focused on the role of melanin in human pathogenic fungi (Jacobson 2000; Nosanchuk and Casadevall 2006; Taborda et al. 2008; Liu and Nizet 2009). Often, the infecting spores of human pathogenic fungi produce melanin as protection against the host immune response, such as phagocytosis, killing by the host cell, and antioxidants (Nosanchuk and Casadevall 2006). For A. fumigatus it has been demonstrated that melanin deficiency leads to a change of conidial surface structure. While wild-type conidia have cell wall protrusions and appear echinulated, conidial color mutants and conidia of a pks knockout strain produce smooth conidia that show an altered cell wall structure, are more susceptible to killing by oxidants and therefore are less virulent (Jahn et al. 1997; Langfelder et al. 1998; Jahn et al. 2002; Pihet et al. 2009). It has been suggested that melanin on the conidial surface may mask the host cytokine response (Chai et al. 2009). Similar results were obtained for the pathogenic basidiomycete C. neoformans. Here, melanins in the cell wall protect against phagocytosis and confer resistance against endogenous antifungal peptides as well as pharmacological agents (for a review, see Liu and Nizet 2009). Melanin is also an important feature of plant pathogenic fungi, especially leaf pathogens (for a review, see Henson et al. 1999). While some fungi penetrate leaf cells through stomata, several fungi are known to produce appressoria (Mendgen et al. 1996). In M. grisea and C. lagenarium, melanization of appressoria is essential for penetration of

the host plant cell, while for A. alternata, melanization of conidia and hyphae could not be directly linked to pathogenicity (Kubo et al. 1982; Tanabe et al. 1988; Kubo et al. 1991; Kimura and Tsuge 1993; Howard and Valent 1996; Kawamura et al. 1999). Extensive studies on M. grisea showed that melanin in the appressorial cell wall is needed to withstand a high internal turgor pressure of 8.0 MPa, and that penetration pegs are pushed into the host cell through a nonmelanized pore (Howard et al. 1991 Howard and Valent 1996; Money and Howard 1996; de Jong et al. 1997). Melanin-deficient mutants of M. grisea are unable to generate high turgor pressure and are nonpathogenic (Howard and Ferrari 1989; Chumley and Valent 1990). Pigmented appressoria have been shown to be essential for plant infection in other fungi, for example, Diplocarpon rosae and several Colletotrichum species; thus, turgor pressure-mediated plant infection seems to be a general scheme of plant pathogenic fungi with melanized appressoria (Kubo et al. 1982; Mendgen et al. 1996; Fujihara et al. 2010; Gachomo et al. 2010).

Some data hint at a function for melanin in sexual development. Color mutants of Podospora anserina and Ophiostoma piliferum have been described to be defective in the production of sexual fruiting bodies (Esser 1966, 1968; Zimmerman et al. 1995), and several studies imply a correlation between melanin biosynthesis or melanin biosynthesis gene expression and stages of sexual reproduction in N. crassa, S. macrospora and Tuber species (Hirsch 1954; Prade et al. 1984; Ragnelli et al. 1992; Engh et al. 2007). However, additional studies are necessary to reveal the components linking both processes.

VII. Conclusions Fungi produce a vast spectrum of natural products, and with the availability of genome sequences, it quickly became apparent that this spectrum is even greater than previously expected. Genomic mining and phylogenetic studies have already yielded novel insights into the evolution of genes for secondary metabolism and have facilitated the identification of new metabolites. It is now clear that many fungi have the genetic potential to produce dozens of metabolites per species, many of which may be species-specific.


Mechanisms involved in this high diversity are recombination events including duplications and deletions as well as horizontal gene transfer. Genes for secondary metabolites are generally fast-evolving, often clustered, and co-regulated through common transcription factors or at the level of chromatin organization. What is not yet known is the biological function for the majority of metabolites and biosynthesis genes; however, several detailed studies over recent decades have already indicated that the roles of these metabolites for the fungal lifestyle are as diverse as their biochemistry and include functions ranging from core cellular processes to development, pathogenicity factors, and protective agents. Combined bioinformatics/genetic/physiological approaches will help to further clarify the origins and roles of fungal secondary metabolites. Acknowledgements We would like to thank Prof. Dr. Ulrich Ku¨ck for his support. Funding for our work comes from the German Science Foundation (Deutsche Forschungsgemeinschaft, DFG, grant NO 407/2-1) and the Protein Research Department of the Ruhr-University Bochum.

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Wheeler MH (1983) Comparisons of fungal melanin biosynthesis in ascomycetous, imperfect and basidiomycetous fungi. Trans Br Mycol Soc 81:29–36 Wheeler MH, Bell AA (1988) Melanins and their importance in pathogenic fungi. Curr Top Med Mycol 2: 338–387 Wilkinson HH, Siegel MR, Blankenship JD, Mallory AC, Bush LP, Schardl C (2000) Contribution of fungal loline alkaloids to protection from aphids in a grass-endophyte mutualism. Mol Plant Microbe Interact 13:1027–1033 Williamson PR, Wakamatsu K, Ito S (1998) Melanin biosynthesis in Cryptococcus neoformans. J Bacteriol 180: 1570–1572 Wolf JC, Mirocha CJ (1973) Regulation of sexual reproduction in Gibberella zeae (Fusarium roseum “Graminearum”) by F-2 (Zearalenone). Can J Microbiol 19:725–734 Woloshuk CP, Foutz KR, Brewer JF, Bhatnagar D, Cleveland TE, Payne GA (1994) Molecular characterization of aflR, a regulatory locus for aflatoxin biosynthesis. Appl Env Microbiol 60:2408–2414 Woo PCY, Tam EWT, Chong KTK, Cai JJ, Tung ETK, Ngan AHY, Lau SKP, Yuen KY (2010) High diversity of polyketide synthase genes and the melanin biosynthesis gene cluster in Penicillium marneffei. FEBS J 277:3750–3758 Xu J, Saunders CW, Hu P, Grant RA, Boekhout T, Kuramae EE, Kronstad JW, DeAngelis YM, Reeder NL, Johnstone KR, Leland M, Fieno AM, Begley WM, Sun Y, Lacey MP, Chaudhary T, Keough T, Chu L, Sears R, Yuan B, Dawson TL (2007) Dandruff-associated Malassezia genomes reveal convergent and divergent virulence traits shared with plant and human fungal pathogens. Proc Nat Acad Sci USA 104:18730–18735 Yamaguchi S (2008) Gibberellin metabolism and its regulation. Annu Rev Plant Biol 59:225–251 Yang G, Rose MS, Turgeon BG, Yoder OC (1996) A polyketide synthase is required for fungal virulence and production of the polyketide T-toxin. Plant Cell 8: 2139–2150 Young C, McMillan L, Telfer E, Scott B (2001) Molecular cloning and genetic analysis of an indole-diterpene gene cluster from Penicillium paxilli. Mol Microbiol 39:754–764 Young CA, Bryant MK, Christensen MJ, Tapper BA, Bryan GT, Scott B (2005) Molecular cloning and genetic analysis of a symbiosis-expressed gene cluster for lolitrem biosynthesis from a mutualistic endophyte of perennial ryegrass. Mol Genet Genomics 274:13–29 Young CA, Felitti S, Shields K, Spangenberg G, Johnson RD, Bryan GT, Saikia S, Scott B (2006) A complex gene cluster for indole-diterpene biosynthesis in the grass endophyte Neotyphodium lolii. Fungal Genet Biol 43:679–693 Young CA, Tapper BA, May K, Moon CD, Schardl CL, Scott B (2009) Indole-diterpene biosynthetic capability of Epichloe endophytes as predicted by ltm gene analysis. Appl Environ Microbiol 75:2200–2211

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11

Carbonic Anhydrases in Fungi and Fungal-Like Organisms – Functional Distribution and Evolution of a Gene Family

SKANDER ELLEUCHE1

CONTENTS I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Distribution of Carbonic Anhydrases in Fungal-Like Organisms and ‘Basal Fungi’ . . . . . . . A. a-, b- and g-Carbonic Anhydrases in Phytophthora infestans . . . . . . . . . . . . . . . . . . . . . B. b-Class Carbonic Anhydrases in the Social Amoeba Dictyostelium discoideum . . . . . . . . . . . . C. Carbonic Anhydrases in ‘Basal Fungi’ . . . . . . . . III. Diversity of Carbonic Anhydrases in Ascomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. a-Carbonic Anhydrases. . . . . . . . . . . . . . . . . . . . . . . . B. b-Carbonic Anhydrases . . . . . . . . . . . . . . . . . . . . . . . . 1. Plant-Like b-Carbonic Anhydrases . . . . . . . . 2. Cab-Like b-Carbonic Anhydrases . . . . . . . . . IV. Carbonic Anhydrases in Basidiomycetes . . . . . . . . V. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

257 258 259 260 261 263 263 264 266 269 270 270 271

I. Introduction Carbonic anhydrases (EC 4.2.1.1) form a highly diverse family of enzymes. Members of this family are distributed among all three domains of life (Bacteria, Archaea, Eukaryotes). So far, five evolutionary unrelated classes (a, b, g, d, z) with independent ancestors have been described. Mammals solely contain isozymes of the a-class, a group that has also been identified in prokaryotes, plants and fungi. The b-class has been identified in prokaryotes, fungi and plants, but not in mammals. g-carbonic anhydrases were predominantly found in archaeal species and plant mitochondria, while the novel d- and z-classes have been exclusively identified in marine diatoms, so far (Lane et al. 2005; Park et al. 2007; Supuran 2008a). 1

Technische Universita¨t Hamburg-Harburg, Institut fu¨r Technische Mikrobiologie, Kasernenstr. 12, 21073 Hamburg, Germany; e-mail: [email protected]

The definition of a gene family describes a group of genes that have descended from a common ancestor and retained similar sequences that confer structural similarities as well as physiological functions (Demuth and Hahn 2009). Two genes that have undergone duplication from a pre-existing gene within the genome of a single species are defined as paralogues. Comparative genomics identify gene copies, which have diverged by phylogenetic splitting of the organisms. These genes are called orthologues (Gogarten and Olendzenski 1999).

Although all classes of carbonic anhydrases are remarkably different at the sequence level, the structures of the metal-coordinating (zinc ion in most cases, cadmium ion only in z-carbonic anhydrases) catalytic sites show a fairly high degree of identity (Tripp et al. 2001; Xu et al. 2008). The g-carbonic anhydrase Cam from the archaeon Methanosarcina thermophila was even shown to coordinate zinc, cobalt or iron in its catalytic centre (Zimmerman et al. 2004). Nevertheless, members of all groups of carbonic anhydrases have been proven to rapidly catalyse the naturally balanced chemical reaction, namely the interconversion of carbon dioxide and bicarbonate: CO2 þ H2O , HCO3 þ Hþ. Carbon dioxide is a ubiquitous gaseous molecule and the end product of cellular respiration. Bicarbonate represents an important biological substrate that is involved in multiple metabolic reactions (biosynthesis of arginine, uracile or fatty acids), in the degradation of toxic cyanate and in the activation of adenylyl cyclase (Aguilera et al. 2005; Hetherington and Raven 2005; Klengel et al. 2005; Mogensen et al. 2006; Elleuche and Po¨ggeler 2008; Mogensen and Mu¨hlschlegel 2008). In numerous bacterial species, carbonic anhydrases have been found that are either genetically fused to anion transporters or are encoded within operons together with anion transporters. The anion transportres are members of the SulP family which are suspected to be capable of transporting bicarbonate ions (Felce and Saier 2004).

Evolution of Fungi and Fungal-Like Organisms, The Mycota XIV S. Po¨ggeler and J. Wo¨stemeyer (Eds.) © Springer-Verlag Berlin Heidelberg 2011

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In ambient air, 0.033% of the atmospheric gases are represented by carbon dioxide. An immense higher concentration of about 5% is found in respiration active tissues (Bahn and Mu¨hlschlegel 2006). The fact that the spontaneous interconversion of carbon dioxide to bicarbonate is rather slow, but the demand of the important biological substrate bicarbonate is high, necessitates a defined regulation of this interconversion and an immense acceleration up to 10000-fold (Wistrand 1981; Jones 2008). The fine-tuned regulation network caused a highly diverse evolution of the different enzyme classes and led to the development of multiple carbonic anhydrase isoforms in various organisms. In the past years, 16 different organ- and tissue-specific a-carbonic anhydrases have been identified in mammals. They are also located to different subcellular compartments, including the cytosol and mitochondria, and include also membrane-associated and secreted isoforms (Kivela¨ et al. 2005; Supuran 2008a). The green algae Chlamydomonas reinhardtii encodes three carbonic anhydrases of the a-class and six of the b-class, while eight a-, six b- and three g-class carbonic anhydrases have been investigated in the model plant Arabidopsis thaliana. In plants and algae, isoforms have been found to be located to the cytosol, the mitochondria and the chloroplasts and secreted enzymes have also been described (Mitra et al. 2004; Parisi et al. 2004; Fabre et al. 2007; Moroney and Ynalvez 2007; Ynalvez et al. 2008). Multiple carbonic anhydrases of the a- and b-class are also common in fungal species. Physiological functions of carbonic anhydrases have been solely investigated in the yeasts Saccharomyces cerevisiae and Candida albicans, in the basidiomycetous human pathogen Cryptococcus neoformans and very recently in the filamentous ascomycetes Aspergillus nidulans, A. fumigatus and Sordaria macrospora (Go¨tz et al. 1999; Bahn et al. 2005; Klengel et al. 2005; Mogensen et al. 2006; Elleuche and Po¨ggeler 2009a; Han et al. 2010; Kim et al. 2010). All of these described isozymes belong to the class of b-carbonic anhydrases and exhibit diverse functions in fungi (Elleuche and Po¨ggeler 2010). Interestingly, this enzyme family is highly diverse in fungal organisms and often consists of multiple members. The syntrophic bacterium Symbiobacterium thermophilum and several related micro-organisms are the only

species, known to date, that have lost their carbonic anhydrases during the course of evolution. They are able to grow on the high level of CO2 and the spontaneous interconversion product HCO3 in their environment, which is generated by other bacteria (Nishida et al. 2009).

The recent and continuous proliferation of completed fungal genome sequences available at public databases provides an insight into the phylogenetic evolution of genes and proteins within the fungal kingdom. These genomes offer the opportunity to compare the gene content of different fungal orders and to understand underlying evolutionary mechanisms. Carbonic anhydrases are encoded by all fungal genomes sequenced to date and the inventory of novel enzymes during evolution might reflect the individual requirements of different fungal life styles. All known fungal carbonic anhydrases belong either to the group of a- or b-class enzymes, while members of the g-, d- and z-classes have not been identified so far (Elleuche and Po¨ggeler 2009b). In addition, chimeric-types of carbonic anhydrases fused to bicarbonate transporters seem not to exist as well (Felce and Saier 2004). Interestingly, the number and sequence of already described and putative isozymes is highly diverse in fungi, resulting from gene loss and duplication events. This chapter focuses on the distribution and evolution of the gene family of carbonic anhydrases in non-fungal organisms, the ‘Basal Fungi’ as well as in ascomycetes and basidiomycetes. The sequence comparison of isoforms and their predicted cellular localization patterns, in combination with the data of some proteins, which have been already characterized, will shed some light on their physiological functions. Especially the pervasiveness of gene duplication, gene loss and neofunctionalization events seem to play a predominant role in the course of evolution of carbonic anhydrases.

II. Distribution of Carbonic Anhydrases in Fungal-Like Organisms and ‘Basal Fungi’ The number of carbonic anhydrases in the genomes from the non-fungal organisms Phytophthora infestans and Dictyostelium discoideum and from the ‘Basal Fungi’ Allomyces macrogynus, Batrachochytrium dendrobaditis, Spizellomyces

Carbonic Anhydrases in Fungi and Fungal-Like Organisms

punctatus, Phycomyces blakesleeanus and Rhizopus oryzae was evaluated and these enzymes were grouped into different classes. Since a comprehensive analysis is beyond the scope of this review, it will make no claim to be complete.

A. a-, b- and g-Carbonic Anhydrases in Phytophthora infestans P. infestans is a water mould straminipilous organism, which belongs to the phylum Oomycota within the ‘Kingdom’ Chromalveolata. The P. infestans 240-Mbp genome is to date the largest genome within the Chromalveolata. The genome expansion has been discussed to be the result of proliferation of repetitive DNA without a wholegenome duplication event. Only specific gene families (e.g., disease effector gene families) were shown to have undergone rapid expansions (Haas et al. 2009). In this context, it might be not surprising that the family of carbonic anhydrases is highly diverse in the P. infestans genome. Table 11.1 lists the isoforms of class a, b and g identified by screening the genome using BLASTP analyses with fungal carbonic anhydrases (unpublished data). Thirteen of the 17 identified carbonic anhydrase encoding genes are predicted to encode enzymes belonging to the a-class. All of these isoforms contain the catalytic histidine triad within a highly conserved catalytic domain predicted to

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be important for zinc coordination (Fig. 11.1A). The Q-x-H-x-H motif of the catalytic region is conserved from bacteria to fungi and has been also found in mammalian and algal carbonic anhydrases (Ishida et al. 1993; Zimin et al. 2009). In contrast to fungal sequences, the third histidine is embedded into a highly conserved H-F-V-H sequence motif that is found in all P. infestans a-carbonic anhydrases as well as in enzymes of prokaryotic origin (Chirica et al. 1997; Barnett et al. 2001; Elleuche and Po¨ggeler 2009b). Two of these enzymes are 100% identical at the protein level (99.9% at the nucleotide level), but are annotated as different genes (PITG_17848.1 and PITG_18284.1) in the database at the BROAD institute (Project: Fungal Genome Initiative), indicating a very recent gene duplication event or a false assembled or annotated genome region. The remaining proteins are highly diverse in sequence similarity (0–96.9%) and vary considerably in size (148–458 amino acids). The a-class carbonic anhydrase encoded by gene PITG_08497.1 seems to be the most distinct isoform. The flanking sequences of the catalytic region of PITG_08497.1 are so divergent that any try to generate a global alignment is insignificant. Calculation with TargetP program predicts seven (including the 100% identical proteins) a-carbonic anhydrases to be secreted, while five isoforms probably are cytosolic enzymes (Emanuelsson et al. 2007). Furthermore, a weak

Table 11.1. a-, b- and g-carbonic anhydrases in ‘non fungal’ organisms Phylum

Organism

Type

Size (aa)

Subcellular localization

Database description

Oomycota

Phytophthora infestans

a-Class

228 148 219 244 269 269 458 269 189 170 272 183 240 315 356 325 251 276 274

Cytosolic Cytosolic Cytosolic Secreted Secreted Secreted Secreted Secreted Secreted Cytosolic Secreted Cytosolic Mitochondrial Cytosolic Cytosolic Mitochondrial Cytosolic Cytosolic Cytosolic

PITG_21981.1 PITG_17845.1 PITG_17844.1 PITG_17837.1 PITG_18284.1 PITG_17848.1 PITG_17846.1 PITG_17842.1 PITG_21980.1 PITG_17808.1 PITG_14412.1 PITG_01908.1 PITG_08497.1 PITG_00674.1 PITG_00673.1 PITG_00682.1 PITG_19282.1 XP_646739.1 XP_644170.1

b-Class

Eumycetozoa

Dictyostelium discoideum

g-Class b-Class

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Fig. 11.1. Multiple sequence alignments of a- and b-carbonic anhydrases in the oomycoteous organism Phytophthora infestans. (A) Amino acid sequences of the a-carbonic anhydrases of P. infestans were aligned to maximize similarities (Thompson et al. 1997). The region around the zinc-coordinating residues (amino acid residues 77–101; PITG_21981.1 numbering) is given with identical amino acids in all sequenced boxed in black. Gaps in the sequence of PITG_08497.1 are introduced to

maximize similarity. (B) Multiple sequence alignment of the b-carbonic anhydrases of P. infestans. Identical amino acid residues are boxed in black and gaps indicate 13 highly conserved residues that have been omitted for better presentation of the catalytic domain. Sizes (in amino acids) and prediction of subcellular localization are indicated at right. Arrows indicate the position of the zinc-coordinating amino acid residues. Abbreviations for TargetP: -C- cytosolic, -S- secreted, -M- mitochondrial

mitochondrial target sequence has been predicted at the N-terminal part of a-class isozyme encoded by the open reading frame PITG_08497.1. Beside a newly identified carbonic anhydrase within the genome of the zygomycetous fungus Allomyces macrogynus (see Section II.C), this is the first indication of a putative mitochondrial a-carbonic anhydrase encoded by fungal or fungal-like genomes, but this has to be proven experimentally. In contrast to fungi and fungal-like species, mitochondrial a-carbonic anhydrases are widespread among mammalian organisms (Shah et al. 2000; Supuran 2008a). Three carbonic anhydrases of the plant-type b-class are encoded within the genome of P. infestans. They exhibit similarities between 53.8 and 60.0% and are 315, 325 and 356 amino acid residues in length (Table 11.1). According to in silico analysis, PITG_00682.1 encodes an isoform that is translocated into the mitochondria, while PITG_00673.1 and PITG_00674.1 encode cytoplasmic proteins (Fig. 11.1B). Moreover, phylogenetic studies revealed that the P. infestans b-class

carbonic anhydrases seem to be related to the CAS2-group from filamentous fungi (Elleuche and Po¨ggeler 2009b). Furthermore, the open reading frame PITG_ 19282.1 encodes the first putative g-carbonic anhydrase encoded in fungi or fungal-like organisms, so far. It has been annotated as a conserved hypothetical protein and was identified by means of gene index search using the term carbonic anhydrase as query. ClustalX alignment and Superfamily prediction assigned PITG_19282.1 to the g-carbonic anhydrases (data not shown). The identification of this first g-class protein in fungallike species quickly expands the distribution of this group of carbonic anhydrases to a further level.

B. b-Class Carbonic Anhydrases in the Social Amoeba Dictyostelium discoideum In the social amoeba D. discoideum, cyclic adenosine monophosphate (cAMP) plays a crucial role


in differentiation, development and cell movement during its life cycle, which necessitates defined spatial and temporal regulation of cAMP synthesis (Saran et al. 2002; Gregor et al. 2010; see Chapter 4 in this volume). Activation of the enzyme adenylyl cyclase leads to the production of cAMP. There are two distinct classes of cAMPgenerating adenylyl cyclases existing in mammals, a well known group of transmembrane enzymes and a recently described soluble enzyme class that is directly regulated by bicarbonate and more closely related to prokaryotic isoforms, while the transmembrane adenylyl cyclases are G proteindependent (Chen et al. 2000). A homologue of the bicarbonate activated soluble adenylyl cyclases from prokaryotes has been also identified in the forest soil-living amoeba D. discoideum (Chen et al. 2000). This coherence makes the presence of a bicarbonate producing carbonic anhydrase in this micro-organism more conceivable and indeed two putative isozymes of the b-class (XP_646739.1 and XP_644170.1) can be found within the fully sequenced genome of strain AX4 (Eichinger et al. 2005). Both are predicted to be located to the cytosol and are highly similar in length (276 and 274 amino acids) and in sequence composition (75.1% identity in 269 amino acids overlap), indicating a common ancestor (Table 11.1). A homologue (AAB06760.1) to XP_646739.1 (99.3% identity) from another D. discoideum strain has also been shown to be closely related to prokaryotic b-class carbonic anhydrases (Smith et al. 1999). It would be interesting to investigate the enzyme family of carbonic anhydrases in the Amoebozoa, since these organisms are located at the earliest branches from the last common ancestor of all eukaryotes in the evolutionary tree of life (Eichinger et al. 2005).

C. Carbonic Anhydrases in ‘Basal Fungi’ In recent years, the taxonomy of the ‘basal fungi’ has been extensively modified in a process resulting in the replacement of the two established phyla Zygomycota and Chytridiomycota by eight novel groups (Chytridiomycota, Neocallimastigomycota, Blastocladiomycota, Glomeromycota, Mucoromycotina, Entomophthoromycotina, Zoopagomycotina, Kickxellomycotina), to which the phylum Microsporidia was added (Hibbett et al. 2007; see Chapters 1, 2 in this volume). According

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to James et al. (2006), the Microsporidia seem to be closely related to the chytrids. A short overview of putative carbonic anhydrases in the ‘basal fungi’ is given below, although their enzyme activity is unknown so far. Multiple carbonic anhydrases have been identified in the chytrid Allomyces macrogynus (Blastocladiomycota), making this fungus a good example for gene proliferation in this phylum (unpublished data). Using different genome BLAST approaches, nine enzymes have been identified in total. According to their primary structure, three of them were assigned to the a-class and six to the b-class (Table 11.2). Such a high number of carbonic anhydrases makes it seem likely that some of the proteins may have lost their functionality during evolution; however sequence alignments revealed that all zinc-coordinating amino acid residues are highly conserved in the A. macrogynus carbonic anhydrases. In contrast to these results, numerous non-functional a-carbonic anhydrases and pseudogenes have been identified in mammals and in the plant A. thaliana (Fabre et al. 2007; Supuran 2008a). It has been hypothesized that young duplicated genes often become pseudogenes (Demuth and Hahn 2009). The three identified a-class carbonic anhydrases of A. macrogynus are encoded by the genes AMAG_06767.1, AMAG_15928.1 and AMAG_ 15929.1 and the conservation among all the sequences is clearly evident (Fig. 11.2). While the deduced proteins from AMAG_06767.1 and AMAG_15929.1 are predicted to be secreted, AMAG_15928.1 seems to be translocated into the mitochondria. Interestingly, the secreted isoform AMAG_06767.1 and the mitochondrial enzyme share a sequence identity of 90.0% in 260 amino acids overlap of the C-terminus, while amino acids 1–63 of AMAG_06767.1 are highly dissimilar to the N-terminus of AMAG_15928.1. The second secreted enzyme AMAG_15929.1 is 66.7% identical to the mitochondrial carbonic anhydrase and 90.0% to AMAG_06767.1, but has a shorter C-terminus and displays multiple mutations near its catalytic domain, which differentiate this enzyme from AMAG_06767.1 and AMAG_ 15928.1. The N-terminus of AMAG_15929.1 including the predicted secretion signal shares a rather low degree of identity (49.2%) to AMAG_ 06767.1 and exhibits an internal deletion of six amino acids directly located after the signal

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Table 11.2. a- and b-carbonic anhydrases in the ‘basal fungi’ Phylum

Organism

Type

Blastocladiomycota

Allomyces macrogynus a-Class b-Class

Chytridiomycota

Batrachochytrium dendrobatidis Spizellomyces punctatus

Zygomycota

a-Class b-Class a-Class b-Class

Size (aa) Subcellular localization Database description 322 314 286 226 220 222 250 451 438 372

Secreted Mitochondrial Secreted Cytosolic Cytosolic Cytosolic Cytosolic Mitochondrial Mitochondrial Secreted

AMAG_06767.1 AMAG_15928.1 AMAG_15929.1 AMAG_02512.1 AMAG_00999.1 AMAG_02510.1 AMAG_01001.1 AMAG_10420.1 AMAG_03148.1 BDEG_03440.1

358 242 354

Cytosolic Cytosolic Secreted

BDEG_07651.1 BDEG_05606 SPPG_04960.2

Cytosolic Cytosolic Cytosolic

SPPG_07866.2 SPPG_05997.2 Phybl2_62368

Cytosolic Cytosolic

Phybl2_184718 RO3G_10751.3

Phycomyces blakesleeanus

b-Class

374 254 263

Rhizopus oryzae

b-Class

272 375

Fig. 11.2. Multiple sequence alignments of a-class carbonic anhydrases from the zygomycete Allomyces macrogynus. A ClustalX alignment was generated with the protein sequences of AMAG_06767.1, AMAG_15928.1 and AMAG_15929.1 derived from the Fungal Genome Initiative at the Broad Institute (Thompson et al. 1997). Arrows indicate the position of histidines relevant for

zinc-coordination. Boxes indicate A. macrogynus open reading frames, with the grey box marking the region which encodes the protein part used for the alignment at the top. The signal peptides predicted for subcellular localization and secretion are marked M for mitochondrial localization and S for secreted protein

peptide. These results might indicate that the three a-carbonic anhydrases from A. macrogynus are the result of a recent gene triplication event that has been followed by the neofunctionalization of a single secreted enzyme and a modification of the subcellular localization pattern of one isoform. Furthermore, the group of b-class enzymes in A. macrogynus is also highly variable in the protein sequences and localization patterns. Four proteins are predicted to be the cytosolic enzymes,

while the open reading frames AMAG_10420.1 and AMAG_03148.1 encode putative mitochondrial carbonic anhydrases. Interestingly, these two proteins are rather large enzymes (451 and 438 amino acids, respectively) and share an overall sequence identity of 95.6%, indicating that another recent gene duplication event of a CA gene might have occurred in A. macrogynus. Both proteins are split into two parts, with the b-carbonic anhydrase domain at the C-terminus


and a large N-terminal domain (amino acids 1–226) with a so far unknown function. It would be interesting to figure out whether (i) both enzymes show identical expression profiles and share redundant functionalities or (ii) a neofunctionalization process has already been happened or (iii) one of the isozymes lost its function. Moreover, the distribution of carbonic anhydrases in A. macrogynus appears to be highly different from the distribution in the related chytrids Batrachochytrium dendrobaditis and Spizellomyces punctatus (phylum Chytridiomycota; Table 11.2). B. dendrobaditis belongs to the order Rhizophydiales and causes the disease chytridiomycosis in hundreds of amphibian species (Longcore et al. 1999; Rosenblum et al. 2010). In the genomes of these two fungal organisms, two strikingly different cytoplamic b-class carbonic anhydrases have been identified. According to BLAST analyses, these are more closely related to the isozymes of prokaryotic origin than to enzymes from ascomycetous or basidiomycetous species. Phylogenetic analysis revealed that B. dendrobaditis BDEG_07651.1 and S. punctatus SPPG_07866.2 and BDEG_05606 and SPPG_05997.2, respectively, are related, assuming a recent common ancestor in each case and a common ancestor of all four proteins earlier in (fungal) evolution. Additionally, a single secreted a-class isoforms is encoded by B. dendrobaditis and S. punctatus. However, both enzymes are strikingly different at the protein level (29.7% identity in a 236-amino-acid overlap). Carbonic anhydrase encoding genes have also been identified within the genomes of the zygomycetous fungi Phycomyces blakesleeanus and Rhizopus oryzae (subphylum Mucoromycotina; Table 11.2). P. blakesleeanus, a model to study phototropism, encodes at least two carbonic anhydrases within its recently sequenced genome. BLAST searches revealed two related b-class isozymes (72.6% identity in 215-amino-acid overlap), with no signal peptides, indicating that both open reading frames probably encode cytosolic proteins. Finally, a single 375-amino-acid carbonic anhydrase (RO3G_10751.3) with unknown function has been identified in the genome of the zygomycete R. oryzae. Interestingly, this enzyme is unique, because it contains an additional N-terminal WD40 domain, besides the typical b-class carbonic anhydrase region at the C-terminus. Using a multiple sequence alignment, it has

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been recently shown, that RO3G_10751.3 contains all amino acid residues important for zinc coordination and that it exhibits a high sequence identity at the catalytic region between 45.3 and 66.2% in comparison with b-carbonic anhydrases from ascomycetous species (Elleuche and Po¨ggeler 2009a). Continuative investigations will give deeper insights into the evolution and physiology of zygomycetous carbonic anhydrases. So far, no carbonic anhydrase homologues have been described in the ‘Basal Fungi’ of the phylum Glomeromycota, which are the closest relatives of the ascomycetes and basidiomycetes.

III. Diversity of Carbonic Anhydrases in Ascomycetes Most of the carbonic anhydrases identified in ascomycetes belong to the large group of b-class isozymes, but several a-class enzymes were also found within the subphylum Pezizomycotina, which is equivalent to the indication ‘filamentous ascomycetes’ or ‘Euascomycetes’. Beside the Pezizomycotina, two further subphyla (Taphrinomycotina, Saccharomycotina) are currently accepted within the phylum Ascomycota (Weber 2009). b-class carbonic anhydrases were found in every subphylum of the ascomycetes.

A. a-Carbonic Anhydrases The first putative a-class carbonic anhydrase was identified within the genome of Aspergillus oryzae (Bahn and Mu¨hlschlegel 2006). A comprehensive BLAST analyses using this a-carbonic anhydrase sequence as query, led to the identification of additional homologues within the ascomycetous orders Eurotiales, Sordariales, Hypocreales and Onygenales (Elleuche and Po¨ggeler 2009b). Two highly similar isozymes were identified within the genomes of Fusarium graminearum and A. terreus, indicating putative gene duplication events. In contrast to A. terreus, A. oryzae, A. niger and A. flavus, no a-class homologues have been found in the genomes of other Aspergilli such as A. nidulans, A. fumigatus and A. clavatus. A comparative analysis of the A. nidulans, A. fumigatus and A. oryzae genomes revealed that the latter has approximately 20% more genes than

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A. nidulans and A. fumigatus. These additional genes might be the result of putative horizontal gene transfer events from Sordariomycetous species to A. oryzae (Khaldi and Wolfe 2008). Similarly, horizontal gene transfers might be the reason for the distribution of a-carbonic anhydrases genes in the Aspergilli (Elleuche and Po¨ggeler 2009b). The primary structure of all the identified enzymes in this group clearly assigned them to the a-class and revealed a strict conservation of the zinc-coordinating domain, while other regions are not very well conserved (Fig. 11.3). However, in contrast to mammals, none of the identified fungal a-carbonic anhydrase exhibits changes in the three key amino acids, which are essential for the coordination of the zinc ion (Supuran 2008a). The functional role of a-carbonic anhydrases in fungi is so far unknown, but a model has been proposed in which CAS4 of S. macrospora, an isoform that might be secreted according to in silico analysis, is involved in bicarbonate production that will be used outside the cell or be imported through putative transport systems (Elleuche and Po¨ggeler 2010). Experimental investigations in the future will reveal insights on the role of a-class carbonic anhydrases in filamentous ascomycetes.

Fig. 11.3. Multiple sequence alignment and schematic drawing of members of the a-class carbonic anhydrases from filamentous ascomycetes. ClustalX alignment was generated with the protein sequences of the putative a-carbonic anhydrases from ascomycetous species (Thompson et al. 1997). Arrows indicate the position of the amino acids relevant for zinc coordination. Details for genera, species and accession numbers or scaffolds from genome projects are as follows (from the top): Fusarium graminearum_a (XP_391271.1),

B. b-Carbonic Anhydrases The budding yeast S. cerevisiae carbonic anhydrase Nce103 was the first b-class enzyme discovered in fungi in 1996. First it was predicted to be a component of a non-classical export pathway, but in 1999, sequence analysis assigned Nce103 to the carbonic anhydrases (Cleves et al. 1996; Go¨tz et al. 1999). Furthermore, Nce103 has been extensively analysed with regard to its cellular function. A deletion mutant exhibits a typical high CO2-requiring (HCR) phenotype under ambient air conditions, a phenotype well known from prokaryotes lacking carbonic anhydrase activity. This result led to the conclusion that the main physiological role of Nce103 is in the production of bicarbonate for metabolic carboxylation reactions catalysed by bicarbonate dependent enzymes under low CO2 conditions (Go¨tz et al. 1999; Aguilera et al. 2005).

Recent molecular phylogenetic analyses revealed putative b-carbonic anhydrase sequences at least in individual members of each subphylum. Most of the hemiascomycetes (Saccharomycotina) have only one b-carbonic anhydrase. A similar situation has been found in Schizosaccharomyces pombe, a member of the phylum Taphrinomycotina. The only exceptions are the yeasts C. albicans and P. stipitis, both encode a second hypothetical

F. graminearum_b (XP_384779.1), Podospora anserina (XP_001906308.1), Chaetomium globosum (XP_001227267.1), Magnaporthe grisea (XP_363766.1), Neurospora crassa (XP_960214), Sordaria macrospora (CAS4, FN178637), Aspergillus oryzae (XP_001827551.1), A. terreus_a (XP_001210252.1), A. terreus_b (XP001215283.1), Trichoderma virens (Trive1/scaffold_8), T. reesei (Trire2/scaffold_32), T. atroviride (Triat1/scaffold_17), Phaeosphaeria nodorum (XP_001790777.1), Paracoocidioides brasiliensis (ACA28690.1)


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isozyme (Elleuche and Po¨ggeler 2009b). In contrast to hemiascomycetous yeasts and S. pombe, a set of three fungal carbonic anhydrase encoding genes is present in most filamentous ascomycetes (Fig. 11.4A). Interestingly, the b-class encompasses two different subclasses, based on their primary structure. This classification was originally proposed for enzymes from prokaryotes and plants. The highly conserved key active site residues in the plant-type subclass were initially identified in pea Pisum sativum and were found to be variable in cab-type enzymes (Kimber and Pai 2000). The cab-type subclass is named after carbonic anhydrases b from the methane producing archaeon Methanobacterium thermoautotrophicum. It has been shown that the different

subtypes can be also distinguished by differences in inhibition patterns of known carbonic anhydrase inhibitors, indicating that the development of subclass specific inhibitory substances might be possible in the future (Supuran 2008b).

Fig. 11.4. Schematic diagram depicting the distribution of b-class carbonic anhydrases in ascomycetous subphyla. (A) Unrooted tree of b-carbonic anhydrases from ascomycetous subphyla. The phylogenetic tree was made with PHYLIP, based on the ClustalX alignment of the catalytic region shown in B. Plant-like b-CAS1, b-CAS2 and cablike b-CAS3 are encoded within the Pezizomycotina, while plant-like b-CA yeasts are from members of the subphylum Saccharomycotina. Spo Schizosaccharomyces pombe. (B) Consensus sequences of the zinc-coordinating domains from b-carbonic anhydrases in ascomycetes.

Alignment of the consensus sequences of plant-like CAS1, CAS2 and cab-like CAS3 homologues compared to the carbonic anhydrases from hemiascomycetous yeasts (Yeast) and S. pombe (Spo; subphylum Taphrinomycotina). For the generation of the consensus sequences, 17 zinc-coordinating regions from CAS1 homologues, 17 from CAS2 homologues, 29 from CAS3 homologues and 11 sequences from hemiascomycetous yeast-specific carbonic anhydrases were aligned using ClustalX (Thompson et al. 1997)

In prokaryotes, a third subclass of b-carbonic anhydrases, initially designated the e-class, was identified in the carboxysomal shell of the chemolithoautotrophic bacterium Halothiobacillus neapolitanus (Sawaya et al. 2006). Related members in fungal organisms have not been identified so far.

In filamentous ascomycetes CAS1- and CAS2homologues (for carbonic anhydrases of Sordaria macrospora) belong to the plant-type b-carbonic anhydrases and can be distinguished from CAS3-homologues, which are cab-like enzymes

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(Elleuche and Po¨ggeler 2009b). In hemiascomycetous yeasts (Saccharomycotina), plant-like b-class carbonic anhydrases are highly conserved and additional isozymes in C. albicans and P. stipitis have been identified as cab-type (Elleuche and Po¨ggeler 2009b). The active site of b-carbonic anhydrases contains a single conserved histidine and two conserved cysteine residues (Fig. 11.4B). Additionally, an aspartic acid and an arginine are also abundant in all known b-class carbonic anhydrase sequences known from bacterial, plant and fungal origin (Tripp et al. 2001; Elleuche and Po¨ggeler 2009b). It has been proposed that these two residues play roles in substrate binding, proton shuffling, product release or that they might act as a fourth ligand (Mitsuhashi et al. 2000; Cronk et al. 2001; Smith et al. 2002). The crystal structure of the S. cerevisiae carbonic anhydrase Nce103 recently revealed that it resembles the typical structure of plant-like b-carbonic anhydrases in which the zinc ion is coordinated by the three highly conserved catalytic residues (Cys57, His112, Cys115) and a water molecule to complete the tetrahedral coordination (Teng et al. 2009). Various oligomerization states have been described for the plant-type b-carbonic anhydrases of Escherichia coli (dimer), Haemophilus influenzae (tetramer), P. sativum (octamer) and others (Kimber and Pai 2000; Cronk et al. 2001, 2006; Schlicker et al. 2009; Teng et al. 2009).

Three components have been shown to be part of the monomer: a N-terminal arm and a C-terminal subdomain are flanking the zinc-binding a/b core. Activity assays proved that the N-terminal arm of Nce103 is indispensable for functionality. Furthermore, it has been shown that Nce103 assembles as a homodimer, which forms a substrate tunnel to the bottom of the active site with a bottleneck composed of amino acid residues Asp59, Phe97, Leu102 and Gly111 (Teng et al. 2009). Nevertheless, the flanking sequences of the zinc-coordinating domain from distantly related ascomycetes are very divergent or even non-homologous and therefore only the catalytic region indicated in the multiple sequence alignment of Fig. 11.4B has been used for the phylogenetic analysis given in Fig. 11.5. 1. Plant-Like b-Carbonic Anhydrases Almost all filamentous ascomycetes exhibit two closely related plant-type b-carbonic anhydrases

that are very similar at the primary sequence level, suggesting an ancient gene duplication event within the common ancestor of the filamentous ascomycetes. Gene duplication has been described as a major evolutionary force representing the main origin of functional novelty (Otto and Yong 2002). The comprehensive functional characterization of the two members of the plant-like b-carbonic anhydrase family in the filamentous ascomycetes S. macrospora, A. fumigatus and A. nidulans provides first insights into the diverse physiological roles of paralogous carbonic anhydrase pairs in fungi (Elleuche and Po¨ggeler 2009a; Han et al. 2010). Duplicated genes can have several likely fates. Divergence in function or sequence is more likely than the deletion of an open reading frame. An example of this scenario has been recently described. The functional characterization of members of the huge cutinase family in the fungal pathogen Magnaporthe oryzae revealed that cutinase paralogues have carried over differential physiological functions during the course of evolution, driven by the novel acquisition of diverged transcriptional regulation patterns (Skamnioti et al. 2008).

Similarly, plant-type fungal carbonic anhydrases may have acquired signal sequences bringing them to the novel subcellular localizations (Fig. 11.6). CAS1 and CAS2 homologues are localized to the cytosplam or the mitochondria, respectively, moreover they exhibit differential transcriptional regulation, resulting in different temporal and spatial expression levels (Elleuche and Po¨ggeler 2009a). Both processes stabilize the maintenance of paralogues after duplication in the genomes of filamentous ascomycetes. S. macrospora is a model organism for fungal fruiting body development with a genome that has been recently sequenced (Ku¨ck et al. 2009; Nowrousian et al. 2010). The role of S. macrospora CAS2 has been investigated in detail and it was proven that this specific carbonic anhydrase is essential for ascospore germination in a localization-dependent manner. Moreover, deletion of cas2 results in a pleiotropic phenotype with defects in vegetative and sexual growth and a drastically delay in the germination of ascospores. Interestingly, it was not possible to fully complement the Dcas2 deletion strain by expressing a CAS2-variant that was localized to the cytosol (Elleuche and Po¨ggeler 2009a). Although


Fig. 11.5. Evolutionry tree diagram (rectangular cladogram) of fungal a- and b-carbonic anhydrases from ascomycetes. The phylogenetic tree was made using programs from the program package PHYLIP, based on a ClustalX alignment of the catalytic region from carbonic anhydrases. Accession numbers and numbers of hypothetical proteins or scaffolds from genome projects are as follows: Ashbya gossypii (NP_983870.1), Aspergillus clavatus_1 (XP_001271988), A. clavatus_2 (XP_001273459.1), A. flavus (EED55452), A. fumigatus_CafA (XP_751704.1), A. fumigatus_CafB )XP_001481412.1), A. fumigatus_CafC

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(XP_751882.1), A. nidulans_CanA (XP_663215.1), A. nidulans_CanB (Q5BCC5), Aspergillus oryzae_1 (XP_001820193), A. terreus_1 (XP_001213134), A. terreus_3 (XP_001209372), Botryotinia fuckeliana_1 (XP_001555448.1), B. fuckeliana_3 (XP_001561102.1), Candida albicans_NCE103 (Q5AJ71), C. albicans_3 (XP_ 715817), C. glabrata_Nce103 (XP_446428.1), Chaetomium globosum_1 (XP_001226871.1), C. globosum_2 (XP_ 001227705), C. globosum_3 (XP_001225170.1), Coccidioides immitis_1 (XP_001247766.1), C. immitis_3 (XP_001241380), Debaryomyces hansenii (XP_456870),

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homologues cafA and cafB (carbonic anhydrase of A. fumigatus) was unable to grow under ambient air conditions, but the growth defect was fully rescued by high CO2 concentrations, indicating that both genes play redundant roles. However, only the heterologous expression of the cytoplasmic b-class carbonic anhydrase encoding gene cafB fully complemented the HCR-phenotype of a S. cerevisiae nce103 null allele mutant (Han et al. 2010). Fig. 11.6. Localization patterns of b-class carbonic anhydrases from S. macrospora. Fluorescent microscopic analyses illustrate the localization of b-class carbonic anhydrases CAS1, CAS2 and CAS3 in the filamentous ascomycete S. macrospora. CAS1 and CAS2 are localized to the cytoplasm, while CAS2 is translocated into the mitochondria. Strains were analysed after growth for 2 days on solid SWG medium supplemented with hygromycin. DIC Differential interference contrast, GFP green fluorescent protein. Scale 20 mm

In contrast to A. fumigatus, the closely related A. nidulans has only the cas1 and cas2 homologues can1 and can2 (carbonic anhydrase of A. nidulans) with no additional cab-type carbonic anhydrase. According to bioinformatical predictions, none of these proteins seem to be translocated into the mitochondria, but both genes encode functional active isozymes. Only the generation of DcanB strain resulted in a typical HCR phenotype, indicating that CanB plays the predominant role in A. nidulans (Han et al. 2010).

a neofunctionalization process might have occurred for CAS1 or CAS2 in this filamentous ascomycete, both isozymes share partially overlapping redundant functions under physiological conditions, since the Dcas2 phenotype can be partially restored by cas1 under ambient air conditions (Elleuche and Po¨ggeler 2009a). It has been shown in S. cerevisiae that the functional complementation of duplicated genes contribute to the robustness of genetic networks (Gu et al. 2003). In contrast to S. macrospora, A. fumigatus double deletion strain of the cas1 and cas2

A comprehensive in silico analysis revealed the presence of putative mitochondrial target sequences in CAS2 homologues from members of the orders Sordariales, Hypocrales, Pleosporales and in Magnaporthe grisea, but not from species of the order Eurotiales (Fig. 11.5). In this order, the CAS1 homologues are predicted to be translocated to the mitochondria. Because of their subcellular localization it might be likely that CAS1 homologues from members of the order Eurotiales and CAS2 homologues encoded within the genomes of the Sordariales, Hypocreales and Pleosporales are putative functional orthologues.

Fig. 11.5. (continued) Fusarium graminearum_2 (XP_384734.1), F. graminearum_3a (XP_383146.1), F. graminearum_3b (XP_390629), F. oxysporum_2 (supercont2.20), F. oxysporum_3a (FOXG_12252.2), F. oxysporum_3b (FOXG_13574.2), F. verticillioides_2 (supercont_3.16), F. verticillioides_3a (FVEG_10874.3), F. verticillioides_3b (FVEG_01549.3), Kluyveromyces lactis (XP_455263_1), Lodderomyces elongisporus (XP_001527257.1), Magnaporthe grisea_1 (XP_362166.2), M. grisea_2 (XP_366523), M. grisea_3 (XP_364389.1), Neosartorya fischeri_1 (XP_001266926), N. fischeri_2 (XP_001262181.1), N. fischeri_3 (XP_001267068.1), Neurospora crassa_1 (XP_960227), N. crassa_2 (XP_959676), N. crassa_3 (XP_961715), Paracoccidioides brasiliensis_1 (PADG_01697.1), P. brasiliensis_2 (supercont1.12), P. brasiliensis_3 (PADG_00315.1), Phaeosphaeria nodorum_1 (XP_001797035.1), P. nodorum_2 (XP_001 802070.1), P. nodorum_3 (XP_001805507.1), Pichia guilliermondii (EDK37806.2), Pichia stipitis

(XP_001386459.2), P. stipitis_3 (XP_001383682.1), Podospora anserina_1 (XP_001905915.1), P. anserina_2 (XP_001905568.1), P. anserina_3 (XP_001911575.1), Saccharomyces cerevisiae_Nce103 (NP_014362.1), Schizosaccharomyces pombe (NP_596512.1), Sclerotiana sclerotiorum_1 (XP_001598 060.1), Sordaria macrospora_ CAS1 (FM878 639), S. macrospora_CAS2 (FM878640), S. macrospora_CAS3 (FM878 641), Trichoderma atroviride_2 (Triat1/scaffold_4), T. atroviride_3a (Triat1/ scaffold_13), T. atroviride_3b (Triat1/scaffold_5), T. atroviride_3c (Triat1/scaffold_18), T. atroviride_3d (Triat1/ scaffold_14), T. reesei_2 (Trire2/scaffold_25), T. reesei_3a (Trire2/scaffold_30), T. reesei_3b (Trire2/scaffold_5), T. reesei_3c (Trire2/scaffold_3), T. virens_2 (Trive1/scaffold_4), T. virens_3a (Trive1/scaffold_15), T. virens_3b (Trive1/scaffold_12), T. virens_3c (Trive1/scaffold_13), Vanderwaltozyma polyspora (XP_001644582.1), Yarrowia lipolytica (XP_505708.1). Accession numbers of a-class carbonic anhydrases are indicated in Fig. 11.3


Within the members of the genera Trichoderma and Fusarium (order Hypocreales), no cas1homologues were identified in BLAST analyses. This result indicates that a former homologue might have got lost during evolution or that the gene duplication event that generated cas1 and cas2 isozymes only took place in limited groups of the filamentous ascomycetes. It has been hypothesized that bacterial carbonic anhydrases appeared when novel enzymatic roles were needed under changing physiological conditions (Smith and Ferry 2000). Similar results have been found for the carbonic anhydrases in plants, algae and animals, where this family of proteins is also targeted to different cellular compartments and involved in a variety of cellular processes (Hewett-Emmett and Tashian 1996; Moroney et al. 2001; Supuran 2008a). In this context, it was proposed that the neofunctionalization of fungal carbonic anhydrases in combination with the subcellular specialization are thought to be quite recent events, because of the high degree of sequence similarity in the gene sequences of fungal CAS1 and CAS2 homologues (Elleuche and Po¨ggeler 2010). Moreover, Coccidioides immitis and Paracoccidioides brasiliensis, both species of the order Onygenales encode putative mitochondrial carbonic anhydrases that are no homologues. The C. immitis cas1 homologue encodes a protein with a predicted 33-amino-acid mitochondrial target sequence, while a 66-amino-acid signal peptide is predicted at the N-terminus of the CAS2-homologue from P. brasiliensis (Elleuche and Po¨ggeler 2009b). 2. Cab-Like b-Carbonic Anhydrases Beside the well conserved gene subfamily encoding fungal plant-like b-carbonic anhydrases, cabtype isozymes were also identified predominately within fungal genomes of filamentous ascomycetes. A single cab-type carbonic anhydrase has been identified by BLAST searches in members of the orders Sordariales, Helotiales, Onygenales, Pleosporales and in several members of the Aspergilli (Eurotiales), while multiple open reading frames encoding cab-type b-carbonic anhydrases were identified within members of the order Hypocreales, suggesting that cab-like carbonic

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anhydrases have had a unique history of duplication events in different orders during the course of evolution (Fig. 11.5). It seems reasonable to assume that one of the multiple CAS3 homologues might have taken over the physiological function of the plant-like CAS1 homologue, which may have gone lost during the evolution of the order Hypocreales (Section III.1). Furthermore, it might be likely that some of these duplicated genes are encoding non-functional proteins, although all cab-type carbonic anhydrases identified revealed no changes in the three essential key amino acid residues of the zinc-coordinating domain, but this has not been proven yet. The first indication of an active cab-type b-carbonic anhydrase has been given experimentally by the constitutive expression of cab-type cas3 in a Dcas1/2 double deletion background resulting in the partial complementation of the sterile phenotype (Elleuche and Po¨ggeler 2009a). According to bioinformatic approaches and experimentally studies using S. macrospora CAS3 (Fig. 11.6), most of the fungal cab-type b-carbonic anhydrases seem to be exclusively cytoplasmic enzymes, with the A. fumigatus CafD as the sole exception so far (Elleuche and Po¨ggeler 2009a, b; Han et al. 2010). A. fumigatus belonging to the order Eurotiales has been recently identified to encode two cab-type carbonic anhydrases (cafC and cafD). The deletion of cafD did not result in any obvious bicarbonate-dependent phenotype, while cafC was shown to play a minor role in conidiation. Furthermore, the heterologous expression of cafD in the S. cerevisiae nce103 mutant does not complement the HCR phenotype, indicating that cafD might probably not encode a functional carbonic anhydrase, although it was not possible to generate a DcafA/D double deletion mutant. However, cafD is located 288-bp adjacent to the A. fumigatus cas2 homologue cafB and in silico analyses also predicted a 30-amino-acid residues signal peptide at the N-terminus, which has been proposed to be important for mitochondrial translocation (Han et al. 2010). In addition, cafD may encode a novel type of cab-like carbonic anhydrase in filamentous fungi and is probably not a member of the CAS3 group. Furthermore, carbonic anhydrases CafC and CafD might not be the result of a duplication event of a common ancestor, since the sequence identity of both genes is less then 20% in a global alignment. Interestingly, the presence of CafD homologues seems to be restricted to genomes of several members of the order Eurotiales.

The distribution and evolutionary history of b-class carbonic anhydrases in ascomycetes is

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obviously complex, but the characterization of carbonic anhydrases from evolutionarily ancient fungi might shed light on the origin of these fungal enzymes.

IV. Carbonic Anhydrases in Basidiomycetes Carbonic anhydrases are widely distributed within the basidiomycetes. BLAST searches of basidiomycetous genomes revealed the existence of a- and b-carbonic anhydrases. Multiple genes encoding putative a-class carbonic anhydrases have only been identified in the genome of Laccaria bicolor, but these have not been investigated at the experimental level so far (Elleuche and Po¨ggeler 2010). Single b-class encoding carbonic anhydrase genes were identified in the genomes from members of the orders Agaricales (Coprinopsis cinerea, L. bicolor), Malasseziales (Malassezia globosa) and Ustilaginales (Ustilago maydis), while two closely related plant-type b-class carbonic anhydrases – Can1 and Can2 – have been extensively characterized in the ubiquitous human pathogen Cryptococcus neoformans (order Tremellales) indicating that an evolutionary gene duplication event might be restricted to the order Tremellales or even to the genus Cryptococcus within the basidiomycetes (Bahn et al. 2005, 2007). The availability of further genomic sequences will solve this question in the future. Can1 and Can2 of C. neoformans are 36% identical at the protein level and group together in phylogenetic analyses, but either underwent independent functional differentiation or Can1 lost its physiological functionality almost completely (Bahn et al. 2005; Schlicker et al. 2009). It has been suggested that a chromosomal translocation event and/or a segmental duplication event might be a possible reason for the duplication of a C. neoformans b-carbonic anhydrase ancestor, since the complete genome has not been duplicated during evolution (Loftus et al. 2005; Elleuche and Po¨ggeler 2009b). The C. neoformans CAN2 is essential for survival under ambient air conditions and mediates the pathogenic switch in the human host at 5% CO2, which includes the biosynthesis of a polysaccharide capsule (Bahn et al. 2005). This defect arises in the presence of CAN1, although this gene also encodes a catalytic active isozyme, indicating a nonfunctionalization of CAN1 at

least under the conditions tested. Interestingly, the activity of Can2 is dispensable under high CO2 conditions predominating in the human lung (Bahn et al. 2005). The a-carbonic anhydrase family in mammals containing 16 isozymes all originating from a common ancestor can serve as another example for non- and neofunctionalization. Three of them lost their function by substitutions of one or more of the functionally important key amino acids within the catalytic zinc-coordinating domain. These proteins are termed carbonic anhydrase related proteins (CARPs). The remaining 13 enzymes benefited from the great potential of acquiring novel physiological functions, which finally led to tissue or organ specific expression profiles and different subcellular localization patterns (Sly and Hu 1995; Supuran 2008a; Aspatwar et al. 2010). In contrast to the C. neoformans Can1, the three human CARPs completely lost their catalytic activities during evolution.

The crystal structure of the C. neoformans Can2 was solved and exhibited the conserved catalytic site composed of Cys68, His124 and Cys127, important for zinc-coordination. However, it carries a unique N-terminal extension, which forms two a-helices. The N-terminus is not homologous to the shorter N-terminal arm of S. cerevisiae Nce103 (Schlicker et al. 2009; Teng et al. 2009). Interestingly, this N-terminus of Can2 is capable of interacting with a surface groove at the catalytic site entrance of the homodimer. Due to the conservation of this N-terminus in other fungal homologues, it has been speculated that it may be involved in protein–protein interactions. Although the overall structure of Can2 resembles the structures of known planttype b-carbonic anhydrases, the C-terminus of Can2 is more like cab-type carbonic anhydrases from M. thermoautotrophicum and Mycobacterium tuberculosis (Strop et al. 2001; Covarrubias et al. 2006; Schlicker et al. 2009). However, Can2 also revealed a homodimeric crystal structure, similar to the plant-like carbonic anhydrase Nce103 from S. cerevisiae.

V. Conclusion The gene family of carbonic anhydrases encodes five evolutionary unrelated classes (a, b, g, d, z) of enzymes. To date only a-, b- and g-class enzymes have been identified in genomes of fungi and fungal-like organisms, but only members of the b-class are characterized in detail. The


distribution of carbonic anhydrase genes is highly diverse in the genomes of fungal-like organisms and in the ‘Basal Fungi’. It can be assumed that a highly dynamic evolution has taken place in the ancestors of the fungal-like P. infestans and in the chytrid A. macrogynus, because in both multiple isozymes were identified that have still an almost identical sequence composition. The evolution of plant-type b-carbonic anhydrases in ascomycetes is dominated by a putative gene duplication event after the separation of the filamentous ascomycetes from the yeasts and the subcellular translocation of a single isoform into the mitochondria. The specific mitochondrial localization of at least one carbonic anhydrase in different ascomycetous orders might indicate that the acquisition of novel functions went along with the mitochondrial translocation of a specific isoform. In the basidiomycetes, it is hard to propose a model for the evolution of carbonic anhydrases, since only a few enzymes have been identified in this phylum. Acknowledgements I am grateful to Prof. Stefanie Po¨ggeler for her constant support and for critically reading this manuscript.

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Evolutionary Mechanisms and Trends

12

Evolution of Mating-Type Loci and Mating-Type Chromosomes in Model Species of Filamentous Ascomycetes

CARRIE A. WHITTLE1, HANNA JOHANNESSON1

CONTENTS I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. The Mating-Type Locus in Filamentous Ascomycetes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Comparison of Mating-Type Loci Features of Filamentous Ascomycetes and Other Fungi. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Mating-Type Locus and Mating-Type Chromosome Evolution in the Model System Neurospora. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Mating-Type Locus Evolution in Neurospora . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Mating-Type Chromosome Evolution in Neurospora tetrasperma . . . . . . . . . . . . . . . . . . . . 1. Parallels Between N. tetrasperma Mating-Type Chromosomes and Sex Chromosomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Independent N. tetrasperma Phylogenetic Lineages. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Benefits of N. tetrasperma as a Model System for Early-Stage Sex Chromosome Evolution. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4. Recombination Suppression and Genomic Degeneration in N. tetrasperma mat Chromosomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . III. Key Findings Regarding the Evolution of Mating-Type Genomic Regions in Other Fungal Model Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Cryptococcus neoformans . . . . . . . . . . . . . . . . . . . . . . B. Microbotryum violaceum . . . . . . . . . . . . . . . . . . . . . . C. Evidence of Interspecific Gene Transfer from Ophiostoma novo-ulmi . . . . . . . . . . . . . . . . . . D. Mating-Type Loci in a Basal Fungus. . . . . . . . . . IV. Future Directions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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I. Introduction Sexual reproduction in fungi is typically regulated by small chromosomal regions containing one or more mating-type loci. Among the filamentous ascomycetes (i.e., Pezizomycotina) which 1

Department of Evolutionary Biology, Uppsala University, Uppsala, Sweden; e-mail: [email protected]

comprise the largest subphylum in the Ascomycota (Alexopoulos et al. 1996; James et al. 2006; Spatafora et al. 2006; Sugiyama et al. 2006), the genome contains a single mating-type locus composed of two highly dissimilar alleles (referred to as idiomorphs; Shiu and Glass 2000; Casselton 2008; see Chapter 5 in this volume). The two mating-type idiomorphs, each consisting of one or more genes, are associated with various stages of reproduction, including sex organ development, gamete development and attraction, and post-fertilization differentiation (Glass et al. 1990a; Fraser and Heitman 2004). The matingtype locus also regulates sexual compatibility. For example, to produce sexual spores heterothallic (self-incompatible) species require a partner containing nuclei harboring the opposite idiomorph at the mating-type locus (Glass et al. 1990a, b; Shiu and Glass 2000) while homothallic (self-compatible) species reproduce autonomously and often carry genes of both idiomorphs within each haploid nucleus (Coppin et al. 1997). Pseudohomothallic species are also highly selffertile, but are distinguished from homothallics by the presence of two haploid nuclei of opposite mating type within ascospores as well as vegetative cells. Given the key role of the mating-type locus/loci in fungal mating and sexual reproduction and its implications to evolutionary biology and population genetics, much attention has recently been devoted to characterization of this genomic region and the identification of factors altering its evolution in fungi, particularly among the filamentous ascomycetes (Casselton 2008). In this chapter we describe the genomic traits and evolutionary features of the mating-type loci and the mating-type chromosomes in model systems of filamentous ascomycetes, and we highlight the comparable findings from other fungal phyla and other kingdoms. Evolution of Fungi and Fungal-Like Organisms, The Mycota XIV S. Po¨ggeler and J. Wo¨stemeyer (Eds.) © Springer-Verlag Berlin Heidelberg 2011

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Carrie A. Whittle and Hanna Johannesson

A. The Mating-Type Locus in Filamentous Ascomycetes Most available research on the mating-type locus to date has been derived from heterothallic filamentous ascomycetes (Heitman et al. 2007; Idnurm et al. 2008). In particular, the matingtype locus has been well characterized in a wide range of species, including Neurospora crassa (Glass et al. 1990a; Staben et al. 1990; Ferreira et al. 1996), Cochliobolus heterostrophus (Turgeon et al. 1993), Coccidioides immitis, Coccidioides posadasii (Fraser et al. 2007; Mandel et al. 2007), Pyrenopeziza brassicae (Singh and Ashby 1998), Podospora anserina (Debuchy et al. 1992, 1993), Magnaporthe grisea (Kang et al. 1994), Ascochyta rabiei (Barve et al. 2003) and Histoplasma capsulatum (Bubnick and Smulian 2007; Fraser et al. 2007). The consensus from these analyses is that mating types in filamentous ascomycetes are conferred by highly dissimilar non-recombining idiomorphs at the mating-type locus, each of which range in size between 1 and 6 kb (Coppin et al. 1997; Dyer 2008) and normally contain one to three genes (Dyer 2008). DNA sequences of mating-type loci idiomorphs have also been identified in genomes of homothallic taxa, including Sordaria macrospora, Aspergillus nidulans and species of Cochliobolus and Neurospora (Beatty et al. 1994; Coppin et al. 1997; Po¨ggeler et al. 1997; Yun et al. 1999; Galagan et al. 2005). Data from the N-eurospora africana mat A-1 gene suggests that this gene is functional, that is, it activates mating and vegetative incompatibility in N. crassa (Glass and Smith 1994). Mating-type genes have also been detected in asexual filamentous ascomycetes such as Bipolaris sacchari (Sharon et al. 1996) and in the supposedly asexual taxon Aspergillus fumigatus (Paoletti et al. 2005). In totality, the data suggests that the matingtype genes are often maintained in the genome even after the divergence of mating systems but whether they are evolutionarily constrained or a vestige of an ancestral heterothallic stage still remains to be determined. The mating-type genes in filamentous ascomycetes have been shown to encode transcription factors, including high mobility group (HMG) domain proteins and a-box domain proteins (Coppin et al. 1997; Fraser and Heitman 2004; Casselton 2008). The simpler heterothallic systems (e.g., Cochliobolus heterostrophus) harbor

one gene in each idiomorph, one that encodes an HMG domain protein and the other that encodes an a-box protein (Yun and Turgeon 1999). More complex systems are inherent to specific taxa, such as N. crassa, wherein three genes are encoded by the mat A idiomorph, namely mat A-1, mat A-2 and mat A-3 (which encode an a-domain protein, an acidic a-helix and a HMG protein, respectively) while one gene, encoding a HMG protein, is found in its mat a counterpart (Coppin et al. 1997; Shiu and Glass 2000). These transcription factors direct sex-specific gene regulatory pathways, which determine cell fate, identity and underlie reproductive processes (Coppin et al. 1997; Casselton 2002). For example, the mat genes in N. crassa have been associated with the expression of genes that regulate pheromones and their receptors, which is required for attraction among mating types, fertilization and sex-specific development (Casselton 2002; Bobrowicz et al. 2005; Kim and Borkovich 2006; see Chapter 5 in this volume). The mat A idiomorph in particular has been associated with expression of the pheromone gene ccg4 and the receptor gene pre-1. Both genes are highly expressed in mat A reproductive cells during mating, and have been shown to be closely involved in sexual attraction and female functionality and fertility (Kim and Borkovich 2004). Expression of the pheromone gene mfa-1 and the pheromone gene pre-2 is primarily associated with mat a strains (Kim and Borkovich 2006; Karlsson et al. 2008). The evolution of the structure of the mating-type locus across fungal groups may be revealed from comparative examination among filamentous and non-filamentous ascomycetes and basidiomycetes. B. Comparison of Mating-Type Loci Features of Filamentous Ascomycetes and Other Fungi Data to date suggests that certain features of the mating-type loci are highly conserved among filamentous ascomycetes, yeast and the basidiomycetes. For example, in all of these taxonomic groups the mating-type loci are associated with pheromone-pathways (Kronstad and Staben 1997; Casselton 2008) and encode transcription factors (e.g., HMG, a-box, or homeodomain proteins, such as HD1 in yeast; Banham et al. 1995; Coppin et al. 1997; Hull and Johnson 1999; Tsong et al. 2007). Other features of the mating-type

Evolution of Mating-Type Loci and Mating-Type Chromosomes

loci differ markedly among the taxonomic groups. For instance, the mating-type loci in certain basidiomycetes contain pheromone and pheromone receptor genes (Wendland et al. 1995; Kothe 1996; O’Shea et al. 1998) while pheromone genes are indirectly activated through the mating-type genes in other fungal taxa including the filamentous ascomycetes, that is, the mating-type genes encode transcription factors (Kim and Borkovich 2004; Casselton 2008). In addition, the matingtype loci in some yeast and basidiomycete species contain genes encoding homeodomain proteins (e.g., Candida albicans, Saccharomyces cerevisiae) and certain yeast species can undergo mating-type switching wherein silent copies of MAT sequences are activated, neither of which is typically observed among the filamentous ascomycetes (Haber 1998; Scott et al. 2004; Casselton 2008). Moreover, the Basidiomycota contain many species with tetrapolar mating systems, which are often multi-allelic, and other species with bipolar mating systems with only two alleles that can span more than 100 kb and contain more than 20 genes. Tetrapolar mating systems are common in the Basidiomycota, which give rise to a vast array of possible mating types; these traits are not normally characteristic to filamentous ascomycetes (Kothe 1996; Kronstad and Staben1997; Casselton and Olenicky 1998; Casselton 2002; Fraser et al. 2004). In combination, these findings in various fungal taxa reveal a complex history, potentially including common ancestry of mating-type loci among lineages and/or independent divergence events (Casselton 2008) and highlight the novel features inherent to the matingtype loci of filamentous ascomycetes.

II. Mating-Type Locus and Mating-Type Chromosome Evolution in the Model System Neurospora A. Mating-Type Locus Evolution in Neurospora Neurospora is emerging as a model for studies on the evolution of the mating-type locus in filamentous ascomycetes. The currently available data from N. crassa indicates that the mat a idiomorph contains the open reading frame mat a-1, which is the main regulator of sexual reproduction

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in mat a strains and is required for mating identity, post-fertilization functions and for vegetative incompatibility (Glass et al. 1990a, b; Staben et al. 1990; Po¨ggeler and Ku¨ck 2000). The equivalent sexual regulator in the mat A idiomorph is mat A-1 (Glass et al. 1990a, b). The additional two genes encoded at the mat A idiomorph, mat A-2 and mat A-3, enhance the level of reproductive output but are not essential for reproduction (Ferreira et al. 1996, 1998). Although the structure of the mat locus is less well studied in homothallic species of Neurospora, hybridizations conducted using the genes in N. crassa as probes classified the homothallic taxa into three possible groups: those containing both mat A and mat a gene regions, those containing mat A sequences and lacking the mat a sequence, and those wherein mat a-1, mat A-1 and mat A-2 are present but the A-3 genomic region is absent (Beatty et al. 1994). Although the evolutionary dynamics leading to this complex variability in the mat locus among Neurospora lineages remains to be ascertained, recent comparative analysis has revealed some of the factors associated with their evolution. Comparative analysis of mat sequences among a wide range of Neurospora species (including N. crassa, N. tetrasperma, N. africana, N. intermedia, N. discreta), representing an array of mating systems and mat gene compositions, has revealed that mat genes evolve at a higher rate than non-reproductive genes (such as actin, EF1-alpha, glyceraldehyde 3-phosphate dehydrogenase; Po¨ggeler 1999; Wik et al. 2008; see Chapter 8 in this volume). This finding is consistent with the higher rate of protein evolution observed for sex-linked than autosomal genes observed in many animals, such as Drosophila, birds, mice, rats and humans (Charlesworth et al. 1987; Tucker and Lundrigan 1993; Swanson and Vacquier 2002; Mank et al. 2007). The underlying cause of the rapid evolution in Neurospora varies with the type of mating system. For example, likelihood analysis of codon sites among taxa indicates that positive selection drives the high rates of evolution in mat a-1 and mat A-2 sequences among heterothallic species (e.g., see mat A-2 in Fig. 12.1; Wik et al. 2008). Such rapid divergence of genes via positive selection can have important evolutionary consequences; for instance, this could give rise to barriers to fertilization and/or alter hybrid vigor, and thereby lead to speciation

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A

Heterothallic taxa, classes of ω:

Prob. 0

B

0.2

1.0

7.9

9

100

155

179

*

*

*

*

HPG-domain

304

334

357

*

*

*

1

Homothallic taxa, classes of ω:

0.0

1.0

Prob. 0

HPG-domain

1

Fig. 12.1. Posterior probabilities that codons of the mat A-2 gene belong to different site classes (o ¼ dN/dS, o < 0 indicates negative selection, o ¼ 1 indicates neutrality, o>1 indicates positive selection). Data is shown for heterothallic (A) and for homothallic (B) filamentous ascomycetes (based on analyses of Neurospora and

Gelasinospora species). The bar height indicates the probability. Asterisks indicate probability >0.95 that the numbered site belongs to class with o >1, consistent with positive selection. Arrows indicate intron splicing positions. Data is based on Wik et al. (2008)

Table 12.1. Mating-type genes detected among various species of homothallic filamentous ascomycetes. Genes having loss of function mutations (frameshift and stop mutations) are in italics. The Fungal Genomic Stock Center (FGSC) identification number for each strain is shown. Data taken from Wik et al. (2008) Taxon Taxa containing all MAT genes Gelasinospora calospora (Mouton) Moreau and Moreau Neurospora sublineolata (Furuya and Udagawa) von Arx Gelasinospora cerealis Dowding Neurospora pannonica Krug and Khan Taxon lacking MAT A-3 Neurospora terricola Gochenaur and Backus Taxa lacking MAT a-1 Neurospora africana Huang and Backus Neurospora galapagosensis Mahoney and Backus Neurospora dodgei Nelson and Novak Neurospora lineolata Frederick and Uecker

FGSC number

ORF detecteda

958 5508 959 7221

a1, A1, A2, A3 a1, A1, A2, A3 a1, A1, A2, A3 a1, A1, A2, A3

1889

a1, A1, A2

1740 1739 1692 1910

A1, A2b,A3 A1, A2, A3 A1, A2, A3 A1, A2, A3

a Gene regions were amplified by PCR using isolated cDNA and primers designed from published mating-type gene sequences or Neurospora crassa. b Failed to amplify from cDNA.

events (Swanson and Vacquier 2002). In contrast to heterothallic species, the high rate of sequence evolution in homothallic species at mat a-1, mat A-2 and mat A-3 is attributable to relaxed selective constraint, as demonstrated by a relatively high proportion of neutrally evolving codons (Fig. 12.1; Wik et al. 2008) and by enhanced levels of stop and frameshift mutations in these genes (Table 12.1; Wik et al. 2008). Except for mat A-2 in N. africana we were able to PCR amplify cDNA from all mat genes present in the genomes of the investigated homothallic taxa. This result indicates the mat genes are actively expressed in homothallic taxa of Neurospora; although this is likely to have a deleterious effect on fitness (i.e., particularly for the genes having loss of function mutations; Charlesworth 1996; Bachtrog 2006). Whether the genes that

remain functional in each homothallic species (e. g., mat A-1) can solely support reproductive development in a particular taxon needs further investigation. Recent data from Neurospora suggests the mat genes have novel evolutionary patterns as compared to other regions of the genome. In particular, a species tree generated using concatenated sequence data from four non-coding DNA regions for 11 outcrossing Neurospora taxa (N. crassa A, B, C, which are defined as distinct phylogenetic subgroups of N. crassa; Dettman et al. 2003), N, discreta. N. hispaniola N. intermedia (phylogenetic subgroups A, B; Dettman et al. 2003), N. metzenbergii, N. perkinsii, N. sitophila and the pseudohomothallic N. tetrasperma) (Dettman et al. 2003) differs markedly from the gene tree


generated using genes encoded by the mat A and mat a idiomorphs (Strandberg et al. 2010). Although the causes of such variation remain to be ascertained, the finding suggests that novel processes/forces such as selective introgression drive the evolution of mat idiomorphs. Another plausible explanation is the existence of ancestral polymorphism for genes encoded by each of the mat idiomorphs (Tajima 1983; Takahata and Nei 1985; Pamilo and Nei 1988); evidence suggests that such polymorphism can occur for individual nonrecombining sex chromosomes due to recurrent mutations and haploid selection (e.g., Repping et al. 2003). It is notable that gene trees derived from the pheromone receptor genes (pre-1, pre-2) also deviate from the Neurospora species tree, however not in a manner parallel to the mat genes (Strandberg et al. 2010), suggesting this type of phenomenon could be inherent to genes that play a key role in mating and reproduction. Overall, it is evident that genes involved in reproduction have novel divergence patterns within Neurospora, and thus, could play a major role in reproductive isolation and speciation events within this taxonomic group (see Chapter 5 in this volume). B. Mating-Type Chromosome Evolution in Neurospora tetrasperma The model species Neurospora tetrasperma is pseudohomothallic. It exhibits several distinct reproductive features including the presence of a predominant heterokaryotic state, a novel meiotic program that leads to progeny containing two haploid nuclei of opposite mating type (mat A, mat a) and a very high level of self fertilization (Dodge 1927; Raju and Perkins 1994; Jacobson 1995; Powell et al. 2001). The N. tetrasperma mating-type (mat) chromosomes are highly novel among fungi as they have very limited recombination; data shows that this feature has been recently acquired in this lineage (approximately 4 to 6 MYA; Menkis et al. 2008) and is not inherent to other fungi, including Neurospora sister taxa, wherein the area of restricted recombination is normally limited to small genomic regions containing the mat locus (Merino 1996; Lee et al. 1999). The suppression of recombination in the region surrounding the mat locus in N. tetrasperma suggests parallels to the limited recombination widely observed in the dimorphic and

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recombinationally suppressed sex chromosomes of animals and some plants (Charlesworth and Charlesworth 2000; Menkis et al. 2008).

1. Parallels Between N. tetrasperma Mating-Type Chromosomes and Sex Chromosomes Marked parallels have been reported between the traits observed in animal/plant dimorphic sex chromosomes and the mat chromosomes in N. tetrasperma. Specifically, evidence indicates that a region of suppressed recombination in N. tetrasperma chromosomes spans an approximately 7-Mbp region containing the mat locus. This region represents more than 75% of the chromosome (Fig. 12.2; Menkis et al. 2008), consistent with the large region of suppressed recombination observed around the reproductive SRY gene in human Y sex chromosomes, which represents 95% of the chromosome (Rappold 1993; Fraser and Heitman 2004, 2005). In addition, the region of suppressed recombination in the mat chromosomes contain segments with different levels of sequence divergence (i.e., evolutionary strata), suggesting that there has been expansion of the recombinationally suppressed region over time (Fig. 12.2). Since evolutionary strata were first described for the human X/Y chromosomes, they have also been detected in the sex chromosomes from a wide range of species including mice, chicken and dioecious plants as well as for the mating-type chromosomes from the basidiomycete Microbotryum violaceum (Lahn and Page 1999; Lahn 2001; Fraser et al. 2004; Handley et al. 2004; Nicolas et al. 2004; Sandstedt and Tucker 2004; Menkis et al. 2008; Volintseva and Filatov 2009). Evolutionary strata are genomic regions with suppressed recombination within sex chromosomes which have arisen successively over time, and wherein younger regions normally have lower synonymous substitution rates (among gene alleles located on opposing sex chromosomes) than older regions (Fraser and Heitman 2005).

It has also been reported that the region of suppressed recombination in the N. tetrasperma mat chromosomes is flanked by regions with obligate crossovers, allowing a proper alignment of the mating-type chromosomes (Fig. 12.2; Gallegos et al. 2000; Jacobson 2005; Menkis et al. 2008), a phenomenon also inherent to other eukaryotic


recombining right flank 0.5Mbp (5.6%)

6.6Mbp (75%) old stratum

0.3Mbp (3.4%) recent stratum

recombining left flank 1.4Mbp (16%)

282 mat chromosome

2

Gene

dS mat A vs mat a

ro-10

0

nit-2

0

krev-1 sod-1 mus-42 rid-1 leu-4 cys-5 se r-3 tef-1 un-3 mat aut1 upr-1

0 0 0.0013 0

erg-8 arg-1

0.060 0.047

eth-1 lys-4

0.029

cys-9

0.082

ad-9

0.070

al-1

0.073

lys-3

0.063

os-1

0

prd-4

0

arg-13

0

phr

0

0.028 0 0.04 0.029 0.04 0.055 0.056

0.062

1

Fig. 12.2. Schematic representation of the mating-type chromosomes from Neurospora tetrasperma (strain P581) including the recombining flanking regions, the nonrecombining regions (stratum 1, stratum 2) and specific genes (listed by their relative gene order) from each genomic region. The genetic distance (dS) between single mating-type strains of the heterokaryotic N. tetrasperma (mat A vs mat a) are provided for each gene based on findings by Menkis et al. (2008)

systems including human X/Y chromosomes (Rappold 1993). Another trait common to animal sex chromosomes and N. tetrasperma mat chromosomes is that the recombination block is associated with chromosome rearrangements and genetic modifiers of recombination rates (Brooks 1988; Fraser and Heitman 2004, 2005; Charlesworth et al. 2005; Jacobson 2005). Moreover, homologous gene regions located in the recombinationally suppressed segment of the mat A and mat a chromosomes in N. tetrasperma demonstrate substantial sequence divergence (Fig. 12.2), which is consistent with the high rates of divergence observed in the X/Y chromosomes in other organisms such as humans, plants and Drosophila miranda (Rice 1987; Charlesworth 1996; Merino et al. 1996; Bachtrog 2003a; Charlesworth et al. 2005; Jacobson 2005; Marais et al. 2008; Menkis et al. 2008). It is notable that the N. tetrasperma mat chromosomes have not yet revealed evidence of massive gene losses as occurs on the permanently heterozygous sex chromosomes in animals (e.g., the Y or W chromosome), which likely results from selective sweeps and/or the sheltering or recessive mutations (Nei 1970; Charlesworth 1996; Charlesworth and Charlesworth 2000; Carvalho 2002; Skaletsky et al. 2003; Charlesworth et al. 2005). This apparent lack of gene loss may be because the degeneration is in its earliest stages and/or that haploid selection prevents degeneration in this predominantly heterokaryotic taxon. Altogether the data suggests that the mat chromosomes in N. tetrasperma share parallels with the sex chromosomes in animals and dioecious plants, in both the regions of suppressed recombination as well as in the recombining flanking regions (Menkis et al. 2008). 2. Independent N. tetrasperma Phylogenetic Lineages Recent data has suggested that N. tetrasperma has a complex phylogenetic history. In particular, comparative analysis of a genetically and geographically diverse selection of strains using multi-locus data (i.e., the non-repeat regions of the microsatellite loci DMG, TMI, TML, QMA; Dettman and Taylor 2004) and biological species recognition data (i.e., reproductive output and viability of laboratory crosses) has revealed that N. tetrasperma is a monophyletic group of


Neurospora and comprises at least nine distinct phylogenetic species, contained within one traditional, broadly defined, biological species (based solely on spore pigmentation criteria; Menkis et al. 2009). This pattern of genetic isolation preceding reproductive isolation has also been reported for heterothallic taxa such as N. discreta, which is comprised of eight distinct phylogenetic lineages within a single broadly defined biological species (Dettman 2006).

However, when integrating data on spore viability and self-fertility into the analyses, we scored a significantly higher reproductive success in crosses of strains from the same phylogenetic species than of different phylogenetic species, and when using a more narrow biological species recognition based on these criteria we found seven biological species of N. tetrasperma. Taken together, we conclude that N. tetrasperma is a species complex consisting of at least nine phylogenetic, reproductively isolated, species, which will be referred to as N. tetrasperma phylogenetic lineages 1–9 until the formal nomenclature is settled. Data from divergence of the mat chromosome between single mating-type component strains of wild heterokaryons representing the nine distinct N. tetrasperma phylogenetic species indicates a marked variation in the size of the recombination block between lineages. The region of limited recombination in the mat chromosomes may have grown, or shrunk, independently in each lineage (Menkis et al. 2009; Menkis et al. 2010), further supporting the conclusion that there has been autonomous evolution among these lineages. 3. Benefits of N. tetrasperma as a Model System for Early-Stage Sex Chromosome Evolution In order to better understand the early stages of sex chromosomes divergence, including the evolution of sex chromosomes from autosomes and the divergence of sex chromosomes from each other, the examination of relatively primitive (i.e., less evolved) systems is needed (Charlesworth and Charlesworth 2000). In addition to the recently emerging system from N. tetrasperma, one of the

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few other systems for such research is the neo-Y and neo-X chromosomes in Drosophila miranda. The neo-X/Y chromosome represents an autosomal region that is fused to the ancient X/Y chromosomes, and is believed to have arisen approximately 1.2 million years ago (Bachtrog 2005). This system has proven valuable in revealing traits associated with suppressed recombination along the sex chromosomes; for example, molecular evolutionary analysis has shown that neo-Y chromosome has reduced gene expression levels (found for >80% of genes), enhanced levels of frame-shift and stop mutations, increased transposition frequency, faster protein evolution and enhanced levels of non-preferred codons as compared to the neo-X chromosome (Bachtrog and Charlesworth 2002; Bachtrog 2003a, b, c, 2005, 2006; Bartolome´ and Charlesworth 2006).

Another model system is the dioecious plant Silene latifolia, wherein it has been demonstrated that the relatively young Y chromosome has lowered gene expression, fast protein evolution and enhanced transposable element frequency than its X-chromosome counterpart (Marais et al. 2008). Given the limited availability of model systems for the study of the early stages of evolution among sex chromosomes, the N. tetrasperma mat chromosomes offer valuable new opportunities to expand our understanding of this process. The mat chromosomes in N. tetrasperma are a particularly valuable system for the study of sex chromosome evolution due to several specific and novel traits. For example, in contrast to neo-X/Y chromosomes in Drosophila, the entire region of recombination suppression has been recently acquired within the N. tetrasperma mat chromosomes (Menkis et al. 2008). For the neo-X/Y chromosomes, the small autosomal region has been inserted into highly evolved ancient sex chromosomes (Charlesworth and Charlesworth 2000), and thus the evolutionary forces in the neo-X/Y chromosomes may not be reflective of those inherent to the earliest stages of suppressed recombination in proto-sex chromosomes, that is, wherein the regions of suppressed recombination in the sex chromosomes are all relatively young. In addition, the fact that N. tetrasperma is comprised of a wide range of closely related phylogenetic species allows one to identify and examine evolutionary patterns among the various mat chromosome regions among lineages (e.g., variation in the size of the recombinationally suppressed region, gene conversion events; Menkis et al. 2009; Menkis et al. 2010),

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a trait not inherent to other model systems. Another valuable feature of the N. tetrasperma mat chromosomes is that gene evolution on the limited-recombining region of mating chromosomes can be readily compared to closely related species such as N. crassa, which demonstrates normal recombination within the mat chromosomes. Furthermore, most genes outside of the region of suppressed recombination in N. tetrasperma are identical or nearly identical at each allele (due to the high selfing rates) and most genes are present on both mating-type chromosomes, that is, there has not been massive gene loss in one chromosome (Beatty et al. 1994; Menkis et al. 2008). This is in contrast to the highly evolved sex chromosomes in most animals and in some plants that retain few homologous genes and wherein the genes located on the permanently heterozygous sex chromosome have almost completely degenerated, such that there are few remnants of the historical events that underlie sex chromosome evolution (Charlesworth and Charlesworth 2000; Ellegren 2000; Charlesworth et al. 2005). Moreover, given that N. tetrasperma is a hermaphrodite, and its mating type chromosomes are not associated with morphological sex-determination per se (i.e., the male or female organs may contain either of the two mating type chromosomes; Coppin et al. 1997), sex-specific mutation and selection biases can be excluded as factors affecting gene/chromosome evolution (Jacobson 1995; Li et al. 2002; Ellegren and Parsch 2007; Hedrick 2007; Menkis et al. 2008). Altogether, these combined traits suggest that N. tetrasperma is a particularly valuable research system for the identification and characterization of molecular changes in the early stages of sex chromosome evolution.

4. Recombination Suppression and Genomic Degeneration in N. tetrasperma mat Chromosomes Recently generated large-scale genomic data from the N. tetrasperma mat chromosomes has provided new insights into the genomic events associated with the evolution of regions of recombination suppression. In particular, comparative analyses of synonymous substitution rates (Ks) among gene alleles (of protein coding DNA) from various segments of the N. tetrasperma mat chromosomes (from an older and from a younger

evolutionary stratum – stratum 1 and stratum 2, respectively – and from two pseudoautosomal regions, using 290 genes in strain P4492) has revealed that recombination and/or gene conversion events are prevalent at the earliest stages of recombination suppression (Whittle et al. 2011). Specifically, due to the self-fertilizing nature of N. tetrasperma, we assumed a very low sequence divergence (e.g., Ks¼0) between the mat chromosomes in regions of recombination or gene conversions (as gene conversions are associated with recombination and limit divergence; Szostak et al. 1983; Ohta 1989) whereas Ks>0 was presumably indicative of restricted recombination (Menkis et al. 2008). The data showed that the region of suppressed recombination contains segments with genes having Ks>0 interspersed with cases wherein Ks¼0, wherein the later cases are particularly frequent in the younger stratum (stratum 2) as compared to the older stratum (stratum 1; Whittle et al. 2011). Thus, the data suggest that recombination suppression does not arise uniformly across a particular genomic region in N. tetrasperma mat chromosomes, but rather emerges gradually and in a manner that affects a greater fraction of genes per strata over time. Such empirical data supports earlier postulations that recombination suppression might arise in a gradual manner in sex chromosomes (Brooks 1998; Charlesworth et al. 2005). Additional genomic data for gene alleles from the nine distinct N. tetrasperma phylogenetic species has also shown that specific recombination and/or gene conversion events have occurred in the region of suppressed recombination during the history of this taxonomic group (Menkis et al. 2010), and thus further supports a role of these genomic events in early mat chromosome divergence (Menkis et al. 2010). Gene conversion and recombination events could play a critical role in countering the deleterious effect of suppressed recombination (which leads to the accumulation of mutations, e.g., Muller’s ratchet; Charlesworth and Charlesworth 2000; Charlesworth et al. 2005; Khakhlova and Bock 2006) in the early stages of sex chromosome evolution. The genomic data for N. tetrasperma mat chromosomes (strain P4492, see above) has also revealed evidence that genomic degeneration arises within the early stages of recombination suppression. Specifically, comparative analyses of preferred codon usage, one of the key traits


associated with genomic adaptation (for efficient and accurate translation; Duret and Mouchiroud 1999; Duret 2000; Stoletzki and Eyre-Walker 2006), have shown that marked levels of genomic deterioration (toward non-preferred codons) are inherent to early stages of suppressed recombination in the mat chromosomes. Furthermore, the patterns of changes in preferred codon usage among genes and among gene alleles have lead to the identification of key traits associated with the process of genomic degeneration. These include findings that: the earliest stages of recombination suppression are characterized by marked and largely independent degeneration in preferred codon usage among gene alleles, that the level of degeneration is magnified over greater time periods of recombination suppression, with less deterioration in stratum 2 (the younger stratum) than stratum 1, that both mat a and mat A chromosomes are subjected to degeneration (such that one chromosome has not been assigned the “degenerative” chromosome early in sex chromosome evolution), and that the rate of degeneration in preferred codon usage is greater for shorter genes (which have evolved for high efficiency) than for longer genes, suggesting these genes may play an especially significant role in sex chromosome evolution. Moreover, comparative analysis of preferred codon usage and GC content of introns with GC content at third codon positions (GC3) among genes and gene alleles located on the mat chromosomes, has revealed that these results are best explained by reduced selective efficiency for preferred codon usage in the region of suppressed recombination and are not attributable to mutational and/or selection bias (Whittle et al. 2011). The accumulation of non-synonymous substitutions has also been reported for the region of suppressed recombination (Whittle and Johannesson 2011). Thus, the data demonstrate that shifts in selective efficiency underlie genomic degeneration in early stages of recombination suppression in mat chromosomes.

III. Key Findings Regarding the Evolution of Mating-Type Genomic Regions in Other Fungal Model Systems Key information regarding the evolution of mating-type loci and/or chromosomes has recently

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been revealed from research based on model species of basidiomycetes, including Cryptococcus neoformans and Microbotryum violaceum, the filamentous ascomycete Ophiostoma novo-ulmi and the basal fungus Phycomyces blakesleeanus. In particular, molecular evolutionary analysis and/ or population level studies in these species have revealed possible mechanisms underlying matingtype locus evolution and parallels to animal/plant sex chromosomes.

A. Cryptococcus neoformans Recent studies of the mating-type loci in the heterothallic basidiomycete fungi C. neoformans have suggested that these relatively large DNA regions, that is, larger mating-type loci than in filamentous ascomycetes, share marked parallels to sex chromosomes in higher eukaryotes. Unlike the tetrapolar basidiomycetes, C. neoformans contains only a single bipolar MAT locus that encodes both pheromones and pheromone receptors as well as homeodomain protein genes. This locus is relatively large, spanning over 100 kb and 6% of the chromosome (note that the mating-type loci are normally less than 6 kb in filamentous ascomycetes; Coppin et al. 1997), consists of more than 20 genes with no signs of degeneration, lacks mating type-switching and is recombinationally suppressed (Lengeler et al. 2002).

Comparative examination of this large MAT locus among subspecies of C. neoformans and to the bipolar mating system of the ascomycete S. cerevisiae and the tetrapolar system of basidomycete Ustilago maydis has led to the theory that this recombinationally suppressed locus has likely evolved in a manner similar to that observed in sex chromosomes in animals. Specifically, the analysis suggests that the C. neoformans MAT locus, which is highly divergent from other basidiomycetes, likely originated through the acquisition of sex-determining genes into one of two gene clusters (Fraser et al. 2004). Subsequently, the DNA regions fused through translocation events, resulting in an intermediate tripolar mating system, which converted to a bipolar system through gene conversion and/or recombination. Subsequent gene conversion events and inversions, which suppress recombination, likely gave rise to recombination suppression and the current

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MAT loci (Fraser et al. 2004). These traits largely correspond with the steps involved in the evolution of animal/plant sex chromosomes (Fraser et al. 2004). Parallels with the human Y chromosome were also identified in C. neoformans including the fact that there is a clustering of genes with common functions; for example, a majority of genes encoded by the predicted ancient MAT locus are involved in pheromone production and sensing (Fraser et al. 2004). Nonetheless, it is notable that this large recombinationally suppressed MAT locus differs from human sex chromosomes in that one of the MAT chromosomes has not degenerated and lost functionality, as the human Y chromosome (Charlesworth and Charlesworth 2000), nor is it reduced in size; each of these traits could be explained by the fact that unlike humans these fungi also exist in a haploid stage during parts of their life cycle wherein selection may act to preserve gene functionality (Fraser et al. 2004). Overall, these findings from C. neoformans, combined with data from N. tetrasperma, highlights a high degree of convergent evolution of sex chromosomes among animals, plants and fungi (Fraser et al. 2004; Menkis et al. 2008). Evidence from C. neoformans has also provided insight into the possible mechanisms driving mating-type chromosome evolution. Specifically, data from this species suggests that the MAT locus may be subjected to elevated recombination events within the genome (Hsueh et al. 2006). In particular, evidence indicates that although recombination is generally suppressed between the two MAT alleles/idiomorphs in this species, recombination is markedly higher in the immediate MAT flanking regions as compared to rest of the genome (Hsueh et al. 2006). In addition, the enhanced recombination is observed for both homozygous and heterozygous alleles, suggesting that this is not attributable to poor alignment during meiosis, but is generally inherent to the MAT locus. Accordingly, it has been proposed that recombinational activation in this region may play a key role in shaping the composition of the MAT locus, including the clustering of genes involved in specific metabolic and biosynthetic processes (Hsueh et al. 2006). These findings suggest that the MAT locus may be switched onto different genetic backgrounds, which could play a key role in the steps involved in the evolution of this locus in C. neoformans (Lengeler et al. 2002; Fraser et al. 2004) and potentially other fungal species.

B. Microbotryum violaceum In many fungi, including the basidiomycete Microbotryum violaceum, mating compatibility is determined during a haploid stage of development

(Hood 2002). This differs from the pseudohomothallic Neurospora tetrasperma, which is heterokaryotic throughout the majority of its life cycle. It also differs markedly from most animals where the sex chromosomes are maintained at the diploid stage throughout the vast majority of an organism’s lifespan (except for the gamete stage); it has been postulated that the diploid nature of animal sex chromosomes facilitates the sheltering of mutations from selection in the permanently heterozygous chromosome and gives rise to mutation accumulation (e.g., Y chromosome in humans) via processes such as genetic hitchhiking and Muller’s ratchet. The latter process is defined by an accumulation of mutations in an irreversible manner due to lack of genetic recombination (Felsenstein 1974; Rice 1987; Charlesworth and Charlesworth 2000; Hood 2002). Because the mating-type specific chromosomes in M. violaceum experience haploid selection, it is believed that they would be expected to have limited opportunity for sheltering, since only genes that are nonessential for haploid growth should be sheltered and accumulate mutations (Hood 2004). Analysis of genome data from M. violaceum has shown that the mating-type chromosomes in this species contain high levels of transposable elements as compared to the autosomes, are dimorphic in size and have unequal levels of functional genes (Hood 2002; Hood 2004), thus sharing many traits inherent to the dimorphic sex chromosomes of animals and plants. These findings have been proposed to indicate that restricted recombination in itself is sufficient to account for the degeneration in sex chromosomes and that this process does not require chromosomal sheltering (Hood 2004). Subsequent data, however, has questioned this proposition. In particular, an analysis of gene linkage and gene divergence within and among the M. violaceum mating-type chromosomes (Votintseva and Filatov 2009; see below) has shown that the recombinationally suppressed segment comprises a small region of the chromosomes (about 1 Mb), and does not include most of the genes analyzed by Hood (2004). This suggests that factors other than restricted recombination, such as random concentration of transposition frequency (Votintseva and Filatov 2009) and/or gene losses on the mating-type chromosomes could explain the observed degenerative patterns. Further studies will be needed to explain the findings of degeneration (Hood 2004) in


regions with normal recombination (i.e., absence of suppressed recombination; Votintseva and Filatov 2009) within the M. violaceum matingtype chromosomes. Recent DNA sequence analysis and genetic mapping of a sample of genes from the matingtype chromosomes (A1/A2) in M. violaceum has demonstrated that the recombinationally suppressed mating-type region spans about a 25% contiguous segment of the entire chromosomes in this species (and 1 Mb of the chromosome, which is notably smaller than the region in N. tetrasperma; Menkis et al. 2008; Votintseva and Filatov 2009). Comparative analysis of A1/A2 divergence levels suggests that the mating-type chromosomes are comprised of three distinct strata: one oldest stratum with very high divergence, one with about 5–9% divergence and one with 1–4% divergence, which is consistent with the presence of evolutionary strata as found in the mat chromosomes from N. tetrasperma and in the sex chromosomes from animals and plants (Lahn et al. 2001; Fraser and Heitman 2004; Handley et al. 2004; Nicolas et al. 2004; Menkis et al. 2008; Volintseva and Filatov 2009). In addition, certain genes from the recombinationally suppressed region of the mating-type chromosomes were found to amplify (by PCR) from the haploid genome of only one of the two mating types, potentially reflecting substantial degeneration and/or rapid sequence divergence (Votintseva and Filatov 2009). Altogether, the data suggests that there are marked parallels between the regions of suppressed recombination of the mating-type chromosomes of this basidiomycete, the filamentous ascomycete N. tetrasperma and the mature sex chromosomes inherent to plants and animals.

C. Evidence of Interspecific Gene Transfer from Ophiostoma novo-ulmi Although not an extensively researched model species for the mating-type locus evolution per se, population-level studies among the filamentous ascomycete Ophiostoma nova-ulma, which causes Dutch elm disease, has provided key insights into one of the possible mechanisms influencing the evolution of the mating-type locus. In particular, data suggests that MAT locus composition in this species may be altered

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by interspecific gene transfers. This species, which originated in Europe as a clonal population containing only a MAT-2 locus and a mating incompatibility locus (VIC), has spread into worldwide populations. This global expansion has been accompanied by the acquisition of a MAT-1 locus as well as several incompatibility loci (Brasier 1988). Genetic mapping and sequence analysis within and among populations has shown that the newly acquired genes have been transferred to O. nova-ulma from another species, O. ulma (Paoletti et al. 2006), a process that has occurred independently many times during the species expansion. This process may be facilitated by the enhanced selective advantage of the heterogeneous genomic composition (MAT-1/MAT-2 combinations, various VIC loci) against infection by viruses (Brasier 1988; Paoletti et al. 2006). The MAT locus in particular may be subjected to rapid adaptive evolution since a virus exclusion mechanism during reproduction acts to limit vulnerability to parasites, including viruses (Paoletti et al. 2006). These findings demonstrate that interspecific gene transfer, which has historically been believed to primarily occur in prokaryotes, is inherent to the MAT and VIC loci in this group of fungi. Further data is needed to ascertain whether this phenomenon, which could play a key role in mating-type locus evolution, is inherent to a wide group of fungal species.

D. Mating-Type Loci in a Basal Fungus Recent research in Phycomyces blakesleeanus (Zygomycota) by Idnurm et al. (2008) has provided insight into the regulation of sexual traits in this early diverged fungus (Dyer 2008). Specifically, two genes, SexM and SexP have been identified and shown to comprise an early form of the mating-type locus. This conclusion is based on the findings that each gene is strain-specific, upregulated during mating and is linked to the sexual genotype of progeny produced among crosses. In addition, rare strains containing both SexM and SexP were found to be partially self-fertile (Idnurm et al. 2008). Meiotic mapping delimited the sex locus within a 38 kb region encompassing the sexM/sexP genes within the ~65 Mb genome. Further evidence that these genes are ancestral forms of the mating-type locus includes findings that the Sex genes are flanked by genes conserved

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among mating types, have asymmetrical positions on the chromosomes, and that the genes are inverted relative to each other, similar to features found at the mat locus in filamentous ascomycetes such as Neurospora (Debuchy and Turgeon 2006; Idnurm et al. 2008; Menkis et al. 2008). The findings of the HMG-based transcription factor as an ancestral form of the mating-type locus led to the hypothesis that the current locus composition in filamentous ascomycetes, wherein the matingtype locus is characterized by the presence of HMG and a-domain transcription factors and absence of homeodomain protein genes, is most parsimoniously explained by the gain of an a-domain protein gene and the conservation of the HMG-protein gene in this lineage following divergence from the zygomycetes (Dyer 2008). The sex locus has also been identified in other zygomycetous fungi within the Mucorales, including Rhizopus oryzae and Mucor circinelloides (Idnurm et al. 2008; Lee et al. 2008). Thus, the comparative analysis of these highly divergent taxa provides a baseline of the large-scale events (gene losses/gains) in the long-term history of the mating-type locus within fungi.

IV. Future Directions Although marked advances have been made in our understanding of the evolution of matingtype loci in fungi, including the filamentous ascomycetes, many key factors remain to be ascertained, including: (i) the steps giving rise to the various types of mating-type loci among ancestral and extant fungal species, (ii) the co-evolution of mating-type loci and genes involved in reproduction, (iii) the interaction between mating systems and mating-type loci evolution, and (iv) the characterization of mating-type loci among a large range of taxa. As demonstrated herein, the various model taxa from Neurospora contains valuable features that allow the identification and characterization of traits associated with its mat locus, such as the novel evolutionary patterns of the locus across lineages. Future research of the mat chromosomes of N. tetrasperma holds promising new opportunities to reveal the key molecular features associated with early stages of sex chromosome evolution, such as the rate of degeneration, the level of gene silencing, and the role of expan-

sion of the region of suppressed recombination on sequence/gene evolutionary rates among the distinct subspecies. Such data will be highly complementary to the growing data derived from the mating-type loci and chromosomes with suppressed recombination in basidiomycetes (e.g., C. neoformans, M. violaceum) which also share features of highly evolved eukaryotic sex chromosomes. These basidiomycete model species are also likely to continue to serve an essential role in the identification of molecular traits (e.g., elevated recombination) among highly complex mating-type loci. In the future, population level studies such as that inherent to O. novo-ulmi, demonstrating the presence of interspecies gene transfers at the MAT locus, will also likely prove essential for further revealing the in vivo dynamics contributing to mating-type locus evolution. A key factor in future research includes the recent identification of the mating-type loci in the basal Zygomycota (P. blakesleeanus); this can serve as an effective starting point to further reveal the patterns of gene gains, losses and molecular evolution in the mating-type loci among ascomycetes, basidiomycetes and basal fungal species. Overall, the conglomeration of molecular evolutionary analysis and population level genomics research across various fungal model systems will be essential for further disentangling the evolutionary steps giving rise to the complex array of matingtype loci and mating-type chromosomes in fungi. Acknowledgements The authors gratefully acknowledge research funding to C.A.W. from The Royal Swedish Academy of Sciences (Hierta-Retzius Research Grant), The Royal Physiographic Society in Lund, The Lars Hierta Minne Foundation and The Wenner-Gren Foundation, and research funding to H.J. from The Swedish Research Council. We thank Joseph Heitman and Michael Hood for useful comments on an earlier draft of the manuscript.

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Szostak JW, Orr-Weaver TL, Rothstein RJ, Stahl FW (1983) The double-strand break repair model for recombination. Cell 33:25–35 Tajima G (1983) Evolutionary relationship of DNA sequences in finite populations. Genetics 105:437–460 Takahata N, Nei M (1985) Gene genealogy and variance of interpopulational nucleotide differences. Genetics 110:325–344 Tsong AE, Tuch BB, Johnson AD (2007) Rewiring transcriptional circuitry: mating-type regulation in Saccharomyces cerevisiae and Candida albicans as a model for evolution. In: Heitman J, Kronstad JW, Taylor JW, Casselton LA (eds) Sex in fungi: molecular determination and evolutionary implications. ASM Press, Washington DC, pp 75–89 Tucker PK, Lundrigan BL (1993) Rapid evolution of the sex determining locus in Old World mice and rats. Nature 19:715–717 Turgeon BG, Bohlmann H, Ciuffetti LM, Christiansen SK, Yang G, Schafer W, Yoder OC (1993) Cloning and analysis of the mating type genes from Cochliobolus heterostrophus. Mol Gen Genet 238:270–284 Votintseva AA, Filatov DA (2009) Evolutionary strata in a small mating-type-specific region of the smut fungus Microbotryum violaceum. 182:1391–1396 Wendland J, Vaillancourt LJ, Hegner J, Lengeler KB, Laddison KJ, Specht CA, Raper CA, Kothe E

(1995) The mating-type locus B alpha 1 of Schizophyllum commune contains a pheromone receptor gene and putative pheromone genes. EMBO J 14:5271–5278 Whittle CA, Johannesson H (2011) Evidence of the accumulation of allele-specific non-synonymous substitutions in the young region of recombination suppression within the mating-type chromosomes of Neurospora tetrasperma. Heredity DOI:10.1038/ hdy.2011.11 Whittle CA, Sun Y, Johannesson H (2011) Degeneration in codon usage within the region of suppressed recombination in the mating type chromosomes of Neurospora tetrasperma. Eukaryot Cell 10: 594–603 Wik L, Karlsson M, Johannesson H (2008) The evolutionary trajectory of mating-type (mat) genes in Neurospora relates to reproductive behaviour of taxa. BMC Evol Biol 8:109 Yun SH, Turgeon BG (1999) Molecular comparison of mating-type loci and adjacent chromosomal regions from self-sterile and self-fertile Cochliobolus species. Plant Pathol J 15:131–136 Yun SH, Berbee ML, Yoder OC, Turgeon BG (1999) Evolution of the fungal self-fertile reproductive life style from self-sterile ancestors. Proc Natl Acad Sci USA 96:5592–5597

13

Evolution of Special Metabolism in Fungi: Concepts, Mechanisms, and Pathways

CHRISTINE SCHIMEK1

CONTENTS I. Scope and Limitations . . . . . . . . . . . . . . . . . . . . . . . . . . . . 293 II. Secondary Metabolites, Special Metabolites, and Special Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . 293 III. Studying Special Metabolism . . . . . . . . . . . . . . . . . . . . . 295 IV. Evolution of Metabolism: General Considerations. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 296 V. Evolution of Metabolism: Mechanisms and Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 298 A. Basic Integrated Metabolism . . . . . . . . . . . . . . . . . . 299 1. Cellobiose Dehydrogenase . . . . . . . . . . . . . . . . . 299 2. Arabinitol Dehydrogenase . . . . . . . . . . . . . . . . . 300 3. Xylose Metabolism in Piromyces sp. E2 . . . 300 4. Alternate Pathway Localization for Fatty Acid Degradation . . . . . . . . . . . . . . . . . . . . 300 B. Carotenoids. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 301 1. Carotenoid Biosynthesis and Modifications 301 2. Trisporoid-Mediated Sexual Communication in Zygomycetes . . . . . . . . . . 302 C. Evolution of Proteins and Enzymes . . . . . . . . . . . 303 1. Biochemical Considerations . . . . . . . . . . . . . . . 303 2. The Role of Aromatic Prenyl Transferases in Indole Alkaloid Biosynthesis. . . . . . . . . . . . 305 D. Evolution of Special Metabolite Diversity . . . . 306 1. Gene Duplication . . . . . . . . . . . . . . . . . . . . . . . . . . . 306 2. Assembly of Functionally Linked Gene Clusters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 306 3. Horizontal Gene Transfer . . . . . . . . . . . . . . . . . . 309 4. Adaptive Evolution . . . . . . . . . . . . . . . . . . . . . . . . . 311 5. Regulation of Special Metabolite Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 311 E. Gene Clusters for Specific Metabolic Pathways. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 312 1. Metabolic Gene Clusters in Yeast. . . . . . . . . . 312 2. Biosynthesis of b-Lactam Antibiotics . . . . . 314 3. Biosynthesis of Epipolythiodioxopiperazine Toxins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 316 4. Gibberellin Biosynthesis . . . . . . . . . . . . . . . . . . . 317 5. Ergot Alkaloid Biosynthesis. . . . . . . . . . . . . . . . 319 6. Other Non-Ribosomal Peptide Synthetases and Polyketide Synthase Pathways . . . . . . . . 320 VI. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 322 References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 322 1

Department of General Microbiology and Microbe Genetics, Institute of Microbiology, Friedrich-Schiller-Universita¨t Jena, Neugasse 24, 07743, Jena, Germany; e-mail: [email protected]

I. Scope and Limitations Metabolism is one of the key characteristics of life, implicating that evolution always had an impact on metabolic processes. As opposed to studying genealogies and confirming the phylogenetic relationships between organisms, analyses of the changes and variations in metabolic processes help to discover some of the underlying evolutionary mechanisms. Studies on fungal metabolism have traditionally concentrated on metabolites of specific interest, namely mycotoxins, pathogenicity factors, antibiotics, and other compounds displaying interspecific activity. Over time, a vast collection of facts and data relating to fungal metabolism has accumulated, which is now available for comparison and analysis. Piece by piece, a picture of the evolutionary history of some metabolic pathways emerges. As evolutionary mechanisms as well as the underlying chemical, biochemical and biophysical laws and limitations apply to all kingdoms of life, for the present review I made use of several theory articles on metabolic evolution, as well as some more specific publications on the situation in plants, in addition to reviews and original articles covering all facets of metabolic evolution in fungi.

II. Secondary Metabolites, Special Metabolites, and Special Metabolism Though the term ‘special metabolite’ is less qualifying than any of the others and will therefore be used throughout this work, the substances in question are generally referred to by a number of other names: More familiar terms to classify these compounds are ‘secondary metabolites’ or ‘natural products’, reflecting the respective viewpoints of biologists or chemists dealing with them Evolution of Fungi and Fungal-Like Organisms, The Mycota XIV S. Po¨ggeler and J. Wo¨stemeyer (Eds.) © Springer-Verlag Berlin Heidelberg 2011

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(see Chapter 10 in this volume). These classifications mirror the traditional view on the importance and functions of these compounds and the related pathways. The concept of distinct metabolic types was introduced by Albrecht Ko¨ssel (1891) and Friedrich Czapek (1922–1925), and it aimed at the description of specific features of plant metabolism. The idea was that all the basic processes of the cell are governed by a primary metabolism producing primary metabolites and that primary metabolism is basically the same for all organisms. In contrast, secondary metabolites are synthesized via distinct pathways, and only in certain species or lineages, and were supposed to be of less general importance. This description was never stringent, as any number of compounds (e.g., polysaccharides, lipids, terpenoids) could not be classified unambiguously into either of these two categories. The definition was later expanded to include the wealth of compounds and metabolic pathways newly identified in all types of micro-organisms. With a certain overlap, such metabolites are often also referred to as ‘natural products’, usually in a mainly chemical or systematic context. With accumulating data, the perception of secondary metabolism changed over time, and this became most conspicuous when the functional propertie of many of these metabolites became recognized. While in the beginning, the secondary metabolites were seen merely as byproducts of primary/basic metabolism with basically no definite function at all, later they were interpreted alternately as metabolic waste, laboratory artefacts, detoxification products, or products of overflow or luxurious metabolism, with the tenor on them being evolutionarily neutral compounds. These classifications became untenable for two reasons. First, it became more and more apparent that the biosynthesis of these compounds was controlled and determined by a complex genetic background. Second, more and more secondary metabolites were identified as chemical weapons, defence compounds, communication signals, or as being involved in metabolite transport. As a compound active only in another organisms or in another cell can exert its activity only if there exists a molecular target or receptor, all these functions imply close interactions, crossadaptations between the interacting partners, and, in the long run, co-evolutionary processes between the organisms involved. As a result, usually a single group of secondary metabolite

dominates within any given lineage (Wink 2003). Considering all these facts, there is certainly nothing second-rate about the so-called secondary metabolites. At the gene level, a similar distinction is evident. Whereas the ‘core genes’ are widely distributed and code for proteins with central functions in the cell, some others (the ‘accessory genes’) are patchily distributed across the lineages and code for taxon-specific functions (Anderson 2009). This general notion is more and more corroborated by whole genome comparison analyses. The high specificity of special metabolism is evident, for example, in Aspergillus spp. where 68% of the A. fumigatus (teleomorph: Neosartorya fumigata) special metabolite biosynthesis genes are absent in the closely related A. clavatus (Perrin et al. 2007), and even more were found to be missing in the more distant A. oryzae and A. nidulans (teleomorph: Emericella nidulans; Nierman et al. 2005). Special metabolites fulfil a number of roles in the life of fungi: Some are virulence factors involved in the recognition of hosts in pathogenic interactions, such as, for example, the product of the avirulence conferring enzyme 1 (ACE1) gene cluster in Magnaporthe grisea (Collemare et al. 2008), others exhibit broad antibiotic activities. The carotene-derived trisporoids mediate recognition between mating partners in zygomycete fungi (Schimek and Wo¨stemeyer 2006; Wo¨stemeyer and Schimek 2007). Mycotoxins may either be decidedly host-specific or act as unspecific toxins in pathogenic interactions, where they do not benefit the producer directly and may only be apparent after the death of the pathogen (Hof 2008). Often the biological or physiological significance of a metabolite or its function and benefit for the fungus are not known at all. Many such metabolites are only produced by a small group of fungi, such as the loline alkaloids, insect feeding deterrents produced by the Epichloe¨/Neotyphodium fungal endosymbionts of grasses (Schardl 2001). These metabolites most probably play a role in niche adaptation and survival and are themselves clearly the product of long-going coevolutionary processes, as the alkaloids are only produced when the fungi are living associated with their host grasses (Tanaka et al. 2005a). The inverse aspect of co-evolutionary adaptation has been established for lichen fungi where lichenicolous taxa are more efficient in parasitizing on lichens than non-lichenicolous taxa because,


during the long period of co-evolution, they became tolerant to lichen defence metabolites and therefore better suited to occupy this ecological niche (Lawrey 1995). During adaptive responses between interaction partners, the nature of signals may change from negative to positive when, for example, a former defence compounds is now used as a recognition signal, or when allelochemicals are recruited for specific new tasks. But a fluid transition, hallmark of ongoing evolution, always exists between the categories: Several signalling compounds acquired essential functions during evolution, but these are essential predominantly during specific stages of the life cycle, for example, reproduction or differentiation. Such functions are know, among other classes, for many terpenoids, among them the gibberellins (diterpenes), abscisic acid (sesquiterpene), and carotenoids (tetraterpenes). The capacity for the production of special metabolites, including the virulence factors and toxins, must have evolved because it provides a selective advantage. Most fungi and many prokaryotes (e.g., the antibiotic-producing Streptomycetes; Challis and Hopwood 2003) are soil micro-organisms. Soil represents a very complex habitat where the physical conditions are subject to rapid changes. Organisms in touch with this habitat are also confronted with a high microbial density and the concurrent metabolic cacophony of chemical signals that constantly need to be interpreted and answered. Such highly interactive surroundings provide ample driving force for the high variability of special metabolism. In the root sphere continuous contact with other organisms is inevitable and communication and reaction to these communities becomes an issue of vital importance. This involves such basic survival strategies as avoiding feeding damage by other soil inhabitants. A. nidulans is more susceptible to fungivory by the springtail Folsomia candida when LaeA, a global regulator of special metabolism is deleted (Rohlfs et al. 2007) and antibiotic production is therefore decreased. Antibiotics and toxins may also become a currency in interspecies interactions when they protect a host organism from damage by other competitors in exchange for nutrients returned to the producer of the special metabolites. And of course, antibiotics will help to defend resources or even provide nutrients by destroying other micro-organisms.

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In an attempt at a comprehensive description, special metabolites are diverse, complex compounds generally limited to a narrow range of species or even isolates. They are not essential for growth or other basic cell functions and are synthesized via often elaborate pathways from simple precursors that are derived from the basic or essential metabolism. Special metabolites integrated into intra- or intercellular communication pathways will effect a response inside the recipient cell or individual, usually manifested as a change in gene expression. Special metabolism as a whole is understood as comprising all interactions of an organism with its environment. Spontaneous reactions, formerly seen as an essential mechanism in biosynthesis, only play a minor role and wherever they have been studied, were found to be tightly controlled by either the preceding or the subsequent enzyme-catalysed reaction step (Hartmann 2007). In general, biosynthesis of special metabolites requires numerous reaction steps which are catalysed by specific and specifically regulated enzymes.

III. Studying Special Metabolism Studying special metabolism poses a number of peculiar problems. The potential for the production of special metabolites cannot be inferred from studying a single isolate of any given fungal species, as significant metabolic diversity is apparent even among isolates of a single species collected from a single site (Galagan et al. 2005; Seymour et al. 2004). Both the developmental stage and the micro-environment effect metabolism control and thus the metabolite pattern. It is inevitable to conclude that an enormous capacity for variations in special metabolism must exist and that neither the spectrum of possible metabolites nor all possible control levels are as yet fully recognized. This is directly related to the observations that many of the organisms producing special metabolites reside in highly dynamic environments supplying numerous and diverse environmental challenges. Schroeckh and coworkers (2009) recently showed that silent and therefore hitherto unknown genes for special metabolite biosynthesis were selectively activated only in co-cultures of A. nidulans and actinomycetes. Moreover, these interactions were highly

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specific, as only one out of more than 50 actinomycete strains elicited this response, and only at close hyphal interaction with the fungus. As another example, 80% of all genes of Saccharomyces cerevisiae seem to be dispensable under nutrient-rich laboratory conditions. In an experimentally verified computer model for calculating knockout fitness, the majority of these dispensable genes was found to be possibly important under altered conditions (37–68%). For this group of condition-specific genes, a more restricted phylogenetic distribution was determined. From the residual genes, 15–28% are compensated by a duplicate and 4–17% are compensated for by metabolic network flux reorganization. Gene duplicates occur in about the same ratio for dispensable and for indispensable genes, indicating that compensation is not the reason for their retention. The duplicates are probably retained because of selection for high metabolic flux, to speed up the pathway when required (Papp et al. 2004).

A strong bias comes from strictly focusing on chemical aspects of the metabolites, and not on metabolism, in many studies. Often the declared aim is to catalogue natural metabolic diversity or to analyse specific effects of metabolites exerted on organisms in no natural physiological connection whatsoever with the producing organism. In special metabolites, displaying strong biological activity is often interpreted as an indicator of purpose and interpreted accordingly as the result of an evolutionary process. This view is inconsistent with the fact that the majority of metabolites analysed in this context do not benefit the producing organism at all. Data supporting this interpretation moreover often were obtained by screening assays using high concentrations of the compounds to be tested and thus allowing for unspecific reactions and unspecific toxic side effects. Firn and Jones (2000) insist that the trait to be studied in evolutionary studies should not be bioactivity (where toxic or promoting effects are observed at the level of the whole organism) but potency. They define potency as the specific biological activity characterized by the strict structural requirements on small molecules to bind to a given target. In customary screening assays, where a large numbers of molecule species are present, such individual specific interactions will be hard to recognize and observe. With the recent increasing flood of gene and genome data, additional tools for the identification of new metabolites and analysis of their biosynthesis, regulation, and evolution have become

available. Nevertheless, it is not a good idea to discount all known genes and consider everything left as a novelty, as this will certainly lead to serious over- or under-estimation of the true situation. Changed functionality of biosynthesis enzymes and altered metabolites as a result of these pathways could be achieved by minor genetic variations leading, for example, to paralogous domains within an otherwise homologous sequence. Several more promising approaches remain. The analysis of conserved gene neighbourhoods is based on the assumption that linked genes are are more likely to participate in the same cellular function. Probing genome sequence data of a target organism with sequence information for a gene or domain characteristic for a specific pathway will help not only in identification of the gene in question, but also allow scrutinizing its neighbourhood for the rest of the gene cluster. This approach was used e.g., for the the identification of a sirodesmin PL gene cluster in Leptosphaeria maculans (Gardiner et al. 2004). New functional backgrounds of a known protein or domain may also be elucidated in that way. Using sequences derived from sirodesmin PL biosynthesis genes helped to identify the gliotoxin gene cluster in A. fumigatus (Gardiner and Howlett 2005).

Analysis of protein domain architecture may help in determining the participation of multidomain – multifunctional enzymes in special metabolite biosynthesis and shared properties because, for example, hydrophobicity patterns may indicate new members of protein families known for their catalytic function in special metabolite pathways. Co-expression analysis or the overexpression of transcription factors implicated in the regulation of special metabolite biosynthesis pathways is also frequently used to identify unknown pathways and their end-products (e.g., Bok et al. 2006; Bouhired et al. 2007).

IV. Evolution of Metabolism: General Considerations The concept of metabolic evolution and research into this field first and foremost needs to address the question what exactly is the main focus. All evolutionary research is based on the comparison of recent phenomena with the intention to deduce their development over time. The fundamental


question is, what should be compared? There are the metabolites themselves, of course, but the mechanisms leading to their existence also underlie evolution. And disregarding the fact that functions also may evolve may lead to serious mis-interpretations indeed. It is also important, and nowhere near to obvious, to make a decision on which organisms or traits should be compared: those that are closely related, or preferably the rather distant ones. Following the profound analysis presented by Firn and Jones in two review articles (Firn and Jones 2000, 2009), evolution, first and foremost, does not act on organisms or specific traits, but acts upon their inherent functional mechanisms. Therefore it is considered imperative not only to study the evolution of individual enzymes, but also to study and define the evolution of metabolism as a whole. The central question in this attempt would be: which rules determine whether a new biosynthetic capacity arising from mutation will be retained in the organism? Many metabolites serve as pheromones, hormones, or regulatory compounds in inter-organismic, intercellular, or intracellular communication. Such compounds need a receptor to adequately fulfil their task. One of the more interesting questions therefore is the evolution of the ligand/receptor couple. It is generally accepted, that such a pair of molecules originates from a co-evolutionary process, meaning that the ligand and its receptor underwent parallel adaptive evolution. If the ligand is a peptide or protein, that is, if both the ligand and its receptor are directly encoded by genes, the concept for this evolution is basically similar to all kinds of protein–protein interactions, and it is easy to understand that both genes will undergo parallel evolution, each acting as a selective factor on the performance of the other. If the ligand is a small molecule, it is not a gene product, but originates from a complex biochemical pathway where a precursor, either indigenous or from outside sources, is converted into the active ligand in a number of catalytic steps. These pathways usually contain a rate-limiting step which is physiologically regulated and most often also catabolic steps for ligand degradation. It is hard to imagine that simple parallel evolution could work here, although some connection between the ligand and its binding protein must exist. This line of thought can be enlarged to encompass all types of interaction of small molecules with

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proteins, namely also their respective biosynthesis enzymes, and the same rules should govern the evolution of a ligand receptor couple as will direct the overall synthesis of small molecules. Each cell exhibits its own contingent of molecules with their respective properties and functions. The properties are what the cell needs, and these properties are the important feature, not any molecule by itself, nor the function this molecule performs. As well as similar functions may be performed by different metabolites in diverse organisms, a given compound may also exhibit more than one property that might be exploited. The biochemical properties of the individual metabolites determine their value to metabolism, and any given pathway may contribute metabolites to different property classes (Firn and Jones 2009). The shared molecular properties explain the high variability existing and being tolerated in lipids, carotenoids and many special metabolites. In these compounds the molecular properties are not strictly linked to any detailed fine structure, and many similar but not identical structures have common physicochemical properties. New variants of these compounds will retain still similar properties, and will most often be evolutionary neutral, leading to the existence of a diversity of chemical types within a single species, as is the case with the carotenoid trisporoid sexual communication signals in zygomycetes (Schimek and Wo¨stemeyer 2006). This will in the end lead to variations between the species, as each species might use different ratios of all possible compounds. Chemical diversity will constitute an advantage for the producing organisms when the chemicals contribute to its physical defence. The resulting chemical diversity will be tolerated as it may serve as a resource pool for the selection of new physiologically active compounds. All this underlines that it must be considered advantageous for an organism to possess metabolic traits that enhance the likelihood of producing or retaining chemical diversity. This hypothesis serves to explain both the existence of highly potent special metabolites and the retaining of metabolites with no special effects. An animal parallel may help in understanding: for animals, it is of vital importance to have an effective immune system, although most of the individual component products of this system will have no effect, neither positive nor negative, in the producer.

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The tremendous evolutionary variance in special metabolism is put down to the fact that these non-essential metabolites are not as strictly required to fit into an established pathway as the metabolites involved in basic cellular functions. High genetic plasticity and diversity are therefore inherent features of the respective pathways and guarantee adaptive flexibility towards challenges imposed by environmental changes. This plasticity extents to one further level: while generally the absolute number of genes does not correlate at all with the diversity of the possible products, in many organisms and in specific pathways, too, the number of metabolites exceeds the number of genes involved in their biosynthesis. This disproportion suggests that some mechanism must exist that grants the production of more than one product with the same set of enzymes. A review on such mechanisms was published by Schwab (2003) for the situation in plants, but it may safely be assumed that the same basic factors and features also apply to fungi. Much of the metabolite diversity originates from variations of common molecular backbone structures. We also know that post-translational modification plays a major role and leads to the processing of much higher numbers of different mature proteins than are encoded by the genome. Diversity in these contexts can be introduced at every possible level: the DNA code, the mRNA transcript, or the translation product. The mechanisms involved include the usage of alternative reading frames, gene fusion as in the carotene synthesis genes carRP (Mucor circinelloides; Velayos et al. 2000) and crtYB (Xanthophyllomonas dendrorhous; Verdoes et al. 1999) coding for enzymes with both lycopene cyclase and phytoene synthase activities, alternative splicing, posttranslational modifications, assembly of multiple proteins into complexes, or altered catalytic specificity.

Of course, selection must also be considered in this context. In chemical interactions, only such target organisms matter evolutionary that indeed have the opportunity to interact with the producer of the active compounds. Others simply cannot act as a focus for selection. In a similarly built argument, only concentrations of the compounds are of any evolutionary importance that really occur under normal circumstances. All the rest may matter for biotechnology and medicine, but not for evolution. Firn and Jones (2009) state that the most common evolutionary scenario for

selection on parts of special metabolism has involved only a few target organisms. If evolution leads to a new compound, the intrinsic properties of this compound are under selection, not the enzyme(s) that produced it: The compound could have properties that either enhance or adversely affect the functions of the cell or organism. It could also have properties that substitute existing necessary properties with either no or negative impact. If the mutation gives rise to several compounds, selection can focus on the properties of either compound and favour a new compound when the cost/benefit ratio is below 1. If the ratio is higher than 1, the new compound will be lost again over time. At the compound level, selection forces will act on the physicochemical characteristics of a molecule. One of these properties is the biomolecular activity determined by the accuracy of ligand– target interaction. The high potency of a compound is the result of a strong ligand–protein interaction and depends on a very specific ligand structure fitting a precise target site on the target protein. These constraints on the evolution of biomolecular activity are considered to be of high importance during the evolution of special metabolism. Tolerating relaxed substrate specificity will increase the capacity to generate chemical diversity. In conclusion, the high substrate specificity of an enzyme as manifested by strict and stringent ligand–target interactions is always the result of intense selection (Firn and Jones 2009). At the organism level, it must be postulated (and it is increasingly supported by solid proof) that there must be evolutionary connections between members of different kingdoms if these are part of a communication network. Understanding of the same signal by different taxa may originate from a simple misunderstanding where an receptor molecule expressed by organism A is not absolutely specific in its ligand-binding properties and interacts with a product of organism B. Such misunderstanding could have been evolutionarily developed into any kind of inter-species signalling.

V. Evolution of Metabolism: Mechanisms and Pathways Any metabolism depends on specific enzyme activities and these in turn have been moulded


by the evolution of proteins. The fusion or division of protein domains as a basic process alters or expands the intrinsic functional properties and gene duplications, horizontal gene transfer (HGT), or rearrangement of gene fragments added to the evolution of enzymes and metabolic pathways. The evolution of metabolism is a process which has accompanied life since its very beginnings. All the basic features were settled a very long time ago and are tightly conserved nowadays. In the beginning, metabolism evolved because chemical diversity was available. It was extended by chance events. This area is categorized by Firn and Jones (2009) as basic integrated metabolism where selection acted not on the intrinsic properties of any new molecules arisen through mutation, but on their compatibility with the existing pathways leading from that point to ensure proper functioning. With time the interactions between adjacent elements of the pathway and also between the diverse pathways contributing to the cellular metabolism became integrated into a vast, strongly interconnected network. The apparent evolutionary conservation and stability of such pathways is a direct consequence of the tight connections within and between the pathways, as it would be difficult to improve any particular function without compromising the functionality of the whole network. The basic integrated metabolism is characterized by the lack of metabolic diversity between different organisms. The more derived metabolic pathways allow for a higher plasticity and free radiation, with the pathways of the basic integrated metabolism acting as donors for new components (Caetano-Anolles 2009).

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material became only possible with the evolution of specialized extracellular degradation enzymes. Among these are the multifunctional cellobiose dehydrogenases responsible for the degradation of cellulose and lignin. Cellobiose dehydrogenase genes are the product of an ancient fusion of two formerly independent genes, recognizable in the gene structure with two independently evolved domains and a linker region between. The functions of the original gene products are retained in the two domains of the holoenzyme, a cytochrome electron transfer moiety with the unique capacity for storing three electrons linked to a FADbinding dehydrogenase, and the cellobiose dehydrogenase in sensu strictu. The flavin-binding domains belong to the glucose–methanol–choline (GMC) oxireductase family, also present in archaea, bacteria, and higher eukaryotes (Cavener 1992). The capability for specific cellulose binding is conferred either by a separate carbohydratebinding module (Subramaniam et al. 1999) or via a cellulose-binding domain on the surface of the FAD moiety (Hallberg et al. 2002). Zamocky and co-workers (2004) propose that the full-length fungal genes share a common evolutionary history starting from one ancestral GMC gene that spread and adapted to its specific function early in fungal evolution. Very soon after branching off this GMC, the cytochrome domain must have been acquired from an as yet unknown origin. This fusion is interpreted as useful by increasing the catalytic efficiency because of the binding of all necessary prosthetic groups to one molecule compared to the separate actions of the cytochrome and the flavoenzyme. The fused gene is supposed to have spread throughout the lineages because it presented an evolutionary advantage for the fungal lifestyle.

A. Basic Integrated Metabolism Compared to the knowledge base on evolutionary aspects of special metabolism in fungi, detailed data on processes categorized into basic integrated metabolism are scarce. Some of them deal with carbohydrate utilization and stress the basic similarity between these pathways and those established in the other kingdoms. 1. Cellobiose Dehydrogenase The fungal lifestyle with its dependency on the extracellular degradation of complex organic

From the amino acid-based sequence comparison of ten cytochrome domains, it became clear that these enzyme domains represent a sequence motif existing exclusively in fungi and that their sequence similarities mirror the phylogeny of the fungal lineages (Zamocky et al. 2004). The ascomycete sequences are more complex, are clearly distinct from the basidiomycete ones, and experienced longer periods of individual evolution, thus reflecting the phylogeny of these two clades. The same applies to the dehydrogenase moiety. Here, the fungal sequences form a distinct clade separated from all enzymes occurring individually in other kingdoms, and also with a distinction between ascomycetes and basidiomycetes. The average pair-wise percentages of sequence identity and similarity for the dehydrogenase FAD domain are 53.4% and 81.4%,

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respectively, those for the cytochrome domain are quite similar with 44.5% and 75.6%. This supports the idea of a long parallel evolution of the two domains. Unfortunately, at the time of the study, zygomycete and chytrid genomes were not available, so information on the situation in the earliest emerging fungi is not included.

2. Arabinitol Dehydrogenase The enzyme L-arabinitol 4-dehydrogenase Lad1 from Trichoderma reesei (teleomorph: Hypocrea jecorina) was found to be an orthologue to plant and animal sorbitol dehydrogenases, probably as an adaptation to the reductive catabolism of the hemicellulose monosaccharides, like L-arabinose or D-xylose, present in the environment of fungi. This classification is based on studies of substrate specify and preferences in the reduction of diverse sugars and on a comparison of sequence data (Pail et al. 2004). In fungi, the enzyme evolved distinctly from the other eukaryotic lineages in that it reduces both pentitols and hexitols, and with a higher catalytic capacity for the first. Nevertheless, all amino acids active in sorbitol (hexose) binding are still present, and the catalytic mechanism has been conserved. The different function pattern is most probably caused by altered amino acids flanking the active cleft of the enzyme (Pail et al. 2004).

3. Xylose Metabolism in Piromyces sp. E2 The anaerobic fungus Piromyces sp. strain E2 metabolizes xylose using xylose isomerase and D-xylulokinase, while the majority of other fungi depend on a pathway via xylose reductase and xylitol dehydrogenase (Jeffries 1983). The situation in Piromyces sp. most closely resembles that in bacteria, because some plant-specific alterations are also not evident (Harhangi et al. 2003). The genes coding for pyruvate formate lyase (Akhmanova et al. 1999) and several extracellular cellulases and xylanases of this fungus (Gilbert et al. 1992; Zhou et al. 1994) also resemble bacterial analogues. It is suggestive to note that the extracellular enzymes provide the substrates for other, intracellular enzymes, so that probably a complete pathway was acquired here via HGT, as was proposed by Garcia-Vallve et al. (2000).

4. Alternate Pathway Localization for Fatty Acid Degradation Despite the inherent functional properties constantly under selective pressure, several other features also mirror evolutionary processes. One of these is the cellular localization of a given pathway, if more than one locus is possible. Fatty acid degradation in the process termed b-oxidation is a sequence of four catalytic steps, resulting in the repeated removal of two carbon units from acetyl-CoA. This pathway is highly diverse and organized differently in the various organismic groups. In mammals, the process takes place in both mitochondria and peroxisomes. In each organelle, a different set of enzymes catalyses identical reactions. In fungi, b-oxidation is also established in both organelle types in Emericella nidulans (Maggio-Hall and Keller, 2004), but in the Saccharomycetes the process is limited to peroxisomes (Hiltunen et al. 1992; Kurihara et al. 1992). Based on the analysis of genome sequence data and the comparison of existing genes and proteins, Shen and Burger (2009) presented an overview on the distribution of this pathway in the fungal lineage. As there is no evidence for the existence of any homologues to the mammalian so-called trifunctional enzyme, characteristic for subtype II of the mitochondrial b-oxidation, only the mitochondrial type I pathway is apparently present in fungi. At least one homologue of each of the four enzyme encoding genes of this pathway, acyl-CoA dehydrogenase, enoyl-CoA hydratase, 3-OH-acylCoA dehydrogenase, and 3-keto-acyl-CoA thiolase, was detected in 30 out of 57 species distributed over all major fungal lineages (Shen and Burger 2009). Usually even gene families exist for one or other of the enzyme encoding genes. This indicates that the intact mitochondrial pathway is present in these 30 species, although in some species probably restricted to only one substrate, butyric acid. An even larger contingent, 48 out of 57 analysed species, contains a functional peroxisomal pathway. A further six species exhibit a modified hybrid pathway, where one of the mitochondrion-specific enzymes functions in an otherwise peroxisomal pathway. Based on the genome comparison data alone, but without experimental proof, the two pathways seem to follow different evolutionary trends. The authors


assume that the mitochondrial pathway got lost early in the common ancestor of all Saccharomycetes, while the Sordariomycetes newly evolved the hybrid peroxisomal pathway. Schizosaccharomyces sp. and Encephalitozoon sp. completely lost both pathways, the latter being an obligate intracellular parasite without typical mitochondria and peroxisomes, while Schizosaccharomyces sp. possesses both types of organelles but is adapted to life in a sugar-rich environmental niche where the usage of fatty acids is probably not needed for survival. In early diverged clades, such as the Zgyomycetes, both pathways are present and apparently functional, similar to the situation in the sister clade animals. In the later diverged lineages, the mitochondrial pathway is lost occasionally, but never the peroxisomal pathway. B. Carotenoids 1. Carotenoid Biosynthesis and Modifications The first steps of carotene synthesis are common to all carotenogenic organisms and are similar to sterol or isoprenoid synthesis pathways (Fraser and Bramley 2004; Sandmann 2002). Enzymes catalysing the various transformations of carotenoids and apocarotenoids are closely related and have been conserved over large evolutionary distances. However, some carotenoids play an important role as signal and communication compounds, placing metabolism of this substance class somewhere between the categories of typical basic and special metabolites. The same applies to lipids and fatty acid-derived substances, such as the oxylipins (Calvo et al. 1999, 2001; Tsitsigiannis et al. 2005, 2006). The early steps of carotenoid synthesis in fungi follow the mevalonate pathway, also providing the substrates for a number of other terpenoid biosynthesis pathways. The first committed enzyme in C40-carotenoid biosynthesis is phytoene synthase, catalysing the condensation of two geranylgeranyldiphosphyte molecules to 15-cis-phytoene. Phytoene synthase is related to the plant and cyanobacterial carotene isomerases, and also to animal retinol saturases catalysing the saturation of all trans-retinol (Moise et al. 2005). Next in the reaction sequence comes a series of desaturation steps. In non-photosynthetic bacteria, archaea, and fungi, the degree of carotene desaturation is determined by a single enzyme of the bacterial crtI type. Four desaturation steps are necessary for desaturation towards lycopene, three steps for neurosporene, and five steps towards 3,4-dehydrolycopene (Sandmann 2001).

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Cyclization of lycopene to b-carotene is performed by a lycopene cyclase derived from the prokaryotic crtY gene. Fungal lycopene cyclase is the product of a fused gene, crtYB, composed of lycopene cyclase and phytoene synthase functional domains. The lycopene cyclase moiety is probably derived from an archaeal enzyme (Peck et al. 2002), also a type 2 lycopene cyclase, as opposed by the type 1 enzymes present in most gram-negative bacteria, cyanobacteria, and plants. The archaeal enzyme consists of two small homologous subunits, CrtYc and CrtYd, supposed to be the result of a gene duplication. The same tandem fusion occurs in the fungal enzyme, where the 200 amino acid residues constituting the lycopene cyclase are localized at the N-terminus, while the phytoene synthase makes up the C-terminus (Hemmi et al. 2003).

The genes coding for the carotene synthesis enzymes are physically linked in all fungi harbouring this trait. Carotenes are frequently further modified by oxidation or addition of various functional groups to the isoprene chain. Other pathways involve enzymatic cleavage at an internal C¼C double bond by carotene cleavage oxygenases using dioxygen, for example the cleavage of violaxanthin as the first step in plant abscisic acid synthesis (Li and Walton 1990), cleavage of b-carotene to retinal, the first step leading to retinol and its subsequent metabolites with functions in vision, immunity, and development (Prado-Cabrero et al. 2007; von Lintig and Vogt 2004), and the cleavage of b-carotene to produce trisporoids (Burmester et al. 2007). The reaction is either limited to one such bond, or the two bonds at structurally similar positions at both sides of the substrate molecule. The reaction products are the corresponding aldehydes or ketones, which are frequently further modified to form the functional apocarotenoids (Kloer and Schulz 2006). The generally high substrate specificity displayed by the carotene cleavage oxygenases is in part determined by the shape and length of a tunnel taking up the substrate and threading it past the active centre. The length of the tunnel determines the cleavage position, while the shape of the tunnel entrances allows entering of ionone rings or of free polyene chains only. Bends in the tunnel accommodate for trans- or cis-acceptance. These structural features were determined for an apocarotenoid cleaving enzyme, but apparently, the overall structure is conserved within all carotene cleavage dioxygenases (Kloer et al. 2005). The variable features of the diverse carotene oxygenase molecules, though all modelled around a well conserved, stable propeller made up of seven beta-sheet regions, allow for

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alteration of the substrates and thus for evolutionary adaptation and selection. Comparable conservation tendencies are also obvious in the other carotenoid modification reactions. Modification of retinoids involves a number of enzymes classified in the short-chain alcohol dehydrogenase protein family and the medium-chain alcohol dehydrogenases. RPE65, involved in retinoid isomerization, also has high sequence similarities to the 9-cis-epoxycarotenoid dioxygenase protein family. Besides the short-chain dehydrogenases, members of the cytochrome P450 family are also often involved in apocarotenoid or retinoid metabolism. Such enzymes catalyse, for example, the inactivation of abscisic acid via oxidation to 8’-hydroabscisic acid, and the hydroxylation of retinoid acid derivatives. The CYP707 abscisic acid inactivation enzymes share similarities with other enzymes oxidizing terpenoids involved in cellular signalling, such as brassinosteroids, gibberellins, and cytokinins.

The chemistry of the carotenoid molecule limits the possible enzymatic reactions used for the diverse pathways in all kingdoms to the following set: pyrophosphate requirement for the condensation of the isoprenoid units, isomerization around a double bond, cleavage of CH2CH2 or HC¼CH bonds, oxidation to epoxides or ketones, oxidation of alcohols to aldehydes and of aldehydes to acids, hydroxylation, saturation and desaturation of HC¼CH bonds, and carbon–carbon cyclization (for a review, see Moise et al. 2005). Furthermore, there is an extraordinary relationship at the molecular level between enzymes of the different kingdoms. Pathways for the biosynthesis of, for example, abscisic acid and retinoic acid follow a common scheme and are mediated by related enzymes. The animal retinol saturase is also a CrtI-like enzyme, the reverse reaction of the same enzymes is used by plants and cyanobacteria to effect saturation, rotation an desaturation, and thus to change the geometric configuration of the substrate molecule. Even the specifically fungal pathway for direct synthesis of abscisic acid by cyclization and oxidation of farnesyldiphosphate is catalysed by enzymes belonging into the same conserved group. In general, the carotenoid-related metabolic pathways are highly conserved throughout all kingdoms reflecting the limited number of chemical reactions these terpenoids can undergo (Moise et al. 2005). That carotenoid metabolism is retained in so many lineages to such a high degree

reflects the general importance of these molecules. As carotene is the starting point for biosynthesis of a number of vital signalling and communication compounds involved in developmental regulation all over the living world, this is not greatly astonishing. For the carotene cleavage and modification steps leading to these highly diverse metabolites, no specialized committed gene clusters have been identified. This may indicate that the signalling function of apocarotenoids is in part a neofunctionalization which led to the recruitment of different cleavage products in diverse organisms for use as regulators compounds of various pathways. However, carotene cleavage oxygenase genes have been identified as part of carotene biosynthesis gene clusters (e.g., in Gibberella fujikuroi) where the product of the carX gene (Thewes et al. 2005) is involved in auto-regulation of carotene synthesis (Prado-Cabrero et al. 2007).

2. Trisporoid-Mediated Sexual Communication in Zygomycetes One specific fungal example for apocarotenoid signals are the trisporoids, active in sexual communication and regulation of the early sexual development in zygomycete fungi. Trisporoids exhibit a broad distribution; they have been identified throughout the order Mucorales. The identification of one of the biosynthesis genes, coding for 4-dihydromethyltrisporate dehydrogenase in several species outside this order suggests that the same signal system is employed throughout the class Zygomycetes. With this very broad activity domain, trisporoids are comparable to other apocarotenoids (e.g., retinoids, abscisic acid) which are also active across large clades. Detailed analysis of the trisporoid signal system showed, that trisporoids are produced in a number of different derivates differing in substitution groups at certain sites of a common backbone structure (Schimek and Wo¨stemeyer 2006). We could not confirm the assumption by Sutter et al. (1989) that these derivates play a major role in establishing species specificity in mating. In Blakeslea trispora as well as in Mucor mucedo and Phycomyces blakesleeanus, compounds of the B and the C derivate series, differing in the oxidation status at one position near the end of the isoprenoid side chain, elicit the same physiological responses up to the level of transcriptional regulation,


although occasionally with variations in the absolute strength of the response. In contrast, we observed marked differences in the activity pattern of several of the trisporoids biosynthetic enzymes at the transcriptional, translational, and even post-translational level. Certain biosynthesis enzymes being active only in specific regions of the mycelium at defined times throughout the life cycle may in fact prohibit interactions between dissimilar partners in a mating reaction and thus provide an effective mechanism for maintaining species-specificity of the mating reaction (Schimek and Wo¨stemeyer 2009). But maintaining species-specific mating systems must not necessarily be the major driving force for this diverging regulation phenomena. For any individual, finding a member of another species in its close vicinity using the same communication signals as the intended partner will be a rare event. In this encounter, misunderstandings are inevitable and will occur. Such misunderstandings are not necessarily disadvantageous, neither when seen from the individual’s point of view nor for the whole species: 1. Due to a number of different niche adaptations in different species, the chances are low to get into such a situation. There will only be a limited number of possible faux partners in a given habitat. 2. Quite comparable to the situation in mammals, where fur colour or even minor cultural preferences may decide partner recognition and species maintenance, fungi show sufficiently different lifestyles to prevent mating encounters between different species. Some zygomycetes, for example, acquire sexual competence only at restrictive environmental conditions, while others are triggered by highly affluent environments. Other abiotic or biotic factors like light, temperature or the morphology of the hyphae also play a role (for a review, see Wo¨stemeyer and Schimek 2007). Also, trisporoids seem to govern only the early developmental steps, future events during sexual development may depend on yet unknown signals or contact mechanisms. All these factors put together will lead in most cases to an abortion of the ‘forbidden’ interaction at some premature stage. 3. Successful sexual reactions between members of different species provide a valuable tool for interspecies sexual or parasexual recombination and HGT. This has also been found in zygomycetes, where several biotrophic fusion parasites

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parasitize exclusively on other members of the same lineage. For one of these, Parasitella parasitica, the involvement of the trisporoid signal system has been proven. P. parasitica undergoes parasexual development with a number of other Mucorales, and with Absidia glauca as partner species, this encounter is truly mating-type-specific (Wo¨stemeyer et al. 1995). Also, transcription analysis of the 4-dihydromethyltrisporate dehydrogenase gene in cross-species mating with A. glauca revealed that the same cooperative trisporoid biosynthesis system is working here as in bona fide intraspecific mating (Schultze et al. 2005).

Over time metabolic networks underlie a strong tendency towards simplification, which add to the difficulties in selectively altering one component without affecting the whole system (Caetano-Anolles 2009). In eukaryotic organisms, especially the classical special metabolism pathways leading to signalling compounds, this simplification is often achieved by increasing the functional redundancy. One such process might be evident in trisporoid biosynthesis: The molecular characteristics of a new trisporoid biosynthesis enzyme in M. mucedo, 4-dihydrotrisporin dehydrogenase (Wetzel et al. 2009) were insofar unexpected, as this enzyme catalyses the same oxidation step at the C4 carbon in the ionone ring of the trisporoid molecule as the earlier known 4-dihydromethyltrisporate dehydrogenase (Czempinski et al. 1996; Schimek et al. 2005; Werkman 1976), but belongs to to a different protein family. While 4-dihydromethyltrisporate dehydrogenase belongs to the aldo-keto reductase superfamily (Schimek et al. 2005), 4-dihydrotrisporin dehydrogenase is a completely unrelated short-chain dehydrogenase (Wetzel et al. 2009). Other observations indicate that in some zygomycete species only one of the two enzymes is active. This suggests that components of several distinct pathways are recruited into the trisporoid biosynthesis pathway and metabolic streamlining and the evolution of a co-regulatory system is still under way.

C. Evolution of Proteins and Enzymes 1. Biochemical Considerations From the biochemical viewpoint, the qualitative and quantitative diversity in special metabolites is

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mainly determined by the selectivity of the enzymes involved in their production, and the supply of suitable substrates available for the entry into that specific pathway. Substrate availability in turn is influenced by the temporal and spatial expression of the respective synthesizing enzymes, that is, by compartmentalization. Metabolic pathways compete for limited pools of substrates, a limitation that in special metabolism is often countered by metabolic channelling (Hartmann 2007). This encompasses the organization of several or all required enzymes of a synthesis pathway into multienzyme complexes, where the product of one reaction step is passed directly on as substrate to the next enzyme. In such enzyme dimers, multimers or complexes, the interactions between the assembled members influence their respective activities and may lead to emergent new functions. Enzymes act specifically at three levels: they may be specific for a substrate, for the catalytic reaction, and for the product, and poor specificity at any of this level contributes to metabolic diversity (Schwab 2003). Substrate specificity can be more or less strict, but any classification of substrate specificity has always to be assessed by the way the data were obtained. In an in vitro assay with a limited number of different reactants, substrate specificity measurements will lead to different results as in an in vivo approach or in assays using a larger complement of reaction partners, that is, unfractionated protein extracts. Anyhow, quite a number of enzymes show marked ambivalence concerning their substrate, among them the O-methyltransferases (Yabe et al. 1988), terpene synthases (Greenhagen et al. 2006), glycosyltransferases (Klutts et al. 2005), polyketide synthases, and cytochrome P450 (Haarmann et al. 2006; Rojas et al. 2004). Despite the importance of highly specific enzymes that are also involved in the synthesis of special metabolites, the direction of evolution in general tends towards a reduced substrate specificity (Madern 2002). Enzymes can also be multifunctional proteins, and multifunctionality may extent to many different features: Besides an ability to catalyse more than one reaction within one route of synthesis (Bornscheuer and Kazlauskas 2004) as with the lycopene cyclase/phytoene synthase, some enzymes are additionally involved in other cellular processes. Saccharomyces cerevisiae contains a hexokinase that also acts as a protein kinase

(Fothergill-Gilmore and Michels 1993). Many other enzymes catalyse the formation of multiple products or fulfil other than catalytic functions, for example as regulator, transporter, or protease. The N-terminal R-domain of the bifunctional lycopene cyclase/phytoene synthase of Mucor circinelloides (carRP) and Xanthophyllomyces dendrorhous (carYB) functions as membrane anchor for the phytoene synthase domain besides its lycopene cyclase catalytic function (Velayos et al. 2000; Verdoes et al. 1999). The number of multifunctional genes and proteins increases with genome size in eukaryote genomes. Multifunctionality may arise from gene fusion, as has been studied in the oomycete Phytophthora sojae (Morris et al. 2009). The resulting proteins are a valuable tool for deciphering pathway evolution in related species: the fusion of two or more catalytic domains as part of one pathway is often the sign that the individual domains were already involved in that pathway in the ancestors. Identification of any orthology to such an individual domain may therefore serve as identification mark for the complete metabolic pathway. For example, the pentafunctional ARO1 enzyme in the fungal amino acid synthesis pathway is the result of a fusion of five domains acting as individual enzymes in Escherichia coli (Duncan et al. 1987). This shows that the fungal pathway is ultimately derived from an ancestral bacterial one. Gene fusion also enforces co-regulation of the fused domains or co-regulated genes in multienzyme complexes. Some domains in eukaryotic proteins are considered ‘promiscuous’ as they may be combined with different other domains/subunits in proteins associated with signal transduction pathways. Overall, there is nevertheless a broad conservation of domain combinations in multifunctional proteins. Analysis of the P. sojae genome for multifunctional proteins containing interkingdom gene fusions revealed 18 new candidate genes with interaction partners in 39 different species. Partial overlap of the multifunctional metabolic enzymes between the different kingdoms suggests that some of the fusion events are ancient and took place before the split of the lineages. Some fusion events, in contrast, occurred independently in the various lineages, for example the fusion of the orotate phosphoribosyl transferase to orotidine 5’-monophosphate decarboxylase in oophyte and plants. The majority of fusion events for the


novel multifunctional proteins occurred after the division between oomycetes and diatoms took place (Morris et al. 2009). Viewed in still greater detail, a relatively small number of enzymatic mechanisms can account for most of the existing metabolic diversity, and these enzymatic mechanisms depend on the underlying protein structure. Proteins are comprised of a very small set of structural elements made up of peptide folds which may be adapted, combined, or modulated for their specific tasks (Hrmova and Fincher 2006). Small variations in secondary or tertiary structure, mediated by pH effects, variations in the hydration status, oxygen availability, or other such factors may influence the specificity of a catalytic reaction (Schwab 2003). Distributed over all types of metabolism, structural homologues in enzymes occur. These enzymes retain the original catalytic functions and co-factor binding properties but make use of different substrates. It seems to be easier to evolve altered substrate binding sites than complete catalytic mechanisms (Teichmann 2002). This, in turn, may be interpreted as the effect of molecular evolution placing very specific constraints on the spatial interactions within catalytic sites. In this context, as in many others, one of the basic principles in evolution may be stated: it is always easier to change something already existing than to invent or discover something entirely new. 2. The Role of Aromatic Prenyl Transferases in Indole Alkaloid Biosynthesis Prenylated indole alkaloids are compounds comprising an aromatic and an indole moiety. In fungi they are mainly produced by the genera Claviceps, Penicillium, and Aspergillus. The prenylated compounds of these taxa often exhibit biological activities differing from those of the non-prenylated precursor compounds. The prenylated indole alkaloids are moreover a chemically highly diverse group of compounds, as the prenylation may occur at almost any position of the indole ring. The best known compounds in that group are lysergic acid and its derivatives, for example ergotamine. These are produced using tryptophan as substrate for the indole and indoline formation, and dimethylallyldiphosphate as precursor of the prenyl moiety. Many compounds contain an additional amino acid, they then form cyclic

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dipeptides with a diketopiperazine structure or a derivative thereof. The key step in the biosynthesis of these diverse compounds is prenylation, catalysed by prenyltransferases. Using available genome sequence data to identify unknown prenyltransferases in different clusters coding for the biosynthesis of indole derivatives in A. fumigatus and Neosartorya fischeri, the evolution and evolutionary change of the new genes was studied (for a review, see Li 2009). Their gene products were found to show significant similarities to known fungal dimethylallyltryptophan synthases. In biochemical tests of the heterologous expressed gene products several enzymes displayed catalytic promiscuity, one of the basic evolutionary mechanisms in pathways leading to the diversity of special metabolites (Bornscheuer and Kazlauskas 2004). In fact, all indole prenyltransferases show catalytic promiscuity; this can also be interpreted as a strong hint on their evolutionary origin from a single common ancestor (Kremer et al. 2007; Yin et al. 2007). The newly assessed enzymes catalyse the prenylation of tryptophan, but they also show aminopeptidase activity with tryptophan being one of the possible reaction products (Kremer and Li 2008). As an indole moiety is a common feature of prenyltransferase activity, the tryptophan structure could be the link between the two activities of these enzymes. Aromatic prenyltransferases in fungi are different from their bacterial counterparts in that they do not depend on Mg2+, or any other metal ions, although their activity is enhanced in the presence of Ca2+. Some bacterial ABBA prenyltransferases, named for their characteristic secondary structure of five a-b-b-a repeats, also do not require Mg2+. In contrast, the aminopeptidase activities require the presence of divalent metal ions, in this case Mn2+. In biochemical characterizations, the rate constants for aminopeptidase activities are comparable to the hitherto known enzymes, but for prenyltransferase activities, these values vary by up to a factor of 100. 4-Dimethylallyltryptophan synthases from different sources show conserved intron positions and 50–74% identity at the amino acid level. Between prenyltransferases catalysing different reactions, 20–34% identical amino acids are found. From sequence data, no conclusions to the prenylation position and other functional characteristics of the enzymes can be drawn (see Li 2009).

The soluble prenyltransferases from fungi form a clade of their own. Relatively close relationships exist between TdiB of A. nidulans and a prenyltransferase from the cyanobacterium

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Lyngbya majuscule and other bacterial prenyltransferases. Some of these can be seen as transitional elements, as they are clearly situated between the fungal indole prenyltransferases and the bacterial ABBA prenyltransferases in phylogenetic analyses (Li 2009). In short, A. fumigatus and N. fischeri contain a number of putative dimethylallyltryptophan synthases with typical fungal characteristics, but additionally some new aromatic prenyltransferases that are markedly different from all known membrane bound prenyltransferases as well as the soluble bacterial ABBA methyltransferases (Li 2009).

D. Evolution of Special Metabolite Diversity

But if the newly modified gene adequately retains the function of the original gene, for example in altering the substrate specificity of an enzyme but leading to products with the same physiochemical properties, gene duplication would not be needed. The genome sequence data of a pathogenic strain of the zygomycete, Rhizopus oryzae, contain evidence for a whole-genome duplication at some point of the species evolution. Interestingly, most of the duplicated information was lost again, but the retained portions gave rise to several gene families related to virulence, cell wall synthesis and signal transduction. Copies of three complex systems for energy generation and utilization were also retained. The expansion of these gene families probably contributed to the evolution of a pathogenic life style in this species (Ma et al. 2009).

1. Gene Duplication One major driving force in the evolution of special metabolite synthesis pathways is certainly gene diversification and recruitment by gene duplication. In this mechanism, a gene directing an essential function is duplicated by genetic recombination events. As the copied gene is free from stringent control and selective pressure, it may eventually be inactivated. On the other hand it may accumulate modifications and finally be recruited to a new function. Continuous modification of function during speciation gave rise to the large gene families known especially for polyketide synthetases, terpene synthases, glycosyltransferases, and o-methyltransferases. Polyketide synthases assemble multiple acyl units to form fused-ring aromatic polyketides occurring in fungi, lichens, other micro-organisms, and plants. The polyketide chrysophanol is a representative for this high variability: Chrysophanol is a defence compound produced by plants, lichen, fungi, and insects, and was also found in bacteria like Myocardia sp. and Streptomyces sp. It is produced by varying organismspecific folding patterns with clear differences between prokaryotes and eukaryotes (for a review, see Bringmann et al. 2009). Any gene duplicate might as well be recruited for a new, stringent function in a new biochemical environment and there develop into a new strictly controlled single copy gene (Hartmann 2007). Overall, the probability of recruitment is higher when the candidate gene(s) are already within the same subnetwork and when they are quite similar.

2. Assembly of Functionally Linked Gene Clusters Across the kingdoms, the genes involved in the biosynthesis of special metabolites are increasingly found to be organized in physically linked functional groups. In fungi, too, quite a number of special metabolite pathways are organized via gene clusters where the intergenic space between the individual component genes is often less than ca. 2 kb. Unlike the situation in bacteria, these genes are transcribed individually, not as a single transcript. In fungi, pathways controlled by clustered genes include those for the antibiotics penicillin (Smith et al. 1990) and cephalosporin, the polyketides, such as lovastatin, sterigmatocystin, aflatoxin, and xenovulene (Bailey et al. 2007; Brown et al. 1996; Cox 2007; Keller and Adams 1995; Keller and Hohn 1997; Kennedy et al. 1999; Woloshuk and Prieto 1998; Yabe and Nakajima 2004; Yu et al. 2004), gibberellins (Tudzynski et al. 1999), ergot alkaloids (Tudzynski and Holter 1998), trichothecenes (Hohn et al. 1993), and toxins like AK toxin (Tanaka et al. 1999), and the HC toxin of Cochliobolus carbonum (Ahn and Walton 1996; Pedley and Walton 2001; Walton 2006). Another gene cluster is responsible for the production of an unidentified product in M. grisea, involved in host recognition (Collemare et al. 2008). The clusters are made up of the core genes responsible for the individual catalytic steps towards the final metabolite. Some of these genes are so significant for clustered special metabolite pathways that they are used in search


for unknown gene clusters, among them the nonribosomal peptide synthases, type I polyketide synthases, terpene cyclases or dimethylallyltryptophan synthetases (Keller et al. 2005). Other genes commonly found in clusters are oxidoreductases, methylases, acetylases, and esterases, all of them involved in modification of the special metabolites. Occasionally, genes for non-biosynthetic functions are also integrated into a group of functionally linked genes. Gene clusters of filamentous ascomycetes often contain a gene coding for a pathway specific transcription factor acting as positive regulator of the assembled biosynthesis genes. In this context, zinc binuclear cluster transcription factors have been identified (Keller et al. 2005), such as AflR in aflatoxin and sterigmatocystin biosynthesis (Georgianna and Payne 2009; Woloshuk et al. 1994), as well as zincfinger proteins regulating trichotecene biosynthesis (Proctor et al. 1995), and the ankyrin repeat protein ToxE as transcription factor for HC toxin production in C. carbonum (Pedley and Walton 2001).

Metabolic channelling is one of the reasons why genetic linkage groups may persist. The genes contributing to a metabolic pathway are necessarily co-expressed, and both the assembly of the component parts into a multienzyme complex as well as coordinated transcription of these components will be facilitated when the respective genes are physically close. Organization of special metabolite biosynthesis genes into physically linked groups represents a striking feature with a number of evolutionary consequences. As most other genes are dispersed throughout the genome without any specific order and as a number of evolutionary mechanisms (e.g., translocation, inversion, unequal crossing over) contribute to the dispersal of genes throughout the genome, clustering must provide a selective advantage. Walton (2000) undertook to formulate a base for understanding gene clustering as an evolutionary mechanism in its own right. Physical linkage provides a tool for retaining functionally related biosynthesis genes and accessory genes under the influence of their regulators. This is accepted as one factor favouring clustering, as co-regulation by cis-acting regulatory elements or epigenetic mechanisms will be facilitated in closely neighbouring sequence elements. Against the exclusive validity of the hypothesis stands the observations that special metabolism in fungi is generally regulated by trans-acting factors that

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can control dispersed genes as efficiently as clustered ones and that transcription of a number of dispersed genes also is co-regulated (Walton 2000). Walton, whose view was obviously strongly influenced by the concept of the ‘selfish gene’, proposes as further explanation for the origin and maintenance of gene clusters for metabolic pathways in fungi that clustering confers a selective advantage to the cluster itself. The evolutionary pressure to maintain the cluster is proposed to be distinct from the pressure on the biosynthesis enzymes or the metabolites themselves. Clustering is also considered to favour the survival of special metabolites, because these genes depend, in part, on HGT for their dispersal and survival (Walton 2000). A mechanism favouring the survival of a gene in a cluster environment as opposed to an individual position in the genome, would over time lead to the accumulation and retention of the genes of a biochemical pathway in a cluster. During vertical transmission from one generation to the next, clustering would not be favoured, as the entire genome is passed to the progeny. Only in horizontal transmission would clustering provide an advantage to any given gene – because its function would be retained in a fully functional environment, the gene would survive. Movement of the entire pathway will then transfer a functioning module to the recipient, which, in one step, acquires a new trait that might provide a significant selective advantage. Thus, if the function is advantageous, the cluster would be useful, and it would be propagated and survive. Adaptive evolution is then only necessary to remove any adverse effects from the new products (Batada and Hurst 2007; Batada et al. 2007; Hurst et al. 2004; Keller et al. 2005; Walton 2000). All this is certainly true and applies to a number of gene clusters or parts thereof. But strong arguments tell against Walton’s selfish operon hypothesis encompassing all mechanisms relating to physical linkage groups and their transmission, for example the special metabolite biosynthesis pathways not organized in gene clusters or genomic islands. As we are certainly looking at evolution as an continuous and ongoing process, the appearance and fixation of the genetic features of special metabolism will be at a different step in the evolutionary process for each pathway considered. Several other authors addressed the topic and collected a number of mechanistic data relating to

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the accumulation and linkage of related genes. Pac and Hurst (2003) conclude that gene clusters can be formed by either selection for genetic linkage or for physical proximity. Integration of a given gene into a physically linked group, according to Firn and Jones (2009) will be advantageous and will occur more readily, when it codes for a metabolite function important enough to be retained for its specific properties supportive in a larger metabolic context. Gene clusters can accordingly be interpreted as small islands of basic integrated metabolism, or as groups of genes on their way there. Conservation of gene clusters extends even to the synteny level, indicating that selection can also act to preserve gene order. The aflatoxin/sterigmatocystin cluster in Aspergillus spp. has been preserved for at least 25 million years (Cary and Ehrlich 2006; Ehrlich et al. 2005). In A. fumigatus, 22 putative special metabolite gene clusters have been identified (Perrin et al. 2007), producing various substances, such as pigments, fumitremorigens, festuclavine, elymoclavine, fumigaclavines, ergot alkaloids, gliotoxin, and others. At least seven of them are located within 300 kb of a telomere. The telomere-proximal clusters, moreover, tend to contain more genes than clusters located more central within a chromosome. Thus, the accumulation of additional genes may be facilitated in this regions (Perrin et al. 2007) characterized by frequent chromosomal rearrangements (Galagan et al. 2005; Nierman et al. 2005). Subtelomeric regions of fungal chromosomes are highly variable, displaying a ubiquitous feature already evident in bacteria. The linear genome of Streptomyces is also organized into a central highly conserved core containing essential genes, and flanking arms with less conserved and more specific genes (Hsiao and Kirby 2008). Genes assumed to be involved in processes leading to niche specialization, accumulate in the neighbourhood of telomeres (Fairhead and Dujon 2006; Rehmeyer at al. 2006). In two strains of A. fumigatus with an overall diversity of 2%, the majority of these divergences were found in the subtelomeric regions and other synteny breakpoint regions. Many special metabolite clusters are either flanked by or contain embedded transposons or transposase-like elements (Niermann et al. 2005; Shaaban et al. 2010). These mobile elements may help to generate the observed diversity between the diverse strains or species. Rearrangements in

these regions may effect the production of special metabolites and have therefore significant influence on the niche adaptation between different strains or species. In the yeast Kluyveromyces lactis, a considerable percentage (30%) of all subtelomeric genes encode products predicted to be localized at either the cell surface of the plasma membrane, for example transporters and lectins (Fairhead and Dujon 2006). Detailed analysis of the evolutionary changes in this genomic region revealed that this location is indeed a hotspot where multiple gene duplication and recombination events superimposed each other, leading to unique combinations of similar genes. The region is still variable, following mechanisms, which additional to the duplication and recombination include template slippage or template jumping during DNA synthesis or repair events (Ricchetti et al. 2003). The accumulation of duplicate genes in this region is highly conspicuous, it has therefore been suggested that gene copies are transferred to this region, when, for whatever reason there might be, a duplicate is needed or when a particular transcriptional regulation is needed, as for example for the FLO genes of S. cerevisiae (Halme et al. 2004).

Studies on various model systems have shown other unusual properties of subtelomeric regions, including reversible silencing of genes mediated by proteins binding to the telomere, and engagement in ectopic recombination with other subtelomeres. A subtelomeric location seems generally to confer a capacity for gene diversification by diverse recombination mechanisms (Rehmeyer et al. 2006). Taking this into account, intragenomic reorganization and vertical transmission is also a conceivable explanation for the formation of a gene cluster. The idea of co-regulation as one of the major driving forces of gene clustering, on the other hand, is supported by the identification of several clusters of virulence genes not encoding any metabolite pathways, but putative secreted proteins in the genome of the corn smut fungus Ustilago maydis (Ka¨mper et al. 2006). Once a cluster has formed, it is under selective pressure as an entity and will certainly adapt to environmental challenges. This can also be deduced from the fate of another group of linked genes: The genes for the utilization of galactose (GAL genes) are clustered in all yeast species that possess them. But the GAL gene cluster is not present in four species that have lost the ability to use galactose. In three of these species, no trace of any GAL cluster gene is detectable any longer, indicating that the clustered genes got lost as


linked entity together. The fourth species still possesses all seven GAL genes, but only in the form of non-functional pseudogenes (Hittinger et al. 2004). Adaptation to a new lifestyle may cost the loss of metabolic pathways not longer under selective pressure, and a whole pathway may also get lost when it becomes disadvantageous under changed conditions. Clustering will facilitate such genome reorganizations. Both arguments are well in line with the evolution of the reduced metabolic capacity observed in many pathogens and endosymbionts. Clustering of genes may also display remarkable species-specific differences. Neurospora crassa was found to be comparatively rare in clustered genes. For example, the genes sAT and sCT needed for sulfur metabolism are clustered in two Aspergillus species (Borges-Walmsley et al. 1995), and the carotene biosynthesis genes carB and carRA are clustered in other species, e.g. in Gibberella fujikuroi (Linnemansto¨ns et al. 2002). In N. crassa, these genes are separated by 100 and 80 kb, respectively. This small distance is interpreted as evidence for the recent separation of the genes in N. crassa. Both small clusters seem to be disintegrating at the moment (Mannhaupt et al. 2003). No theories on cause and mechanisms behind this deviating attitude towards clustered genes in N. crassa have yet been formulated. 3. Horizontal Gene Transfer Phagocytosis as gene transfer mechanism is clearly to be ruled out for both fungi and oomycetes, but anastomosis between hyphae, transduction via mycoviruses or the propagation of retrotransposons are possible uptake mechanisms. Another pathway relays on the direct transfer from a bacterial donor. Agrobacterium tumefaciens, for example, can be induced to transfer its Ti plasmid to yeasts and filamentous fungi (Lacroix et al. 2006), and broad-host-range plasmids and phages may be transmitted via a number of other bacteria (Ragan and Beiko 2009). Bacterial endosymbionts might also introduce part of their genome into their fungal host. The Rhizopus microsporus/Burkholderia rhizoxinica/B. endofungorum endosymbiosis system (where the toxins rhizoxin and rhizonin, formerly ascribed to the fungus, are actually produced by the endosymbionts; Partida-Martinez et al. 2007a, b), might

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well represent a first step in the evolution of a toxin-producing fungus. It is highly probable that a similar lifestyle in the same environment provides the opportunity for large-scale gene or genome transfer between the various organismic groups (Richards et al. 2006). In the counter-argument, the high similarity of lifestyle might also be the result of genetic exchange. Generally, judging from the number of published examples, HGT seems to be far more abundant between microorganisms than between micro-organisms and multicellular eukaryotes (Andersson 2009). The transferred and subsequently retained genes would enable the use of previously non-useable substrates or provide other selective advantages. An example for such a process is cutinase, an enzyme required for the degradation of cutin and essential for plant pathogens (Belbahri et al. 2008). Cutin is an insoluble lipid polyester and a major cell wall component of plants. Cutinase exists only in fungi, oomycetes and actinobacteria, but is most probably of more recent origin than the splitting of these three lineages. Belbahri and co-workers (2008) propose that the gene originated in actinobacteria living in the vicinity of the emerging land plants and was transferred from bacteria to eukaryotes sharing the same habitat.

Transfer of genetic material might extent from single genes to complete chromosomes. Akagi et al. (2009) describe the transfer of a dispensable chromosome conveying plant pathogenicity between strains of Alternaria alternata. A study of the evolutionary history of all predicted 11109 genes of Magnaporthe grisea (Richards et al. 2006) showed that 11 of these genes are probably the result of a HGT event between distinct eukaryotic lineages. For four genes, the transfer from fungi to oophytes was confirmed using stringent phylogenetic analysis methods. The four strongly supported genes are a sugar transporter from the multifacilitator superfamily (AraJ), a putative broad-specificity permease possibly involved in nucleotide uptake (CodB), a protocatechuate 3,4-dioxygenase and a member of the GalM gene family encoding aldose-1-epimerases, also exhibiting broad substrate specificity. All genes originate from the ascomycete lineage within the fungi, and three of them entered the fungal lineage as result of lateral gene transfer from prokaryotes around the time of the diversification between Zygomycota and Dikaryomycota. Some less well supported putative HGT events include putative esterase/lipases and aconitases.

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The same group also presented data supporting bidirectional HGT between plants an fungi (Richards et al. 2009). The genes transferred and retained by the respective recipients are supposed to provide advantages for adaptation and competitiveness in the soil surrounding: they were identified as putative solute transporters for sugars or other small molecules, a siderophore biosynthesis protein involved in iron uptake, and an alcohol dehydrogenase. For some of these HGT events, data analysis suggests a two-step transfer indicating a prokaryote as the original donor or transmitter of the transferred gene. The authors maintain that plant–fungal HGT events must have been extremely rare and ancient. They only involve basal plant lineages prior to the diversification between bryophytes, lycophytes, and angiosperms, and fungal lineages prior to the radiation of the ascomycetes (Richards et al. 2009). The best documented HGT event between fungi is the transfer of a toxin gene ToxA from Stagnospora nodorum to Pyrenophora triticirepentis, only 70 years ago, leading to a new disease of wheat (Friesen et al. 2006; see Chapter 9 in this volume). HGT has also been proposed for the pea pathogenicity cluster in Nectria haematococca mating population VI. This cluster is responsible for the detoxification of the plant phytoalexin, pisatin (Temporini and VanEtten 2004). It is located on a dispensable chromosome, the genes do not have paralogues anywhere else in the genome, and GC content and codon usage differs significantly from the flanking regions and the other chromosomes, all strongly suggesting an origin from outside the species. This cluster also shows a disjunct distribution; it is present in Fusarium oxysporum, but not in the closely related Neocosmospora boniensis (Temporini and VanEtten 2004).

For fungi, few cases for the acquisition of metabolic gene clusters by HGT are supported by experimental data, for example for the origin of a nitrate assimilation gene cluster (Slot and Hibbett 2007), a cluster containing the genes for galactose utilization (Slot and Rokas 2010), and also the T toxin biosynthesis gene cluster of Cochliobolus heterostrophus (Yang et al. 1996), the HC toxin gene cluster of C. carbonum (Ahn and Walton 1996), and the AK toxin gene cluster in A. alternata (Tanaka et al. 1999). The toxin gene clusters are present in only some isolates of a species and are missing in others with an otherwise nearly isogenic genetic background.

A similar picture was found for a gene coding for a trichothecene 3-O-acetyltransferase several close and distantly related ascomycete species (Tokai et al. 2005). In some of the species and isolates, the gene was inactive and non-functional. Nevertheless, its distribution strictly follows the general species phylogeny. The genomic regions surrounding the gene in non-trichothecene-producing strains exhibit no synteny with regions inside the trichothecene gene cluster in trichothecene-producing strains. Tokai and co-workers (2005) propose that this gene has a different evolutionary origin from other trichothecene biosynthesis genes located within the cluster. The persistence of this non-essential gene, and also in most cases of a functional copy, is explained by an selective advantage it might provide for the species in which it exists, although its exact role in these fungi is not known. The discontinuous distribution of gene clusters has been proposed to be caused by several mechanisms: Kroken et al. (2003) proposed a sequence of gene duplication, divergence and gene loss as more probable for the evolution of type I polyketide synthetases than HGT, stating that selective pressure on the function of the compound would also allow for the observed results in many cases. Fungi produce a number of substances that interfere with plant signalling and may thus play a role in environmental fungal–plant interactions. In all cases, the previously suggested probability of acquisition by HGT from the plants to the fungus was discounted as the biosynthesis of the respective compounds shows the distinctive differences between the two lineages. For ethylene, at least two pathways different from those in plants exist in fungi (Chague et al. 2002; Hottiger and Boller 1991). With gibberellins, the situation is somewhat more complex. The enzymes catalysing the individual steps differ considerably while the reaction series in the biosynthesis pathway itself does not deviate very much (Hedden et al. 2002). For abscisic acid, too, biosynthesis pathways completely different from those in plants have been identified in Botryotinia fuckeliana (anamorph: Botrytis cinerea) and Mycosphaerella cruenta (anamorph: Cercospora cruenta; Oritani and Yamashita 1985; Yamamoto et al. 2000a, b; for a review, see Oritani and Kiyota 2003). This might well be a case of the selective force being directed


on the function of the product, as abscisic acid is proposed not only to serve in interspecies communication of plant–pathogenic fungi but also to play a role in the regulation of spore production and germination as well (Marumo et al. 1982). An abscisic acid biosynthesis gene cluster in B. fuckeliana contains two cytochrome P450 monooxygenase genes, the second one with similarities to trichothecene C-15 hydroxylases from some Fusarium species (Siewers et al. 2006). Downstream of these two genes and transcribed in the same direction is a putative short-chain dehydrogenase. The cluster is completed by a gene with similarities to fungal pectin lyase genes. The organization of this cluster is conserved in three strains. The four genes are not strictly co-regulated but are all induced in abscisic acidproducing conditions (Siewers et al. 2006).

4. Adaptive Evolution Strain-specific differences in the metabolite pattern for trichothecene mycotoxins (chemotypes) do not correlate well with a phylogeny of the complete Fusarium graminearum strain complex, based on a six-gene-based phylogenetic tree using single-copy nuclear gene sequences. Polymorphism within the trichothecene gene cluster is trans-specific and has apparently been maintained by balancing selection acting on the chemotype differences that occurred already in the common ancestor of this group. Ancestral polymorphism is supposed to have been followed by adaptive evolution. Each chemotype from the B-trichothecene lineage can be traced back to a single evolutionary origin in the ancestor of this lineage and this polymorphism persists through the multiple subsequent speciation events. The polyphyletic distribution of the trichothecene phenotypes relative to the species phylogeny is explained as the result of non-phylogenetic sorting of an ancestral polymorphism into the descendant species. Balancing selection which provides an advantage for a heterozygote over both homozygotes will keep several alleles of a gene in a population. This form of selection is supposed to have directly acted upon the chemotype differences (Ward et al. 2002). Strains of Fusarium compactum also display genotype variations which determine the metabolite spectrum (Talbot et al. 1996).

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5. Regulation of Special Metabolite Biosynthesis Regulation of special metabolite biosynthesis, as modulated by adaptive evolution, has resulted in some features distinct from other metabolic requirements. As a general feature, enzymes catalysing special metabolite biosynthesis often occur in rather low concentrations and lack feedback regulation (Hartmann 2007). Entry into the biosynthesis pathways is also strictly controlled relating to developmental stage and location within the organism. Many special metabolites are produced only in neglectable amounts or not at all by the undifferentiated, young mycelium. Only the more mature stages are able to commence and maintain high production rates. This can be understood as the consequence of regulation by a network receiving inputs from many other pathways and possibly of external triggers, and integrating these signals. The deduced evolutionary advantage is seen in minimized energy costs or saved regulation and coordinating costs. Any metabolic system producing substances that are not actually necessary or needed at a given time will accumulate products that in turn may influence homoeostasis adversely and may also invite the accumulation of process errors. The production of many special metabolites, especially chemical communication signals, such as pheromones or symbiont metabolites, is only triggered by specific events in the interplay between individuals of the interacting species. The environmental stimuli or internal triggers often induce globally acting transcription factors like LaeA, PacC, or VeA which are regulating multiple pathways and responses (Perrin et al. 2007). Regulation of fungal special metabolism is influenced by the chromosomal arrangement of the genes. As a reason for the clustering it is assumed that this enhances the efficiency of gene regulation. Generally, all genes within a cluster are co-regulated, not only the biosynthetic, but also the regulatory genes themselves. Thirteen of the 22 putative special metabolite clusters of A. fumigatus were found to be regulated by LaeA (Perrin et al. 2007). The regions controlled by this factor are species- or strainspecific, which indicates that they may serve as niche adaptation factors (Bok and Keller 2004). Loss of the LaeA gene leads to a markedly diminished repertoire of special metabolites (Perrin et al. 2007) and this apparently has a negative impact on A. fumigatus pathogenicity.

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When gene clusters are maintained through co-regulation of their component genes, the coregulation is likely to occur at the chromatin level. Here is the connection to the observed effect of histone deacetylases on the control of special metabolite production, and also to the histone methyltransferase LaeA. LaeA preferentially affects expression of genes retained within a cluster; transfer of genes into or out of the cluster results in gain or loss of transcription regulation by LaeA (Bok et al. 2006; Perrin et al. 2007). Keller and co-workers (2005) suggest the regulation of nucleosome positioning and heterochromatin formation as the relevant events in this regulation process. Histone deacetylases do repress the production of several metabolites in A. alternata, Penicillium expansum, and A. nidulans. This may be rated again as support for the idea that the localization of special metabolite biosynthesis gene clusters in subtelomeric regions is meaningful and has an impact on their susceptibility to recombination and epigenetic regulation (Shwab et al. 2007). The gene coding for a transcription factor regulating expression of the genes within a special metabolite cluster is usually also embedded in that cluster. These transcription factors may also act on genes elsewhere in the genome, underlining their global nature. Besides the special metabolite gene clusters, LaeA also regulates eight more groups of linked genes (Perrin et al. 2007). Aflatoxin (AF) and sterigmatocystin (ST) are related compounds, derived from a similar gene cluster. AflR is a Zn(II)2Cys6 zinc binuclear protein that activates transcription by binding to palindromic DNA sequences in the promoter of AF/ST cluster genes (Ehrlich et al. 1999; Fernandes et al. 1998; Payne and Brown 1998). It regulates the aflatoxin clusters of A. flavus and A. parasiticus and the sterigmatocystin cluster of A. nidulans (Brown et al. 1996; Chang et al. 1993; Fernandes et al. 1998; Woloshuk et al. 1994; Yu et al. 1996) and also three genes outside the cluster (Price et al. 2006). Aspergillus sojae does not produce aflatoxin mainly because of a point mutation in the aflR gene causing the truncation of the transcription activation domain of the transcription factor and thus the interaction with the transcription co-activator AflJ (Chang et al. 2007). Zinc binuclear transcription factors are only known from fungi, where they constitute the most common type of incluster regulators. The GliZ transcription factor regulating

the gliotoxin biosynthesis cluster (Bok et al. 2006), and MclR regulating synthesis of the statin compactin in Penicillium citrinum (Abe et al. 2002) also belong in this protein family. For a review on special metabolite biosynthesis regulation, see Shwab and Keller (2008).

Other clusters do not contain any regulatory genes, among them the ergovaline and lolitrem clusters in Neotyphodium/Epichloe¨ (Fleetwood et al. 2007; Young et al. 2001, 2005, 2006; Zhang et al. 2009). Expression of these pathways probably relies on their induction by plant factors, as the expression level is only high for in planta situations (Fleetwood et al. 2007; Young et al. 2005). Transcriptional regulation is the integration point for all stimuli and signals relevant to the induction of a metabolic pathway. In Achremonium chrysogenum, the promoter sequences of all cephalosporin biosynthesis genes contain potential binding sites for several transcription factors, namely the zinc finger proteins PACC and CRE1 already known from other fungal blactam producers, and the RFX transcription factor CPCR1 initially discovered in A. chrysogenum (Schmitt et al. 2004). In addition, VeA regulates development, that is, sexual development in A. nidulans, light responses in P. blakesleeanus, hyphal fragmentation in A. chrysogenum, but also special metabolism in Aspergillus spp. and A. chrysogenum, probably by modulating the expression of other transcription factors (Bayram et al. 2008; Calvo 2008; Dreyer et al. 2007).

E. Gene Clusters for Specific Metabolic Pathways 1. Metabolic Gene Clusters in Yeast Comparison of the sequence data of the first fully sequenced genome of a filamentous fungus, N. crassa, with those obtained from S. cerevisiae, show that the yeast genome has been streamlined during evolution and many genes have been lost (Mannhaupt et al. 2003). In the filamentous fungus, a high proportion of the deduced proteins is involved in metabolism, cellular communication, and signal transduction, and it is needed to retain the full capacity for adaptation to changing environmental conditions. Filamentous fungi may utilize many different substrates and must be able to cope with any number of toxic or detrimental compounds. The highest difference between the


two genomes was observed in the category ‘secondary metabolism’, with only seven entries for S. cerevisiae as opposed to 23 in N. crassa (Mannhaupt et al. 2003). In comparison to filamentous fungi, clustering of genes is a much more rare observation in S. cerevisiae. Nevertheless a number of pathways, especially for specialized metabolism, are also organized in clusters in this unicellular fungus. Among them are pathways for the utilization of some carbon sources (e.g., galactose), allantoin as nitrogen source (Wong and Wolfe 2005), biotin synthesis, vitamin metabolism (Hall and Dietrich 2007; Tanaka et al. 2005b) and resistance to arsenic (Bobrowicz et al. 1997). In S. cerevisiae, neighbouring genes tend to be co-expressed (Cohen et al. 2000) and the spatial linkage of co-expressed genes tends to be conserved during genome reorganization events (Hurst et al. 2002). For the allantoin usage cluster DAL, the events leading to the present day situation have been studied in some detail (Wong and Wolfe 2005). This was facilitated by the fact that the genesis of the DAL cluster is a rather recent development and is not yet completed, so that different situations concerning the localization of the homologous genes are found in a number of yeast species. The cluster contains the three genes required for the breakdown of allantoin to glyoxylate and urea, plus genes coding for allantoin racemase, allantoin permease, and a malate synthase whose function is apparently to dispose of glyoxylate and thus prevent product inhibition of the catalytic process (Cooper 1996).

Several functionally related genes are not clustered with those already mentioned: an allantoate permease gene, a gene encoding an enzyme involved in the degradation of urea to ammonia, and several transcription factor genes (Wong and Wolfe 2005). The six genes are located consecutively near a telomere of chromosome IX. The cluster is only found in a monophyletic group comprising S. cerevisiae and some closely related species, and only in this group is the cluster complete and located at the same position. In S. castelli, the DAL genes are clustered, but with two differences in the internal arrangement and at a somewhat different position in the genome. In the more distant hemiascomycetes, the component genes of the cluster are dispersed individually. Four of the component genes are single-copy

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genes in S. cerevisiae and seem to have transposed to their new position, possibly after gene duplication and subsequent loss of the genes at the original positions. At the same time, the purine degradation pathway has undergone a biochemical reorganization, switching to allantoin instead of urate as the primary substrate. This was possibly a result of the adaptation to an more anaerobic life style and the concurrent loss of several oxygenconsuming enzymes, among them urate oxidase (Wong and Wolfe 2005). Thus a completely new gene cluster has arisen, with a function previously not existing in Saccharomyces. The species without a DAL cluster are still able to metabolize urate, but are limited to aerobic environments. The recombination and re-functionalization of the purine degradation pathway to a higher tolerance against anaerobiosis therefore provides a significant selective advantage. Wong and Wolfe (2005) saw a selective pressure directed at the loss of urate. The metabolic adaptation to a specialized niche is considered a selective force large enough to drive the genome reorganization, and the epistatic selection for the retained linkage of the reorganized genes is supposed to prevent rapid loss of the linkage. An additional factor influencing the stability of the genetic linkage is the need to select for a pathway that minimizes the accumulation of potentially harmful intermediates. In the DAL pathway, the product glyoxal is toxic to yeasts, but it is removed by another product of the same gene cluster, thus strengthening the linkage between the two genes, DAL3 and DAL7. Metabolic channelling through this route is indicated by the observation that Dal3 enzyme activity is reduced when DAL7 is dysfunctional. During reorganization of the purine degradation pathway, two new genes, DAL4 and DAL7 arose from gene duplication. While these new genes were adapting to their new functions, they may have interacted well with only specific alleles at the other DAL loci. Under epistatic selection, the organization of these alleles into a cluster would have been favoured (Wong and Wolfe 2005).

Physical clustering may have been favoured by the location of the DAL cluster. Regions near the telomeres are known to contain numerous genes for the exploitation of unusual nutrients and whose transcription is induced by the respective environmental stress signal. Subtelomeric regions are also a hot-spot for chromatin rearrangement events (Robyr et al. 2002). Selective pressure seems to have acted on the clustering

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itself, not on the sequences of the component genes of the DAL cluster (Wong and Wolfe 2005). In another study, a complex evolutionary process was postulated to explain the distribution and the organization of gene clusters in S. cerevisiae (Hall and Dietrich 2007). Gene clusters in this species contain only few categories of genes, among them such that encode dispensable pathways. One of these, the biotin biosynthesis pathway was apparently lost in an ancestral species and re-acquired again by a combination of HGT of a single gene from a bacterial source and subsequent gene duplication with neofunctionalization of the copy and organization of the pathway into a linked group. Hall and Dietrich (2007) suggested this complex procedure as a way to avoid the effects of a potentially toxic metabolic intermediate. If a pathway confers a selection advantage only in some situations, but strong adversary effects in others, the positive and negative selective forces are both strong and counteracting. As a consequence, the linkage of the pathway genes in a physical cluster might be advantageous, because in the cluster it is less probable that only a part of the clustered genes will be expressed while a possibly existing detoxifying gene remains silent. If such a pathway was lost during evolution, selective pressure would occasionally allow re-acquisition of the pathway by HGT (Hall and Dietrich 2007). Patron et al. (2008) and Kahldi et al. (2008) expand this theory on other ascomycetes. In several lineages, the data also support the frequent loss of ancient gene clusters followed by occasional reacquisition via HGT as an explanation for the recent disjunct distribution of these clusters. 2. Biosynthesis of b-Lactam Antibiotics b-Lactam antibiotics are produced by bacteria and fungi. The hydrophobic penicillins are produced solely by ascomycete fungi, whereas the water-soluble cephalosporins occur in both bacteria and fungi. The early reactions in biosynthesis of the two types of compound are identical (see The Mycota, vol II). At the beginning, the three amino acids L-a-aminoadipic acid, L-cysteine, and L-valine are condensed to the tripeptide ACV by d-(L-a-aminoadipyl)-L-cysteine-D-valine synthetase encoded by the gene acvA in A. nidulans and pcbAB in Penicillium chrysogenum and A. chrysogenum.

In the second step, isopenicillin is formed as a result of the oxidative ring closure of the tripeptide, leading to the bicyclic b-lactam-thiazolidine ring system typical for both antibiotic families. This step is catalysed by isopenicillin N synthetase coded for by the gene ipnA/pcbC. Penicillins are formed from this intermediate when the hydrophilic L- a-AAA side chain is replaced by a hydrophobic acyl group in a reaction catalysed by the acyl coenzyme A-isopenicillin N acyltransferase, encoded by aatA/penDE. In the synthesis of cephalosporins, penicillin N is formed as the L-a-AAA side chain is isomerised to the D-enantiomer. Next, the thiazolidine ring of isopenicillin N is extended to a dihydrothiazine ring by action of deacetoxycephalosporin synthetase, which also catalyses the hydroxylation step leading to the formation of deacetylcephalosporin. Cephalosporin C is finally formed when an acetyl group is transferred to the deacetoxycephalosporin hydroxyl group by acetyl coenzyme A – deacetoxycephalosporin acetyltransferase, a product of the cefG gene in A. chrysogenum (for a review, see Brakhage et al. 2005).

The high similarities between the bacterial and fungal biosynthesis pathways triggered for the first time a discussion on the possible bacterial ancestry of b-lactam biosynthesis and the acquisition of this trait by HGT in fungi. The organization of the biosynthesis genes and the lack of introns in some of them as well as the high GC content in codon third positions was seen as supporting this hypothesis (e.g., Aharonowitz et al. 1992; Penalva et al. 1990), as well as the predominant location of regulatory genes outside the cluster (Bok and Keller 2004; Brakhage et al. 2004). Overall, regulation of the whole pathway is mediated by rather global regulators, leading to the assumption that these took over the regulation of this biosynthesis pathway at a later stage. Arguments against the transfer of the functional linkage group include the fact that, although clustered, in fungi the genes are transcribed separately from individual promoters, and not as polycistronic messages as in bacteria – therefore, the most obvious argument for cluster formation in bacteria is invalidated. Gene linkage is supposed to reflect the origin of the genes here; it is supposed to be derived from the bacterial ancestor. But, encoding acyl coenzyme A-isopenicillin N acyltransferase, an integral component of eukaryotic b-lactam biosynthesis, is not part of the bacterial gene cluster and, moreover, has no close relative in recent prokaryotes. Recent hypotheses (Brakhage et al. 2005, 2009) endeavour to incorporate all known facts about cluster structure and cluster-forming mechanisms. In more detail, the ipnA genes of


fungi and bacteria share a considerable number of identical positions: higher than 60% at the nucleotide level and still 50% at the amino acid level, indicating a common origin. Both the acvA and ipnA genes are organized in a gene cluster in Penicillium chrysogenum and in A. chrysogenum, supposedly reflecting the original prokaryotic situation. After transfer to the eukaryotic lineage, the cluster split, which explains the presence of two cephalosporin biosynthesis gene clusters localized on two chromosomes in A. chrysogenum. The part of the cluster containing the genes coding for the late enzymes of the cephalosporin biosynthesis pathway were lost in the lineage leading to A. nidulans and P. chrysogenum while the aatA gene, having no obvious homologue in bacteria, was recruited to the cluster only during its evolution within the fungal lineage. Activity of the aatA gene product leads to the formation of a product with considerably higher antibiotic activity than the end product of the pathway represented by the originally clustered genes, isopenicillin N. This provides an idea of the selective advantage and the selective pressure on the relocation of aatA into the cluster and its continued retainment. The discussion gained new momentum from some more recent studies on the regulation of blactam synthesis, which showed that none of the regulatory factors, themselves organized in a complex regulatory network, is limited in its action to the b-lactam genes only. All of them are active towards other genes, too. These regulators are typically eukaryotic, basic region helix-loop-helix proteins, CCAAT binding complex proteins or zinc finger proteins. Nothing indicates the presence anywhere of a prokaryotic type regulator associated with the b-lactam synthesis cluster. The major function of the wide domain regulator PacC active on ipnA (Tilburn et al. 1995) and probably acvA (Then Bergh and Brakhage 1998) is pH regulation, as well as AnCF, which also acts upon ipnA and aatA (Litzka et al. 1996, 1998; Steidl et al. 1999; for a review, see Brakhage et al. 2005). b-Lactam synthesis in A. nidulans is also influenced by a number of other transcription factors, such as VeA, a light-dependent regulator of a number of developmental reactions (Calvo et al. 2004; Kato et al. 2003; Spro¨te and Brakhage 2007), and AnBH1, a repressor of the aatA gene (Caruso et al. 2002), and also in A. chrysogenum CreA/Cre1, a protein known for its function controlling carbon source utilization (Jekosch and Ku¨ck 2000). In A. nidulans NRE/AreA, a global regulator of nitrogen metabolism, binds to sequence motifs in the promoter regions of several peniccillin biosynthesis genes (Haas and Marzluf 1995), but its proposed participation in the regulation of penicillin biosynthesis gene transcription

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has not been confirmed in any subsequent publication. The carbon catabolite regulation system also effects cephalosporin biosynthesis. Generally, it was concluded, that cephalosporin synthesis is regulated by a complex interacting or even functionally redundant transcription factor network (Schmitt et al. 2004).

The resulting theory is that upon acquisition of the b-lactam synthesis genes by HGT, existing regulator genes or proteins were recruited to regulate this synthesis pathway. Probably a complete functional cluster was transferred and its bacterial regulator subsequently lost, but it also possible that the regulator gene was already lacking during the original transfer. This last hypothesis is strengthened by the observation that existing transcription factors take over regulation of the b-lactam synthesis cluster when it is transformed into non-producing fungi. N. crassa and A. niger have thus been induced to produce penicillin V (Smith et al. 1990). Recently, also the evolutionary history of the aatA gene has been better resolved by the work of Spro¨te and collaborators (2008). Those authors found that A. nidulans aatA deletion mutants still produce a small amount of penicillin, and they subsequently identified the gene aatB as responsible for this production. The two genes are supposed to be paralogues derived from duplication of an ancestral gene, based on their similarities both concerning sequence and organization. aatA was then recruited to the b-lactam synthesis cluster while aatB remained in its original environment. This story fits well with the observed pattern of recruiting new genes into groups of functionally linked genes. The paralogy of the two genes is further evident in their similar regulation pattern. Whereas the wide distribution of aatB within the ascomycetes is consistent with the phylogeny of this group, aatA-like genes are only found in the penicillin-producing taxa and only as part of the penicillin gene cluster. Because of a sequence identity around 80% between all aatA genes, a second HGT event distributing the complete cluster after the linkage of aatA to the genes originally acquired from bacteria is probable, but cannot be proven unequivocally at the moment. The production of the hydrophobic penicillins was further enhanced by the acquisition of the peroxisome targeting signal PTS1 to aatA, as the last step of the penicillin biosynthesis seems to be more effective in the peroxisome.

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3. Biosynthesis of Epipolythiodioxopiperazine Toxins Epipolythiodioxopiperazines (ETP) are fungal toxins derived from cyclic peptides. They bind to target proteins via cysteine residues after cleaving an internal disulfide bridge, the toxic effect stems from the reactive oxygen species generated in the process. Clustered genes coding for the biosynthesis of the ETP sirodesmin PL were described by Gardiner et al. (2004) for Leptosphaeria maculans and additionally identified, at least partially, in Chaetomium globosum, Magnaporthe grisea, Fusarium graminearum and Aspergillus fumigatus, where the biosynthesis product is gliotoxin, another ETP compound. When compared, the clusters in L. maculans and A. fumigatus (Gardiner and Howlett 2005) contain partially different genes. Those present in both clusters are considered to be ‘common ETP moiety’ genes whose function was deduced from comparison with known genes with similar functions combined with information on the chemistry of ETP synthesis. Among the genes differing in the two clusters are those responsible for the export of the toxin, this efflux is mediated by two different mechanisms in the two species. This supports the notion of cluster stabilization under the pressure of avoiding adverse effects from the metabolites produced. There is no compelling need for this task to be performed in the same way in the various species as long as accumulation of potentially harmful compounds is avoided. Putative ETP gene clusters with some or all of the common ETP moiety genes have been meanwhile found in 14 ascomycete taxa (Fox and Howlett 2008). In other species, namely the ascomycete yeasts Saccharomyces cerevisiae and Schizosaccharomyces pombe, and the dimorphic basidiomycetes Ustilago maydis and Cryptococcus neoformans, they are missing completely (Patron et al. 2007). It is not yet known whether any of the newly identified ETP cluster-containing species really produce ETP metabolites, or whether the lack of one or other genes has any effect on their production. Based on phylogenetic analyses on six of the cluster genes, most of them are of monophyletic origin, and moreover most closely related to nonclustered paralogues throughout the filamentous ascomycetes. All the clusters appear to have derived from a common origin. This suggests that originally dispersed genes were assembled

into a cluster, which from that time onwards has been transmitted relatively intact instead of being newly assembled in the different species. During evolution, the original cluster has developed into a number of distinct phylogenetic classes, but each showing a consistent pattern of subclade relationships, thus also supporting the hypothesis of a common origin of the cluster. As the cluster is distributed rather discontinuously within the ascomycetes, it might have been lost independently in many lineages, but transfer between the lineages by HGT cannot be excluded (Patron et al. 2007). Two of the ETP clusters, nevertheless, seem to be of independent origin: those from Gibberella zeae/Fusarium graminearum and Chaetomium globosum. These differ from the others in several features, among them the number of genes contained in the cluster. The genes recruited for the cluster probably were the results of gene duplication events and being selected into the cluster for the biochemical conformity of the gene products with the ETP pathway. This would explain why, in three closely related Aspergillus species, the common ETP moiety gene encoding dipeptidase J is replaced by another paralogue or why the A. terreus subcluster lacks a gene which is present in the A. fumigatus and N. fischeri subclusters. The phylogeny of the peptide synthetase within the cluster is also somewhat complex. A duplication of the ancestral gene apparently occurred shortly after its recruitment to the cluster but before the various lineages split. The phylogeny of the various ETP clusters differs from that of their harbouring organisms with subclade III being found in Trichoderma virens (Sordariomycetes) and L. maculans (Dothidiomycetes), but not in Phaeosphaeria nodorum (Dothidiomycetes). In the Eurotiomycetes, and the Sordariomycetes, different subclades are distributed between the species (Patron et al. 2007). Analysis of the phylogenetic relationships allows two possible patterns of inheritance. Vertical inheritance and cluster loss as inheritance pattern seem highly improbable: this would infer at least double duplication of the whole cluster to give rise to all the subclades and subsequent multiple incidents of independent cluster loss, estimated to be at least 17 independent incidents, but very likely much more. The scenario has no explanation for the high conservation ratio within some of the subclades despite their presumed origin from an ancestor to the ascomycetes. The second scenario includes HGT


and appears much more likely. It states that the gene cluster evolved over time to the to the phylogenetic relations seen today between the subclades. Horizontal transfer of complete clusters account for the complex distribution of the subclades found today. This scenario infers three HGT events and only five incidents of cluster loss and seems therefore to be more realistic (Patron et al. 2007). Very interesting is the question of inheritance of these special metabolite gene clusters, as their distribution is disconnected. This has also been observed in other filamentous fungi. Dothistroma septosporum contains a number of genes strongly similar to genes of the sterigmatocystin and aflatoxin gene clusters of Aspergillus. These genes are mostly co-regulated, but organized in three ‘miniclusters’ containing also some genes not found in the respective, tightly linked 25-gene Aspergillus cluster. The pathway is functional and produces dothistromin, a toxin with a similar structure to an intermediate in the sterigmatocystin and aflatoxin biosynthesis (Zhang et al. 2007). 4. Gibberellin Biosynthesis Gibberellins are a large group of compounds, comprising currently at least 163 metabolites. Despite their being generally announced as phytohormones, metabolites belonging into this group are produced by bacteria and fungi in addition to plants of all developmental levels. In plants, they are involved in growth stimulation and regulation of developmental processes throughout the life cycle (for a review, see Ferguson and Mathesius 2003). The presence of these compounds in bacteria and fungi has hitherto been ascribed as an adaptation towards a plant pathogenic or mutualistic lifestyle (Rademacher 1994). Not unequivocally proven and only to a minor degree, positive effects similar to those in plants have also been described for N. crassa, G. fujikuroi and P. chrysogenum, whereas no adverse effects of gibberellins were apparent in cultures of G. fujikuroi and Sphaceloma manihoticola cultivated in the presence of gibberellin biosynthesis inhibitors. At present, the true function(s) of gibberellins in fungi are apparently not yet fully understood, and it can by no means be excluded that they may mirror to some extent those in plants.

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As the biosynthesis pathways nevertheless show marked differences between fungi and plants, on the chemical as well as the enzymatic and the genetic level, gibberellin biosynthesis must have evolved independently over a long period in these two organismic groups. No data supporting the earlier proposal (Chapman and Regan 1980) of acquisition of the gibberellin synthesis cluster by HGT from a plant ancestor to fungi have ever been found (Bo¨mke and Tudzynski 2009). Gibberellins are a large family of diterpenoids, all based on a tetracyclic ent-gibberellane skeleton. The compounds contain 19 to 20 carbon atoms forming either 4 or 5 rings, the fifth, variable, ring always being a lactone. While not all of the C19 gibberellins are bioactive, all bioactive gibberellins contain 19 carbon atoms. In fungi, the synthesis starts from acetate and proceeds via the mevalonate pathway and the basic isoprenoid isopentenyl diphosphate. Gibberellin synthesis branches off the general terpenoid pathway from farnesyl diphosphate onwards, with an additional geranylgeranyldiphosphate synthase (GGS2) exclusively used for geranylgeranyldiphosphate entering into gibberellin biosynthesis (Tudzynski and Ho¨lter 1998). In plants, at least in their green parts, isopentenyl diphosphate is also introduced from the methylerythritol phosphate pathway located in the plastids, where it is synthesized from pyruvate via glyceraldehyde 3-phosphate. The following steps leading from geranylgeranyldiphosphate to GA12-aldehyde are similar, but the pathway splits again in the respective final steps, leading to the major products GA4 and GA1 in plants versus GA3 in the ascomycete Gibberella fujikuroi. The GA1 formed in this fungus from GA4 is only a minor product and is not converted into GA3.

One fundamental feature of fungal gibberellin synthesis is the organization of the relevant genes, including the GGS2 encoding gene, into a gene cluster. This fact plus the differences generally observed in localization and reaction mechanism of gibberellin biosynthesis (Kawaide 2006; Malonek et al. 2005a–c) encouraged Tudzynski and coworkers to propose an independent evolution of the pathways in plants and fungi. Of specific interest in the context of this review is the question of gibberellin synthesis evolution within fungi. This was studied in detail by the group of B. Tudzynski (for a review, see Tudzynski and Bo¨mke 2009) for the G. fujikuroi species complex. This clade comprises a number of plant pathogens with marked host specificity. The monophyletic group can by subdivided into several mating populations which differ significantly in their production profiles for

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special metabolites. The gibberellin gene cluster occurs in several mating populations, but most of the individual species have either lost or never possessed the ability for gibberellin production, despite their retaining the complete gene cluster (Malonek et al. 2005c). Comparison of the gibberellin-producing G. fujikuroi with its close relative, a non-producing strain of Fusarium proliferatum showed an identical organization of the gene cluster with an overall sequence similarity of 94%, but with weak or missing transcription of the genes in the non-producing strain. This was attributed to several mutations in the coding and the 5’ noncoding regions of the two cluster genes ggs2 and cps/ks, coding for the pathway-specific GGS2 and a bifunctional terpene cyclase, respectively. In the presence of a functional ggs2 gene, gibberellin production is restored in F. proliferatum (Malonek et al. 2005a; Rim et al. 2005). In other members of the species complex, the gene cluster is further degraded. In F. verticilloides, only two of the seven cluster genes are still present, and only one of the remaining genes is still functional (Bo¨mke et al. 2008a, b). Nevertheless, the regulation network for gibberellin biosynthesis gene expression, including induction of gene expression by nitrogen limitation via the transcription factor AreA, is still functional. Several modifications in the region surrounding the gene cluster indicate the influence of gene rearrangement events in the inactivation, together with the integration of outside sequence elements. These events led to the disintegration of the gibberellin synthesis gene cluster (Bo¨mke et al. 2008b). In Elsinoe brasiliensis (anamorph: Sphaceloma manihoticola), a gibberellin-producing cassava pathogen exhibiting the same pathogenic effects as G. fujikuroi despite its systematic classification in a different order, the functional gene cluster consists of only five genes (Bo¨mke et al. 2008a). Based on sequence comparison and functional characterization, these genes are seen as orthologues to the G. fujikuroi gibberellin biosynthesis genes. The similarity extends to the cluster organization. The genes cps/ks and ggs2 as well as the gene pair SmP450-1 and SM-P450-4 are physically linked in both species, each gene pair is linked by a bidirectional promoter and thus forms a transcriptional unit (Bo¨mke et al. 2008a). Besides the absence of two genes from the G. fujikuroi gene cluster, two sequence inversions also mark the differences between the two species. Absence of

synteny in the regions surrounding the gibberellin gene cluster suggests lateral transmission as a possible mode of acquisition for this gene cluster, but the direction of the putative transfer cannot be inferred from the available data (Bo¨mke and Tudzynski 2009; Bo¨mke et al. 2008a). Several other species from the G. fujikuroi species complex contain functional gibberellin gene clusters, some more are known to produce gibberellins but still do not elicit any pathogenic effects in their host plants (Bo¨mke and Tudzynski 2009). Outside the species complex, Phaeospheria sp. strain L487 exhibits a GA profile different from G. fujikuroi and Sphaceloma manihoticola, and the cluster shows a different organization, too (Kawaide 2006). Although three of the genes and gene products are similar to the G. fujikuroi P450-1, P450-2 and P450-4, an additional P450 gene codes for a plant-like C3-oxidase, shifting the gibberellin profile towards more plant-like products. The cluster also lacks a homologue of the pathway-specific geranylgeranyldiphosphate synthase (Kawaide et al. 2006). These fungal enzymes are nevertheless seen as true orthologues to the G. fujikuroi enzymes, based on the higher sequence similarity between the fungal enzymes compared to that between Phaeospheria and plant enzymes (Kawaide et al. 2006).

The similarities in the gibberellin gene cluster organization makes a common origin of gibberellin synthesis in fungi probable, although at present the exact scenario is still a matter of speculation. A gibberellin gene cluster might have been present in ancestors of specific ascomycete lineages, or the gene cluster might have been present ubiquitously in the last unknown common ancestor of this lineage with subsequent losses in most of the descendants. It is even possible that the gene cluster only evolved later, within one ascomycete lineage, and was horizontally transferred into the other clades. It is also unresolved whether this specialized terpenoid synthesis pathway is rooted in a bacterial metabolism (Bo¨mke and Tudzynski 2009). Apart from analysis of the whole gibberellin biosynthesis gene cluster, special research was also done studying evolutionary aspects relating to the terpene synthases involved in this and other metabolic pathways. Even neglecting the gibberellins, there exist more than 10000 different diterpenoid compounds that are ultimately all derived from geranylgeranyldiphosphate (GGDP). Similar to the gibberellin biosynthesis gene cluster, many of the corresponding fungal diterpene gene clusters contain orthologous and cluster-specific GGDP synthase genes. This led Zhang and


co-workers (2004) to present the hypothesis that the presence of such a dedicated GGDP synthase might be a conserved feature of fungal diterpene gene clusters. Moreover, it might represent the impact of ongoing metabolic channelling, keeping related enzymes and metabolites in close contact with each other to facilitate efficient synthesis of the end products. Saks et al. (2008) recently re-evaluated such micro-compartmentalization effects within the cell on a biophysical background. The fact that the GGDP-1 synthase involved in primary metabolism in G. fujikuroi is unable to complement the function of the gibberellin-dedicated GGDP-2 is interpreted as support for this thesis, as well as the observation that the clusters GGDP-2 gene is often linked to the terpene synthase gene of the pathway by a bifunctional promoter. The terpene synthases themselves also play a pivotal role in metabolic evolution. They are often mediating the committed step in a biosynthesis pathway, and the alteration of their product is a key event in the evolution of novel metabolites (Christianson 2006). The specificity of their product profile can be dramatically shifted by variation of only a few amino acid residues (Xu et al. 2007), which effects not only the direct reaction product, but also increases the overall reaction complexity. Whereas most terpene synthases generate a single product, like the ent-kaurene producing bifunctional CPS/KS of gibberellin-producing fungi, some others are less specific in their catalytic properties and catalyse the production of up to 52 different cyclization products. This highlights the function of such promiscuous enzymes as a playground for the evolution of either more specific pathways or altered enzyme functions (O’Brien and Herschlag 1999). Enzymes with a narrower specificity are then supposed to be the result of divergent evolution of more promiscuous ones with gene duplication and subsequent mutation in the duplicate, followed by metaboliteaimed selection are the proposed mechanisms (Aharoni et al. 2005). Nevertheless, to address enzymes producing multiple metabolites as the result of an ‘incomplete evolution’ of their catalytic site (Christiansen 2006) seems to be somewhat out of place, as evolution does not proceed towards any defined end point and each highly and uniquely specialized enzyme may still undergo future alterations or domain recombination leading to novel functions.

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Based on sequence similarity analysis, the fungal diterpene synthases probably evolved independently from the corresponding plant enzymes, but they are almost certainly all derived from one common ancestor. M. grisea, another putative gibberellin producer, was recently found to also contain two diterpene synthases with high similarity to the G. fujikuroi CPS/KS, for one of them functionality in the G. fujikuroi background could be proven (unpublished data, cited by Bo¨mke and Tudzynski 2009). Other data also hint at the existence of a degenerated gibberellin biosynthesis gene cluster in this species. In contrast, the functional gene might as well be the result of a recent mutation in a cyclase gene belonging into a different diterpenoid pathway. 5. Ergot Alkaloid Biosynthesis Ergot alkaloids are tri- or tetracyclic derivatives of prenylated tryptophan. Other constituents are a methyl group derived from S-adenosyl-methionine and an isoprene unit coming from mevalonic acid (for a review, see Schardl et al. 2006). The best known ergot alkaloid-producing species in Europe, with an exceptionally broad host range, is Claviceps purpurea. Its major alkaloid, D-lysergic acid, has a structural similarity with neurotransmitters like noradrenalin, dopamine and serotonin and can interact with receptors for these neurotransmitters. Biosynthesis of ergot alkaloids begins with the prenylation of tryptophan and the resulting dimethylallyltryptophan, the gene coding for the corresponding L-tryptophan dimethylallylsynthase was the first to be identified from the ergot alkaloid synthesis (EAS) gene cluster (Tsai et al. 1995; Tudzynski et al. 1999). The complete gene cluster, comprising 14 genes, was identified by chromosome walking starting from the L-tryptophan dimethylallylsynthase gene dmaW (Tudzynski et al. 1999; Correia et al. 2003; Haarmann et al. 2005). The cluster contains a set of genes encoding NRPS enzymes. One of these, the D-lysergylpeptide synthetase LPS2, catalyses the production of activated lysergic acid, which may act as recipient for the addition of several different peptide moieties. This may be the base for the broad ergot alkaloid spectrum produced by C. purpurea. An additional NRPS gene, lpsA2 also exhibits a significant degree of sequence identity with one of the other LPS-encoding genes, probably as the result of

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a rather recent gene duplication event (Haarmann et al. 2005). The ability for ergot alkaloid production is not a general property of the genus Claviceps. Some species (e.g., C. africana, C. purpurea) produce ergopeptines while others (e.g., C. fusiformis) end up with the formation of clavine alkaloids or lysergic acid amides, which are intermediates of the ergopeptide biosynthesis. Evolutionary aspects were studied in a comparison of the ergot alkaloid synthesis clusters between such differing strains, namely C. purpurea with ergotamine as the end product, C. fusiformis producing mainly elymoclavine, and C. hirtella with ergometrine and clavine alkaloids as the end products. Compared to the gene cluster in C. purpurea, C. fusiformis is lacking the three NRPS genes accumulated at one end of the cluster in C. purpurea. All other cluster genes are present, although lpsB is not longer functional and some rearrangement took place around that gene. lpsB is only required for one of the final steps in ergot alkaloid biosynthesis, explaining why in C. purpurea the synthesis pathway ends with clavine alkaloids. Another gene also required only for the final steps is CfcloA, a homologue of the C. purpurea cloA, an enzyme involved in the conversion of elymoclavine to paspalic acid. This gene is also expressed only as an inactive product. These findings strongly suggest that the C. purpurea EAS gene cluster is basal to the cluster present in C. fusiformis and that the latter evolved from a more complex ancestral situation (Lorenz et al. 2007). The situation in C. hirtella mirrors the close phylogenetic relatedness of that species to C. fusiformis. This begins with the more preserved sequence homology of the EAS gene products at the amino acid level between these two species. Here, too, the NRPS genes lpsA1 and lpsA2 are missing and the region surrounding lpsB has been rearranged. Further, lpsB itself is still functional and lpsC is also present, although at the other side of the gene cluster compared to C. purpurea. Down to the intron distribution pattern, the EAS gene cluster is highly conserved between these three species, and the presence of the functional synthesis genes explains to a large extent the final product of their respective alkaloid production. The fate of the cloA gene product in this context is still puzzling: despite a high rate of sequence identity, an identical exon–intron structure, and the same heme binding motif, the gene product is active in C. hirtella, but non-functional in C. fusiformis. It is now proposed that the clusters of C. hirtella and C. fusiformis

are derived from the C. purpurea-type, and that the rearrangement as well as the loss of the two NRPS genes took place before the two species separated. All other detail deviations concerning the functionality of LpsB and CloA occurred after that time (Lorenz et al. 2009).

Also, considerable variation in the alkaloid types was observed between strains of the same species. In C. purpurea, the variation in positions 1 and 2 of the tripeptide moiety of the final alkaloid product between the strain P1 and ECC93 (Haarmann et al. 2005) is the result of altered amino acid sequences in the region responsible for substrate recognition of the A domains of the first two modules of the trimodular NRPS. The cluster structure of the two strains is not deviating, suggesting that the sequence variation indeed is the base of the altered alkaloid types. 6. Other Non-Ribosomal Peptide Synthetases and Polyketide Synthase Pathways Non-ribosomal peptide synthetases (NRPS) are multifunctional enzymes synthesizing small peptides. Together with polyketide synthase (PKS) pathways they are also organized in gene clusters. Evolutionary processes in these clusters can be easily visualized as a matter of variations in the domain architecture, leading to either more elaborate and complex pathways or to degenerate and non-functional pathways (Hoffmeister and Keller 2007). (a) The ACE1 Avirulence Gene Cluster In M. grisea, the special metabolism gene cluster containing the ACE1 avirulence gene is specifically expressed during plant infection. ACE1 is a gene coding for a PKS-NRPS hybrid expressed specifically during penetration. Exchange of a single amino acid in the b-ketoacyl synthase catalytic site of ACE1 abolishes recognition of the fungus by resistant plant cultivars. This suggests that ACE1 biosynthetic activity is required for avirulence and that the detection of the invading fungal pathogen involves recognition of a secondary metabolite whose biosynthesis depends on ACE1 (Bo¨hnert et al. 2004).

The cluster consists of 15 genes, coding for two PKS-NRPS hybrids, two enoyl reductases, four cytochrome P450 monooxygenases, an a/bhydrolase, an oxidoreductases, a transporter, a zinc finger transcription factor, an O-methyl


transferase, and one protein of unknown function (Collemare et al. 2008). In a study aimed at finding evidence for HGT events between eukaryotic lineages, 26 fungal genomes were searched for the presence of at least three linked orthologues out of that cluster (Khaldi et al. 2008). Some single homologues to ACE1 cluster genes occur in several species, at diverse locations within the genome. Nine recognizable similar clusters were identified in seven ascomycetes, all belonging into the same subphylum as M. grisea, Pezizomycotina. Gene by gene analysis of the clusters led to the classification of two types, probably the result of an early division. The larger type retains eight or more genes of the cluster present in M. grisea, the smaller cluster type comprises only three to six of the genes. Complete ACE1 clusters were found in four species, A. clavatus and Chaetomium globosum contain two ACE1 clusters each. For three of the clustered genes the physical linkage is apparently stronger as for the others, as these genes are present in all identified clusters. The authors interpret this gene group as the ancient core of the recent cluster, existent already in the common ancestor of all genomes containing the ACE1 cluster. The recent large clusters are the result of repeated internal gene duplications events, in M. grisea evidence for additional tandem duplication within the cluster has been found. The cluster phylogeny roughly resembles the species phylogeny of the analysed species within the Pezizomycotina. The pattern of cluster organization and distribution within the lineages is therefore proposed to be the result of vertical gene transfer and multiple losses of individual genes or the complete pathway (Khaldi et al. 2008). One event of HGT is nevertheless inferred to have occurred for at least five cluster genes from a donor closely related to M. grisea into A. clavatus. (b) NRPS Pathways in Epichloe¨/Neotyphodium Species Members of the grass endophytic genus Epichloe¨ sp. (anamorph: Neotyphodium sp.) contain NPRS genes as core components of special metabolite gene clusters (see Chapter 10 in this volume). One product are loline alkaloids, potent fungal insecticides. The corresponding genes form a cluster with slight variations between the various isolates. All recent Neotyphodium spp. seem to have origi-

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nated from ancestral hybridization events between different species, with the only sexual loline alkaloid producing species, E. festucae (Schardl 2001) as one of the partners in these interspecific hybridizations. This view should better be extended to include common ancestors of the recent E. festucae and Neotyphodium species, as evidence from phylogenetic analyses not always supports E. festucae as direct ancestor of the recent loline producing species (Clay and Schardl 2002; Kutil et al. 2007). It is suggested that a selection for alkaloid producers established the descendants as endophytes (Schardl 2001). Compared to several Neotyphodium clusters, in E. festucae, a 19-kb region separates the genes lolC and lolD whereas in the other isolates, a maximum stretch of 1.7 kb separates these genes. This region contains residues of a degenerate reverse transcriptase suggesting a retrotransposon entering into this cluster at a time. This disruption is a probable reason for the observed low production level of lolines in E. festucae. For the analysed clusters, the order and orientation of the genes within the cluster is strictly conserved, each cluster has evolved as a single sequence. One bias for this strict conservation may be that this mirrors the effect of shared regulatory elements. The recognition elements for transcription factors also seem to be conserved for all genes within the clusters, and the position and order of the recognition elements within a promoter region was also conserved. The five clusters analysed have three evolutionary origins, and overall the trait seems so be on the verge of being lost in one more recent species (Kutil et al. 2007). Extending the analysis to related gene clusters, at least 12 NRPS genes were identified using a degenerate primer PCR approach in a number of symbiotic species of this species complex (Johnson et al. 2007). Most of the NRPS genes distributed among the different lineages seem to be derived from a common ancestor where they evolved through a combination of domain duplication (Damrongkool et al. 2005) and divergence. Their patchy distribution between the individual lineages is probably due to gene loss during species evolution, where most of the lost genes code for enzymes with non-essential functions within the pathway. In interspecies hybrids, common in this species complex, the number of NRPS genes is conserved or even lower that in the non-hybrid

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species, indicating that hybridization is frequently correlated with gene loss. Hybridization during species evolution may also be the cause for the loss of complete special metabolite pathways from some species. The authors discount the possibility of HGT as the cause for this discontinuous distribution because this would require the assumption of an improbably high number of independent HGT events.

VI. Conclusions From its beginning in studying metabolite diversity, understanding of the evolution of this branch of metabolism has gone a long way till its recent focus on elucidating how alterations in the product pattern between the species come about, and it is quite unpredictable where future research may lead. All facts, data and interpretations are pieces of a gigantic puzzle extending in four dimensions, and the complete structure will forever remain hazy in parts, as the past will never be accessible to direct analysis and it is still difficult to outline experiments suited to provide scientific evidence for evolutionary mechanisms at work. Evolution is an ongoing process, and while certainly none of the very first steps determining metabolic pathways will be in evidence anwhere today, all modulating and modifying evolutionary processes continue. Due to the complex network of factors determining metabolic processes at every step, very ancient, highly conserved pathways are retained besides ongoing extreme specialization, and all levels in between. Only true novelties will probably escape detection with the available set of research tools. Special metabolites underlie selective adaptation, with selection directed mainly at the function of a given compound within a rapidly changing, densely populated environment where all kinds of interactions between vastly different organisms and lifestyles take place. Selection, in contrast, also focuses on the conservation of success, leading to pathways more streamlined, with fewer opportunities for biochemical errors and uneffective side reactions, and with a higher level of overall redundancy, tightly controlled and linked interactively with other physiological regulators of metabolism.

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Biosystematic Index

A Absidia glauca, 303 Acanthoecidae, 12, 14–15 Acarosporomycetidae, 193 Acaulospora, 168, 177 Acaulosporaceae, 168 Acremonium chrysogenum, 237, 312, 314, 315 Agaricaceae, 139 Agaricales, 100, 110, 135–144 Agaricomycetes, 99, 100, 104, 111, 113, 123, 134–143 Agaricomycetidae, 135, 139, 145 Agaricomycetides, 146 Agaricomycotina, 97, 99, 102, 104, 115–117, 122, 123, 126–145 Agaricus species, 142 Aglaophyton, 171 Agrobacterium tumefaciens, 309 Agrocybe aegerita, 101 Allomyces macrogynus, 233, 260–263, 271 Alternaria A. alternata, 207, 238, 241, 243, 309, 310, 312 A. brassicae, 207 A. brassicicola, 207, 241 Ambispora, 166, 168, 178 Ambispora brasiliensis, 167 Ambisporaceae, 168 Amoebidiales, 10 Amoebidium parasiticum, 8 Amylostereaceae, 100 Amylostereum areolatum, 100 Aphelidea, 15 Arabidopsis thaliana, 172 Archaeospora, 168 Archaeosporaceae, 168 Archaeosporales, 168, 178 Arthoniomycetes, 194 Arthopyreniaceae, 193 Ascochyta rabiei, 241 Ascomycete, 101, 110, 115, 121 Ascomycetous yeasts, 106, 115, 116 Ascomycota, 97, 101, 102, 106, 108, 115, 122, 144 Aspergillus, 305, 308, 317 A. clavatus, 263, 294, 321 A. flavus, 240, 263, 312 A. fumigata, 294 A. fumigatus, 85, 90, 214, 233, 239, 241, 243, 258, 263, 266, 268, 269, 294, 296, 305, 306, 308, 311, 316

A. nidulans, 90, 101, 214, 233, 235, 237, 258, 266, 268, 294, 295, 300, 305, 312, 314, 315 A. niger, 233, 263 A. oryzae, 233, 263, 294 A. parasiticus, 312 A. sojae, 312 A. terreus, 263, 316 Auricularia A. auricula-judae, 99, 134, 135 A. polytricha, 99, 134, 135 Auriculariaceae, 99 Auriculariales, 99, 134–135 B Baker’s yeast, 106, 115 Barley smut, 119 Basidiomycetes, 97–101, 103–110, 113, 115, 116, 122, 126, 127, 129, 135, 139, 142, 144, 145 Basidiomycetous yeasts, 97 Basidiomycota, 97, 101, 102, 106–108, 115, 116, 122, 144–146 Basidiospores, 132 Batrachochytrium dendrobatidis, 233, 258, 263 Bdelloidea, 121 Bilateria, 9 Bipolaris oryzae, 243 Blakeslea trispora, 302 Blastocladiomycota, 6 Blastomyces dermatitidis, 241 Botryotinia fuckeliana, 310 Botrytis cinerea, 79, 225, 233, 310 Bullera, 133 Burkholderia B. endofungorum, 309 B. rhizoxinica, 309 C Candida albicans, 73, 82, 101, 241, 258, 264, 266 Candidatus, 174 Cantharellales, 99, 135 Capsaspora, 7, 15–16 Ceratobasidiaceae, 99 Cercospora, 236 C. beticola, 207 C. cruenta, 310 Chaetomium globosum, 79, 214, 316, 321 Chaetothyriomycetidae, 192

Evolution of Fungi and Fungal-Like Organisms, The Mycota XIV S. Po¨ggeler and J. Wo¨stemeyer (Eds.) © Springer-Verlag Berlin Heidelberg 2011

332

Biosystematic Index

Chytridiomycota, 6, 115, 145, 169 Ciliate, 102 Cladosporium carrionii, 241 Claroideoglomeraceae, 167, 168 Claroideoglomus, 168 Claroideoglomus claroideum, 172 Claviceps, 239, 305 C. africana, 320 C. fusiformis, 320 C. hirtella, 320 C. purpurea, 239, 319, 320 Coccidioides C. immitis, 233, 269 C. posadasii, 241 Cochliobolus C. carbonum, 238, 241, 306, 307 C. heterostrophus, 101, 236, 310 C. heterostrophus (anamorph Bipolaris maydis), 206 C. miyabeanus, 241 C. victoriae, 207 Codosigidae, 12 Colletotrichum lagenarium, 241, 243 Coprinellus disseminatus, 100, 105, 109, 125, 139, 140, 143, 145 Coprinopsis, 138 C. cinerea, 100, 102, 105, 110, 111, 123, 125, 127, 135–138, 140, 141, 143, 144, 233, 270 C. scobicola, 100, 105, 136, 137 Corallochytrea, 16–17 Corallochytrium, 16 Corynespora cassiicoli, 207 Cronartiaceae, 98 Cronartium quercuum, 98, 114 Cryphonectria parasitica, 77, 89 Cryptoccocus, 131–133 C. amylolentus, 99, 130, 131, 134 C. gattii, 99, 101, 105, 109, 110, 123, 126, 130, 132–134 C. heveanensis, 99, 105, 110, 123, 129–131, 133, 134 C. neoformans, 101–103, 106, 107, 109, 121, 125, 127, 130, 132, 133, 135, 139, 145, 146, 233, 241, 244, 258, 270, 316 mating-type locus, 285 C. neoformans var. grubii, 99, 105, 110, 123, 126, 128, 132, 133 C. neoformans var. neoformans, 99, 105, 110, 111, 123, 126, 127, 130, 132–134, 139, 142 species, 101, 128–134 species complex, 126, 130 varieties, 132, 134 varieties/species, 131 Curvularia protuberata, 241 D Dictyostelium D. deminutivum, 57 D. discoideum, 57–59, 102, 232, 258, 260, 261 D. menorah, 57 D. minutum, 67 Dikarya, 5, 44, 97, 115, 122, 144 Diplocarpon rosae, 241, 244 Diplodia gossypina, 241 Diversispora, 168

Diversisporaceae, 167, 168 Diversisporales, 167, 168 Dothideomycetes, 193 Dothistroma septosporum, 317 E Eccrinales, 10 Elsinoe brasiliensis, 318 Embryophyta, 163 Emericella nidulans, 294 Entrophospora, 168 Entrophosporaceae, 168 Epichloe¨, 239, 294, 312, 321 Epichloe¨ festucae, 84, 238, 321 Escherichia coli, 304 Eurotiomycetes, 192, 316 Eurotiomycetidae, 193 Exobasidiomycetides, 99, 121, 143 F Filamentous ascomycetes mating-type loci, 277 Filasterea, 15 Filobasidiaceae, 99 Filobasidiales, 99, 123, 126–133, 142 Filobasidiella, 127 F. amylolenta, 99, 131 F. depauperata, 99, 131 F. neoformans, 99 Folsomia candida, 236 Fomitopsidaceae, 100 Fomitopsis palustris, 100, 125, 126 Fonsecaea pedrosoi, 241 Fungi, 5–7 Allomyces, 28 harpellalean fungi, 29 Magnaporthe grisea, 32 Malassezia, 30 Neocallimastigaceae, 31 Pneumocystis carinii, 30 Rozella, 28 Rozella allomycis, 28 Saccharomyces cerevisiae, 30, 32 Funneliformis, 168 Funneliformis mosseae, 167, 169 Fusarium, 311 F. avenaceum, 238 F. graminearum, 214, 263, 311, 316 F. moniliforme, 234 F. oxysporum, 84, 310 F. proliferatum, 318 F. venenatum, 234 F. verticilloides, 84, 318 G Gaeumannomyces graminis, 241 Geosiphon, 168 Geosiphonaceae, 168 Geosiphon-Nostoc, 172 Geosiphon pyriformis, 166, 167, 172

Biosystematic Index Gibberella G. fujikuroi, 239, 302, 309, 316–319 G. zeae, 84, 90, 232, 233, 236 Gigaspora, 168 G. margarita, 174 G. rosea, 174 Gigasporaceae, 167, 168, 170, 174, 175 Glarea lozoyensis, 241 Glomeraceae, 165, 168 Glomerales, 164–168 Glomeromycete, 48 Glomeromycota, 6, 115, 163, 164, 166–171, 173, 174, 176–178, 180 Glomeromycota-Mortierellales, 168 Glomus, 48, 165, 166, 168 G. macrocarpum, 165 G. mosseae, 165 Guignardia mangiferae, 241 Gymnosporangium asiaticum, 98, 114 H Histoplasma capsulatum, 241 Holozoa, 3, 16, 17 Hydnaceae, 100 Hydnangiaceae, 100 Hypocrea jecorina, 84, 300 I Ichthyophonae, 11–12 Ichthyosporea, 10–12 J Jelly fungi, 97, 134 Jelly-like fungi, 97 K Kluyveromyces lactis, 101, 308 Kwoniella, 133 K. heveanensis, 99, 133 K. mangroviensis, 99, 133 L Laboulbeniomycetes, 192 Laccaria bicolor, 100, 105, 106, 110, 111, 124, 127, 136–138, 140, 141, 143, 233, 270 Lecanoromycetes, 193 Lecanoromycetidae, 193 Lentinula edodes, 100 Lentinus squarrosulus, 100 Leotiomycetes, 194 Leptosphaeria L. biglobosa, 206 L. maculans, 238, 296, 316 L. maculans (anamorph Phomalingam), 206 Lichinomycetes, 192 Lyngbya majuscule, 306 M Macrophomina phaseolina, 241 Magnaporthe

333

M. grisea, 79, 214, 232, 233, 241, 243, 268, 294, 306, 309, 316, 319, 320 M. oryzae, 241 Malassezia M. globosa, 99, 105, 109, 110, 117, 121, 233, 270 M. restricta, 99, 121 Malasseziaceae, 99 Malasseziales, 99, 120–121 Malbranchea aurantiaca, 239 Marasmiaceae, 100 Melampsoraceae, 98 Melampsora M.larici-populina, 98, 105, 106, 108, 115, 122 M. lini, 98, 114 Mesomycetozoa, 3, 10 Metazoa, 8–9 Microbotryaceae, 98 Microbotryales, 98, 114, 142 Microbotryomycetes, 98, 109–114 Microbotryum, 114 Microbotryum species complex, 102, 114 Microbotryum violaceum, 98, 104, 110, 114, 116, 121 mating-type chromosomes, 286 Microsporidia, 6, 37 Antonospora locustae, 28, 30–33 Antonospora (Nosema) locustae, 27 Brachiola algerae, 33 Edhazardia aedis, 33 Encephalitozoon cuniculi, 25, 27, 28, 30–33 Encephalitozoon hellem, 27 Enterocytozoon bieneusi, 28, 30, 32, 33 Nosema ceranae, 33 Spraguea lophii, 27, 32 Trachipleistophora hominis, 31 Vairimorpha necatrix, 26 Ministeria, 15, 16 Mitosporic Marasmiaceae, 100 Mitosporic Tremellales, 99, 133 Moniliophthora perniciosa, 100, 105, 124, 141 Monosiga brevicollis, 8, 13 Mucorales, 303 Mucor M. circinelloides, 233, 298, 304 M. mucedo, 302, 303 Mucoromycotina, 115 Mushroom, 102, 105, 106, 144 Mycosphaerella M. cruenta, 310 M. fijiensis (anamorph Pseudocercospora fijiensis), 207 M. graminicola (anamorph, Septoria tritici), 207 N Nectria haematococca, 84, 233, 310 Neocallimastigomycota, 6 Neocosmospora boniensis, 310 Neosartorya N. fischeri, 101, 305, 306, 316 N. fumigata, 85, 90, 294 Neotyphodium, 239, 294, 312, 321 Neotyphodium lolii, 240

334

Biosystematic Index

Neurospora crassa, 79, 85, 89, 121, 214, 232, 233, 309, 312, 313, 317 Neurospora tetrasperma, 82 mating-type chromosomes, 281 phylogenetic species, 282 Nodulisporium sp., 241 Nostoc punctiforme, 166 Nuclearia, 7, 8, 15 Nucleariida, 7–8 O Ophiostoma O. floccosum, 241 O. piceae, 241 O. piliferum, 244 Ophiostoma nova-ulma mating-type locus, 287 Opisthokonta, 3, 5 Opithokonts, 43 Ostropomycetidae, 193 Otospora, 168 P Pacispora, 168 Pacisporaceae, 168 Paracoccidioides brasiliensis, 241, 269 Paraglomeraceae, 168 Paraglomerales, 168 Paraglomus, 166, 168 Parasitella parasitica, 303 Passalora fulva (syn. Cladosporium fulvum), 207 Penicillium, 305 P. chrysogenum, 85, 233, 237, 314, 315, 317 P. citrinum, 312 P. expansum, 312 P. marneffei, 241, 243 P. paxilli, 240 Pezizomycotina, 115, 192 Phaeosphaeria, 240, 318 P. nodorum, 316 P. nodorum (anamorph Stagonospora nodorum, syn. Leptosphaeria nodorum, syn. Septoria nodorum), 205–206 Phanerochaetaceae, 100 Phanerochaete chrysosporium, 100, 105, 124, 139, 140, 143 Pholiota P. microspora, 100, 105, 139–141 P. nameko, 100, 105, 139 Phycomyces blakesleeanus, 259, 263, 302, 312 mating-type locus, 287 Phytophthora P. infestans, 258–260, 271 P. sojae, 304 Pichia stipitis, 264, 266 Piromyces, 300 Pleospora infectoria, 241 Pleurotaceae, 100 Pleurotus djamor, 100, 102, 105, 124, 137, 138, 143 Pneumocystis carinii, 73 Podospora anserina, 79, 217, 233, 241, 244

Polyporaceae, 100 Polyporales, 100, 135–142 Polyporus palustris, 100, 125, 126 Polyshondylium P. pallidum, 67 P. violaceum, 67 Populinus, 165 Postia placenta, 100, 233 Psathyrellaceae, 100 Pucciniaceae, 98 Puccinia, 114 P. coronata, 98, 114 P. graminis, 98, 102, 105, 110, 111, 114–116, 122 P. graminis f. sp. tritici, 115 P. triticina, 98, 105, 115 Pucciniales, 98, 114–115, 122 Pucciniomycetes, 98, 111, 113–115 Pucciniomycotina, 97, 98, 102, 104, 109–116, 121, 122, 135, 142, 145 Pyrenophora tritici-repentis, 233, 310 Pyrenophora tritici-repentis (anamorph Drechslera triticirepentis, syn. Helminthosporium tritici repentis), 206 R Racocetra, 167, 168 Radiata, 9 Redeckera, 168 Red yeast, 109 Rhinosporideacae, 10 Rhizoctonia solani, 99, 110, 111, 135 Rhizophagus, 165, 168 R. intraradices, 178 R. irregularis, 168, 175, 178 Rhizopus R. microsporus, 309 R. oryzae, 233, 259, 263, 306 R. stolonifer, 73 Rhodosporidium R. babjevae, 98, 104, 112 R. toruloides, 98, 108–111, 113 Rotifers, 121 Rozella, 6–8, 45 Russulales, 100, 135–142 Rust, 102, 106, 108, 114, 115, 122 Rust fungi, 97 S Saccharomyces, 300 S. castelli, 313 S. cerevisiae, 73, 75–77, 101, 102, 106, 107, 110, 115, 116, 144, 214, 233, 258, 264, 266, 269, 270, 296, 304, 308, 312–314, 316 Saccharomycotina, 115, 191 Salpingoecidae, 12 Schizophyllaceae, 100 Schizophyllum, 138 Schizophyllum commune, 100, 102, 105, 110, 111, 124, 125, 127, 136–139, 141, 143, 144 Schizosaccharomyces, 301, 316 Schizosaccharomyces pombe, 73, 76, 101, 214, 233, 264

Biosystematic Index Sclerocystis, 165, 168 Sclerotinia S. minor, 241 S. sclerotiorum, 233, 241 S. trifoliorum, 241 Sclerotium cepivorum, 241 Scutellospora, 168, 177 Scutellospora heterogama, 167 Scutellosporites devonicus, 170 Scytalidium dimidiatum, 241 Sebacinales, 172 sensu lato Kwoniella clade, 99, 131, 133 Silene latifolia, 114 Sistotrema brinkmannii, 100, 135 Slime mold, 102 Smut, 97, 102, 114, 116, 118–120 Smut-like fungi, 97 Soil clone group 1, 45 Sordaria, 301 Sordaria macrospora, 79, 85, 90, 101, 214, 233, 241, 243, 258, 265, 266, 268 Sordariomycetes, 194, 316 Sphaceloma manihoticola, 240, 317, 318 Spizellomyces punctatus, 258–259, 263 Sporidiales, 145 Sporidiobolaceae, 98 Sporidiobolales, 98, 108–113, 122, 124, 142, 146 Sporidiobolus S. johnsonii, 98, 104, 110, 112, 116, 125 S. metaroseus sp. nov., 112 S. pararoseus strain CBS 484, 112 S. salmonicolor, 98, 104, 108–113, 116, 121, 122, 125 S. salmonicolor (strain IAM 12258), 109 spec. strain IAM 13481, 110, 111 Sporisorium, 126 S. reilianum, 99, 105, 110, 117–120, 125, 126, 134, 146 S. scitamineum, 120 Sporobolomyces S. metaroseus, 98 S. roseus, 98, 108, 112 Sporobolomyces spec. IAM 13481, 98 Sporobolomyces spec. strain IAM 13481, 104, 108, 112, 113, 116, 122 red yeasts strain IAM 13481, 112 strain IAM 13481, 112 Sporothrix schenckii, 241 Stagonospora nodorum, 233. 310 Streptomyces, 308 Strophariaceae, 100 T Taphrinomycotina, 115 Taphrinomycotina, 191 Tetragastis panamensis, 179 Tetrahymena thermophila, 102 Thanetophorus cucumeris, 135 species complex, 99

335

Tilletiaceae, 99, 121 Tilletia T. controversa, 99, 121 T. tritici, 99 Tilletiales, 99 Tremella T. atroviride, 84 T. brasiliensis, 99, 134 T. mesenterica, 134 Tremellaceae, 99, 133, 134 Tremellales, 99, 123, 133–134 Tremella mesenterica, 99, 105, 123, 134 Tremellomycetes, 99, 100, 104, 110, 111, 113, 126–135 Trichoderma T. reesei, 84, 300 T. virens, 84, 316 Truffle Tuber melanosporum, 232 Trypetheliaceae, 193 Tsuchiyaea wingfieldii, 99, 130, 131 Tuber melanosporum, 233 U Unikonta, 5 Ustilaginaceae, 98 Ustilaginales, 98, 116–120, 126, 142 Ustilaginomycetes, 98, 116–120 Ustilaginomycotina, 97, 98, 100, 102, 104, 110, 111, 113, 115–126, 135, 142, 145 Ustilaginomycotina incertae sedis, 99, 120–121 Ustilago, 126 U. hordei, 98, 102, 105, 109, 110, 117, 118, 120–122, 124–126, 139, 145 U. maydis, 98, 105, 106, 110, 113, 116–122, 125, 126, 133, 134, 141, 233, 270, 308, 316 U. scitamineum, 99 Ustilagomycetes, 126, 134, 143 V Venturia inaequalis, 207 Verticillium V.albo-atrum, 241 V. dahliae, 241 V. nigrescens, 241 V. tricorpus, 241 X Xanthophyllomonas dendrorhous, 298, 304 Y Yeast, 97, 109, 112, 113 ascomycetous, 116 Z Zygomycetes, 164, 297, 301–303 Zygomycota, 5, 6, 115, 122166 Zygomycota sensu lato, 106

.

Subject Index

A a/a cell identity, 127, 128 a1 and a2 mating type genes, 126 a1 and a2 mating type loci, 118 Aa sublocus, 136, 137 Abscisic acid, 295, 302, 311 Ab sublocus, 136, 137 Acetate, 232 ACG, 63, 64, 67 ACV synthetase, 237 Adaptive evolution, 311 Adaptive responses, 295 Adenylate cyclase A (ACA), 61 a-factor, 106, 144 a-factor, 116, 144 Aflatoxin, 235, 312 Aflatrem, 240 a gene pair, 136 Aggregation, 57–59 Aglaophyton major, 170 AKOR, 122 Alcohol dehydrogenase, 302 Algae, 173 Alkaloids, 232, 239 A mating type, 134 A43 mating type, 137 a mating type alleles, 126 A mating type loci, 144 A mating type locus, 133, 135, 139, 140, 142, 143, 145 archetype, 136 A43 mating type locus, 136 a mating type locus (P/R), 116 Aminoadipate reductase, 238 Aminopeptidase, 75 Ammonia, 60, 62, 63 Amoebozoa, 57 Amphimixis, 101 Anamorphic, 103 Anamorphic (asexual), 144 Anastomosis compatibility groups hyphal networks, 176 vegetative incompatibility, 176 Ancestral character state reconstruction, 196 Angiocarpous development, 188 Anisogamy, 101 Antibiotics, 237, 294 A pathways, 141

a-pheromone MFa, 115 a-pheromone MFa, 116 a-pheromone receptor Ste2, 115 a-pheromone receptor Ste3, 116 Appressoria, 244 a1-protein, 116 Arbuscular mycorrhiza fungi, 163 acaulosporoid spores, 177 arbuscules, 165 Arthur Schu¨bler and Christopher Walker, 166 asexual fungi, 176 biogeography, 179 co-evolution, 164 concerted evolution of rDNA repeats, 175 DAOM197198, 169, 176 DAOM198197, 178 diversification, 172, 177 diversity, 179 evolution, 164 germination shields, 175, 177 germ tube, 177 heterokaryotic, 175 homokaryotic, 175 hyphal network, 175 isolate recognition, 178 LSU rDNA, 178 microsatellite, 178 mitochondrial markers, 178 molecular clock estimates, 170 molecular diversity, 179 molecular ecological studies, 179 molecular phylogeny, 166, 178 molecular species concept, 178 natural systematics, 166–167 phylogenetic species concept, 177 Red Queen hypothesis, 175 speciation, 180 species recognition, 178 spore ontogeny, 165 systematics, 167, 178 taxonomy, 167 Arbuscules, 163, 169 Aromatic prenyl transferase, 305 Ascogenous hyphae, 91 Ascoma development ascohymenial, 188 ascolocular, 188

338 Ascomycota, 205 Ascus types bitunicate, 189 prototunicate, 189 unitunicate, 189 Asexual (anamorphic), 112 Asexual eukaryotes, 163 Asexuality, 142 Asexual lineages, 121 a-specific genes, 116 a-specific genes, 115 a strains, 127, 132 a strains, 127, 132 Autosomes, 108 Axl1, 107 B Ba and Bb, 138 Bacteria-like organisms (BLO), 174 Ba receptor, 138 Basidia, 97, 101 Basidiospores, 97, 101 Bb receptor, 138 bE and bW genes, 119 bE1 and bW1 genes, 120 Beetles, 38 bE1 gene, 121 b-fg, 136, 139, 140, 143 b gene pair, 136 Biogeography, 166–167 Biological activity, 296 Biotin biosynthesis yeast, 314 Bipolar heterothallic species, 102 Bipolarity, 121, 122, 125, 142, 145 b-Lactam antibiotics, 314 B locus, 113 B mating type, 140 B41 mating type, 138 B43 mating type, 137 B mating type-like locus, 140 B mating type locus, 116, 133, 135, 140, 142, 143, 145 archetypal model, 137 multi-allelic, 119 B mating type pathway, 141 b-oxidation, 300 B-regulated development, 140 Bryophytes, 170, 172, 173 BSP1, 129 BSP2, 129, 131 Burkholderia, 174 bW1 gene, 121 C CaaX protease, 75 Carbonic anhydrases, 257 b-class, 257, 258, 261, 263, 265, 268–270 cab-type, 265, 266, 268, 269 a-class, 257, 258, 261, 263, 264, 270 plant-type, 265, 266 Carboxy-methylation, 75

Subject Index Carboxypeptidase, 75 Carotene apocarotenoids, 302 cleavage oxygenase, 301 developmental regulation, 302 synthesis, 301 Carotenoids, 295 CcSte3.1, 137, 138 CcSte3.3, 137, 138 CcSte3–2151, 137, 138 CcSte3–2153, 137, 138 CcSte3–7395, 138 CcSte3.2a, 137 CcSte3.2b, 137 CDPHB1, 140 CDPHB2.1, 140 CDPHB2.2, 140 CDSTE3.1, 139, 140 CDSTE3.3, 139, 140 Cell-cell fusion, 97 Cell invasion aquaporin, 25 germination, 25 polar filament, 25 polar vacuole, 25 sporogony, 26 Cellobiose dehydrogenase, 299 Cellular and genomic reduction degeneration of organelles, 29 DNA repair, 30 electron transport chain, 30 genome compaction and reduction, 30 genome reduction, 30 Golgi, 32 Krebs cycle, 30 mitosome, 31 peroxisome, 32 signaling processes, 31 vacuole, 32 Cellular fusion, 143 Cell wall, 37, 39, 45, 49 Chemical diversity, 297 Choanoflagellates, 12, 14–17 Chromatin, 235 modifications, 312 CID1, 121, 129, 131 Cilium, flagellum, 37 Clamp cells, 91, 127, 132, 135, 140 Clamp connections, 101, 132 Classification a-and b-tubulin, 28 archezoa, 26 early-branching eukaryotes, 27 EF–2, 27 EF1A, 28 EF1-a, 27 long-branch attraction, 27 RPB1, 28 RPB2, 28 sporozoa, 26 Cluster, 235, 237

Subject Index

339

CnnCpr2, 142 Codon usage, 284 Co-evolution, 294, 297 Communication, 311 inter-specific, 298, 303 pathways, 143 sexual communication, 302 Compound property, 297 Condensation, 237 Conjugation tube, 129, 134 Conjugation tubes, 101 Core chytrids, 6 Courtship, 107, 134 Cpr2, 134 Cryptic diversity, 38 Cryptic sexual reproduction, 175 Cyanobacteria, 171, 173, 180 cyclic adenosine monophosphate (cAMP), 260, 261

F Fatty acid synthases (FAS), 232 Fertilization, 86 Flavoenzyme, 299 Fluorescence in situ hybridisation, 48–49 Fossil record, 169, 170 germination shields, 170 Rhynie chert, 169, 170 Fruiting body, 59, 60, 91, 237 light-regulated, 141 A mating type-induced, 141 Fungal–grass symbiosis, 239 Fungi, 57 homothallic, 101 Fusarium compactum, 311 Fusion cell-cell, 101 nuclear, 101

D Deep-sea, 43 deep-sea fungal pathogens, 43 Degeneration, 285 Dehydrogenase sorbitol, 300 Deletion, 235 Development, 236, 302 d gene pair, 136 DIF–1, 62, 67 Dikaryon, 101 Dimethylallyl tryptophan synthase (DMATS), 239 Dimorphic fungi, 134, 139 Dioecism, 129 Dispensable gene, 296 Diversity, genetic, 298 Dmc1, 132 DNA barcoding, 178 Dothideomycetes, 205

G Galactose utilization, 310 Gametes, 101 GEF1, 129, 131 Gene co-expression, 313 dispensable, 296 duplication, 235, 257, 258, 262, 266, 270, 271, 299, 306, 321 family, 257, 258, 270 fusion, 304 homeodomain transcription factor, 103, 108 mating-type-like, 103 pheromone precursor, 106 for pheromone precursors, 115 pheromone receptor, 106, 109 regulation, 311 sex-related, 130 silencing, 308 Gene cluster, 231, 235, 296, 306, 312 co-regulation, 312 evolution, 307 location, 308 transcription factors, 307 yeast, 313 Gene expression co-regulation, 308 Gene family evolution, 306 Genome duplication, 259 reorganization, 313 sequencing project, 175 structure, 164 Genome ‘finishing,’ 217 Geosiphon-Nostoc symbiosis, 173 Germination, 64, 177 Germination shields, 165 Gibberellin, 239, 295, 310 Gibberellin biosynthesis evolution, 317, 318 Glomeribacter, 174 Glomeromycota asexual evolution, 174 endobacteria, 174

E Ectomycorrhizas, 172 e gene, 137 Endogone, 165 Endopeptidase, 81 KEX2, 81 Endoprotease, 75 Endosymbionts, 294, 309 Enniatin, 238 Environmental DNA, 38, 39, 44, 47–50 Environmental gene library analysis, 39, 48 Epipolythiodioxopiperazine toxins, 316 Ergot, 239 Ergot alkaloids biosynthesis, 319 D-lysergic acid, 319 Ergotamine, 239 Ergovaline, 239 Evolution eukaryotic asexuals, 174 flagellum, 169 neutrality, 297 tubulin, 169

340

Subject Index

GPB1, 132 GPD, 122 G protein activation, 86 b-subunit, 132 heterotrimeric, 108 G protein-coupled receptor, 133 Gymnocarpous development, 188 H Haploid fruiting, 132 HC-toxin, 238 HD, 135, 142 HD1 and HD2 homeodomain transcription factors, 119, 135 HD1 and HD2 SXI1/SXI2, 129, 131 HD1 gene, 137, 140, 141 HD2 gene, 136, 137, 140 HD1/HD1, 103, 111, 112, 131, 142 HD2/HD2, 103, 111, 112, 131, 142 HD1-HD2 fusion genes, 141 HD1/HD2 gene pair, 111, 113, 114, 120, 134, 140, 145 HD1/HD2 genes, 112, 119, 121, 122, 125, 130, 132, 133, 139, 145 bE, 113 bW, 113 pair locus, 115 HD1 homeodomain transcription factor, 127 HD2 homeodomain transcription factor, 127 H/D loci, 119 HD locus, 108, 109, 130, 131, 145 Heterodimeric complex, 127 Heterothallic, 89–90, 97, 112, 135 Heterothallic bipolar, 144 Heterothallic mating, 132 Heterothallic tetrapolar, 144 Heterothallism, 73 Heterozygote, 311 Histidine kinase, 62 Histone deacetylase, 235, 312 Histone methyltransferase, 312 Homeodomain, 111 atypical, 103 canonical, 103 DNA-binding motif, 103 HD1, 103 HD2, 103 protein, 74 TALE, 103 Homeodomain transcription factors, 102, 144 classical, 144 TALE, 144 Homopolymer 454 errors, 48 Homothallic, 90–91, 97, 112, 135, 141, 144 secondary, 137 Homothallism, 73, 132, 142 biparental, 101 facultative, 139 primary, 97 secondary, 97 uniparental, 101 Horizontal gene transfer (HGT), 232, 235, 264, 299, 300, 309, 310, 315 Hornworts, 174

Host recognition, 294 Hydroxynaphtalene reductase, 243 Hyphae, 101 I Idiomorph, 74, 101, 117 Immunosuppressive drugs, 237 Inbreeding, 101, 122, 135 Inbreeding avoidance, 122 Inbreeding potential, 142 Inbreeding restriction, 144 Indole-diterpene, 239 Inter-species interaction, 295 Intraspecific variability, 178 Iron homeostasis, 238 Isoprenoids, 239 Isotropic cell growth, 129 Isozyme, 258, 261, 263, 265, 268–271 ITS, 38, 41, 45, 46, 48 K KAP95, 122 L Laccase, 242 LaeA, 235, 236 Land plant evolution, 164, 166, 169, 171–173 Lateral gene transfer, 224 LbSte3.1, 138 LbSte3.2, 138 LbSte3.3, 138 LbSte3.4, 138, 141 LbSte3.5, 138 LbSte3.6, 138 LbSte3.7 to LbSte3–13, 138 lga2, 117 Lichen, 294 Ligand-induced endocytosis, 86 Ligand receptor-pair, 297, 298 Lipids, 294 Lipopeptides, 107 Liverwort, 169 LKM11, 45, 49 Localization cytosolic, 259, 262, 263, 269 mitochondrial, 260–262, 268, 269, 271 secreted, 259, 261, 262, 264 Loculoascomycetes, 188 Loline alkaloids, 239, 321 Lolitrems, 240 Long-branch attraction, 4 Lower fungi, 43, 44 LPD1, 129, 131 LSm7, 112, 122 Lysergic acid, 239 Lysin biosynthesis, 238 M Macroconidium, 86 Macrocyst, 58, 66 MAP kinase cascade pheromone-sensing, 129

Subject Index MAP kinase pathway, 130 MAP kinase signalling pathway, 130 Marine fungi, 42–43 MAST group, 48 MAT, 111, 121 MAT–1, 120, 121 Mat2, 132 MAT–2, 120 MATa, 115, 127, 131 MATa, 115, 121, 127, 131 MATA1, 109 MATA2, 109 MATaa cells, 129 mat chromosomes, 284 MAT genes, 199 Mating, 102 behaviour, 97 bipolar, 102 heterothallic, 101 intratetrad, 114 molecular control type, 97 same-sex, 101 self-fertile, 101 species specificity, 303 tetrapolar, 102 unipolar, 102 Mating behaviour bipolar, 111 Mating factor, 74 Mating system ancestral, 122 bipolar, 114, 139 evolutionary stable, 142 mushroom, 144 tetrapolar, 108 transitory, 142 Mating type, 102 cassettes, 101 compatible, 107 configuration, 97 control, 126 determination, 142 diverged alleles, 144 genes, 103, 106 homeodomain transcription factors, 103 idiomorphs, 101 incompatibility, 102 intercompatibility, 118 locus, 101 MAT locus, 101 multiple, 102 paralogous genes, 144 pheromone precursor, 106 pheromone receptor, 106 self-compatible alleles, 146 specificities, 118, 144 specificity, 103, 136 specific phylogeny, 129 Mating type (MAT), 73 Mating type alleles multiple, 122

Mating-type chromosomes Neurospora tetrasperma, 281 parallels to sex chromosomes, 281 Mating type genes allelic, 136 duplication, 144 functional redundancy, 136 paralogous, 136, 144 Mating type-like genes, 142 Mating type loci alleles, 102 duplication, 135 filamentous ascomycetes, 277 hyperallelism, 135 mating system, 279 multi-allelic, 117 recombination, 135 subloci, 135 Mating type locus biallelic, 106 gene conversion, 112 HD1/HD2 locus, 105 P/r locus, 106 Mating type system biallelic, 109 multi-allelic, 143 one-locus, 109 tetrapolar, 143 MAT loci, 113, 115, 129, 130, 133, 145 MAT locus, 115, 120, 127, 130 Meiosis, 101, 129 Melanin, 232, 240 DHN, 241 DOPA, 241 GDHB, 241 Metabolic channelling, 307, 319 Metabolic network, simplification, 303 Metabolism, 294, 304 Metabolism/metabolic, 299 Metabolite potency, 296 Methyltransferases, 235 MFa, 129, 132 mfa, 120, 126 MFa1, 129, 131 MFa2, 129, 131 MFA2, 133 mfa2, 117 mfa2.3, 118 Mfa3, 126, 129, 131 mfa1/MFA1, 117, 133 MFa, 106, 129 MFa1, 116, 129, 132 MFa2, 116, 129, 132 MFa3, 129, 132 MFa4, 129 MFa pheromones, 106 mfa pseudogene, 117 MF genes, 129 MgMfa1, 120 Microbial loop, 45 Microcyst, 58, 66

341

342 mip, 136, 139, 140, 143 Mitochondrial genomes, 219–221 Mitochondrial intermediate peptidase, 136 Mitochondria, uniparental inheritance, 117 Mitogen-activated protein (MAP) kinase pathway, 129 Mobile genetic element, 308 Modified tetrapolarity, 125 Molecular clock, 173 Molecular phylogeny, 166 actin, 168 a-tubulin, 168 EF1a, 167 EP1a, 168 H+-ATPase, 168, 169 LSU rRNA, 167 P-type II ATPases, 169 RPB1, 167 RPB2, 167 rRNA gene, 167 5.8S rRNA, 167 SSU rDNA, 168 SSU rRNA gene, 167 tubulin, 169 Molecular property, 297 Mollicutes, 174 Monokaryons, filamentous, 101 Monokaryotic fruiting, 132, 139, 143 Monophyletic origin, 97 MTL1, 114 Multi-allelism, 142 Multicellularity, 57, 58 Multi-drug resistance pump Mdr, 107 Multifunctional proteins, 304 Multi-gene analyses, 167 Mutant B mating type locus, 141 A mating type genes, 141 Mycelium, 97 Mycoplasma, 174 Mycorrhizal, 37, 46, 48 Mycorrhizal fungi, 166 Mycotoxins, 294 MYO2, 131 N Natural products, 231, 293 Neofunctionalization, 258, 262, 263, 268–270 Neurospora mating-type genes, 279 Nonribosomal peptides, 232, 237 Nonribosomal peptide synthetases (NRPS), 233, 237, 319, 320 Nostoc, 173 Novelty, 296 Nuclear fusion (karyogamy), 101 Nuclei, haploid, 101 Nutrient acquisition, 172 O Obligate symbionts, 172 Opisthokont phylogenies, 4

Subject Index Opposite-sex mating, 132, 133 Orchid-mycorrhizas, 172 Origins of multicellularity, 17–18 Osmotrophic, osmotrophy, 37 Outbreeding, 144, 146 Outbreeding efficiency, 145 Outbreeding potential, 116, 122, 142 Outcrossing, 101, 135 Outcrossing potential, 142 Oxidative stress, 236 Oxylipins, 301 P p21-activated protein kinases, 129 Paracrine pheromone signalling, 132 Parasitism biotrophic fusion parasitism, 303 Pathogenicity, 223–225 generation of diversity via RIP, 225 pathogenicity effector genes, 224 Pathogenicity factors, 236 Pathway basic integrated, 299 primary, 299 secondary, 299 special, 299 Pathway conservation, 299 Paxilline, 240 PdCLA4, 138 PdSTE3.1, 138 PdSTE3.2, 138 PdSTE3.3, 138 Penicillin biosynthesis, 314 regulation, 314 Peramine, 238 Perithecia, 91 Ph, 106 Phagocytosis, 244 Phagotrophy, 37, 45 Phenoloxidase, 242 Pheromone-like peptides, 139–141 Pheromone-pheromone receptor system, 106 paralogous groups of genes, 137 Pheromone precursor genes non-mating type, 142 SrRHA1, 113 SrRHA2, 113 SrRHA3, 113 Pheromone precursors, 102 processing, 107 Pheromone receptor constitutive, 142 ligand, 107 non-mating type, 142 Ste2, 116 Ste3, 106, 116 Pheromone receptor gene, 115, 141 SrSTE3, 113 Pheromone-receptor-like protein Cpr2, 133 Pheromone receptors, 75–86, 89–91 B mating type specific, 140 G protein-coupled receptors (GPCRs), 76

Subject Index G protein-coupled 7-transmembrane, 102, 107 non-mating type, 139 non-mating type specific, 140 STE2, 77, 85 STE3, 77, 85 Pheromone response pathway, 132 B mating type, 141 Pheromone-response signalling pathway, 120 Pheromones, 74–86, 89–91, 101 compatible, 107 H-type, 84 lipid-modified, 106 lipopeptide, 82 non-mating type, 143 non-self, 107 peptide, 81 and pheromone receptor genes, 142 self, 107 Phylogeny of Opisthokonta, 17 Phytohormone, 239 Pink basidiomycete yeasts, 42 p21-kinase, 138 Plasticity genetic, 298 Polarized cell growth, 129 Polyketide, 62, 67, 232 Polyketide synthases (PKS), 232, 233, 242, 320 Polysaccharides, 294 Post-translational modification, 75, 298 P/R, 135, 142 genes, 108 loci, 119, 139 pra1, 118, 120, 121, 126 receptors, 118 pra2, 118 receptors, 118 pra3, 118, 126 receptor, 118 Prespore, 60 Prestalk, 60, 67 Prf1, 141 Primary metabolism, 294 P/R locus, 106, 108, 126, 130, 131, 133, 142, 145 biallelic, 134 multi-allelism, 134 triallelic, 134 pr-MATA1, 114 pr-MATA2, 114 Propionate, 232 Protein, domain architecture, 296 Protein kinase (PKA), 61–64, 67 Proterospongia colonies, 12, 14 PRT1, 131 Pseudo-bipolar, 106, 124 Pseudo-bipolarity, 112, 122, 145 Pseudogene, 261 Pseudo-heterodimer, 141 Pseudo-homothallic, 97 Pseudothecium, 188 Pycnidia, 114 Pycniospores, 114

Q Quorum sensing, 143 R Radiation damage, 244 Radioactivity, 243 Radionuclides, 244 Radiotropism, 244 Rare biosphere, 47 Rearrangement of gene fragments, 299 Receptor, 294 agonists, 143 non-self, 108 pheromone, 108 self, 108 Receptor genes non-mating type, 139 Recognition process, 143 Recombination, 235, 308 mating type genes, 111 parasexual, 303 purifying, 122 repression, 114 sexual, 303 suppressed, 111, 120 suppression, 130 Recombinational activators, 130 Red yeast, 112 RegA, 62, 63 Repeat induced point (RIP), 121 Repeat-induced point mutation, 222 Repetitive DNA, 221–222 Reproduction bipolar, 146 homothallic, 101, 146 inbreeding-homothallic, 143 isogamous, 101 sexual, 97, 101, 108 tetrapolar, 146 unipolar, 146 unisexual, 101, 132, 143 unisexual mode, 146 Reproduction (apomixis), 101 Reproduction (heterothallism), 97 Retinoic acid, 302 rga2, 117 RHA1, 109 RHA2, 109 RHA3, 109 RHA2.A2, 109 Rhodotorucine A1, 109 Rhodotorucine A2, 109 RibL6, 122 RibL18ae, 122 RNAPOL, 122 Root sphere, 295 Rozella, 49 RPL39, 131 RPO41, 129, 131 RtSTE3.A1, 109 RtSTE20.A2, 109 RUM1, 129

343

344 S Same-sex mating, 129, 132, 133 ScBar3/Scbar3, 138 ScBbr2/Scbbr2, 138 Scbrl1, 138 ScBrl2/Scbrl2, 138 ScBrl3/Scbrl3, 138 Scbrl4, 138 Scytalone dehydratase, 242 SDF2, 63 Secondary metabolism, 294 Secondary metabolites, 231, 293 Selection, 298 balancing, 311 Self-compatibility constitutive, 141 Self-fertile, 97, 101, 112 Self-fertility, 101, 132, 144 Selfing, 101 Self-sterile, 141 454 Sequencing, 47–48, 50 Serine/threonine kinase, 120 Sex chromosomes, 108, 109, 114, 121, 142, 145 Sex determinants, 108 Sex determination bipolar, 122 tetrapolar, 122 Sexual cycle, 101 Sexual development, 103 Sexual reproduction, 109 Siderophore, 238 Signalling, 59, 61–62, 66, 67 autocrine, 129 cAMP, 59, 65, 67 paracrine, 129 Signal sequence, 75, 81 Signal transduction cascade, pheromone-activated, 108 Silent gene, 295 Sirodesmin, 238 Sister-group to Opisthokonta, 17 SjSTE3.A1, 112 Slug, 60 Soil, 295, 310 Soil clone group 1, 48 Special metabolite, 293, 294 description, 295 Species concepts, 164, 176 herbarium specimens, 179 intraspecific variability, 179 molecular characterisation, 177, 178 morphospecies, 167 species descriptions, 179 spore ontogeny, 165 SSU rDNA, 178, 179 Species discrimination, 143 Species recognition ITS-region, 179 ITS region variability, 179 LSU rDNA, 179 PCR primers, 179 Spermatiun, 86 Spo11, 132

Subject Index SPO14, 129 Spores, meiotic, 97 Sporidia, 114 Sporulation, 129 asexual, 141 repression, 141 SrMfa1.2, 119 SrMfa1.3, 119 SrMfa2.1, 119, 120 SrMfa2.3, 119 SrMfa3.1, 119, 120 SrMfa3.2, 119 SrSTE20, 113 SsRHA1, 109 SsRHA2, 109 SsRHA3, 109 SsSTE3, 109, 111 SsSTE3.A1, 109, 111 SsSTE3.A2, 109, 111 STE2, 116 Ste2, 144 Ste3, 129, 144 STE3, 106, 129, 131, 133 Ste6, 133 STE11, 120, 129–131, 133 STE12, 129, 130, 133 STE20, 122, 129–131, 133 Ste23, 107 Ste24, 107 STE3a, 129 STE3a, 129 STE11a, 129 STE11a, 129, 132 STE12a, 129 STE12a, 129, 132 STE20a, 129 STE20a, 129 Ste12a transcription factor, 132 Ste12-like transcription factors, 129 STE20-like gene (SsSTE20), 109 Ste20-like p21-activated kinase (PAK), 109 Sterigmatocystin, 235, 312 Subtelomeric regions, 308 Subunit ribosomal RNA, 39 Sugar reduction, 300 Switching, mating type, 101 Sxi1, 127 Sxi2, 127 SXI1a, 127 Sxi1a, 133 Sxi2a, 133 SXI2a, 127 SXI1 and SXI2, 131 Symbiosis, 163 Synonymous substitution rates, 284 Synteny, 143, 214, 296 mesosynteny, 217 T Taxonomy, 165 Teliospores, 112, 114 Telomere, 235, 308, 313

Subject Index Terpenes, 232, 239 Terpene synthases, 318 Terpenoids, 239, 294 Terrestrial ecosystems, 180 Tetrapolar modified, 116 Tetrapolar heterothallic species, 102 Tetrapolarity, 112, 122, 125, 142, 145 Thiolation, 237 TmSTE3, 134 TmSTE11, 134 TmSTE12, 134 TmSTE20, 134 Transcription factor AreA, 318 Transcription factors, 312 a-domain, 115 HD1-HD2, 105, 108 HMG box, 115 heterodimeric, 105 homeodomain, 108, 115 mating-type specific, 115 Transposons, 308 Tremerogen A–10, 134 Tremerogen a–13, 134 Tremerogen A-I, 134 Tremerogen A–9291-I, 134 Tremerogens, 134 Tremorgens, 240 Trichothecene, 239, 240 Tricyclazole, 243 Trisporoid, 294, 302 sexual communication, 297 Trychogyne, 86, 91 T-toxin, 236

T–2 toxin, 240 Ttrichothecene 3-O-acetyltransferase, 310 Tup1, 129 Tyrosinase, 242 U UhMfa1, 120 UhMfa2, 120 UhPra1, 120 UhPra2, 120 Uncultured, 38, 41, 48–49 Unipolarity, facultative, 146 Unisexual mating, 132 Unisexual reproduction, 132 ura1, 143 URA5, 131 UV radiation, 243 V Vad1, 129 Velvet, 235 Virulence factors, 294 V4 SSU, 47 V6 SSU, 47 V9 SSU, 47 Y Yeast cells, haploid, 101 Yeast phase, 134 Z Zearalenone, 236 ZNF1, 131 Znf2, 132

345

Evolution of Fungi and Fungal-Like Organisms (The Mycota, 14)

Evolution of Fungi and Fungal-Like Organisms

Evolution and Emergence: Systems, Organisms, Persons

Lizards: Windows to the Evolution of Diversity (Organisms and Environments)

Industrial Applications (The Mycota)

The science and philosophy of the organisms

Gravity and the behavior of unicellular organisms

Fungal Genomics (The Mycota XIII)

The Diversity of Living Organisms

Human and Animal Relationships (The Mycota)

The diversity of living organisms

The classification of lower organisms

Chronobiology of Marine Organisms

Invisible Radiations of Organisms

Biology of Marine Fungi

Chemistry of Fungi

Fungi - Biology and Applications

Molecular Identification of Fungi

Outline classification of organisms

Chronobiology of Marine Organisms

The Fungi, Second Edition

Fungal Genomics (The Mycota XIII)

Genomics of Plants and Fungi (Mycology)

Fungi, Algae, and Protists

Fungi: Biology and Applications

Straminipilous fungi

Molecular identification of fungi

Pictorial Atlas of Soil and Seed Fungi: Morphologies of Cultured Fungi and Key to Species

Ainsworth and Bisby's Dictionary of Fungi

Evolution of Fungi and Fungal-Like Organisms (The Mycota, 14)