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

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. Ku¨es 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)

viii

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 KU¨ES, 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

xvi

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

xviii

List of Contributors

URSULA KU¨ES (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

List of Contributors

xix

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

.

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

4

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

The Protistan Origins of Animals and Fungi

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

6

Martin Carr and Sandra L. Baldauf

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

The Protistan Origins of Animals and Fungi

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

7

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

8

Martin Carr and Sandra L. Baldauf

(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,

The Protistan Origins of Animals and Fungi

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 (Jime´nez-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 (Jime´nez-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; Jime´nez-Guri et al. 2007; Wallberg et al. 2004;). The monophyly of Porifera has been questioned in a number of studies (Borchiellini et al. 2001;

9

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 (Jime´nez-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; Jime´nez-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).

10

Martin Carr and Sandra L. Baldauf

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).

The Protistan Origins of Animals and Fungi

11

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 (<10) of round endospores are formed. The cell walls break open to release the endospores, which are then released into the external environment (Baker et al. 1999). As is the case with many ichthyosporeans, the complete life cycle has not been documented, so it is currently unknown if motile stages are produced or if dispersal is passive. Eccrinids produce two distinct dispersal cells. New arthropod hosts are infected by swallowing free-floating sporangiospores that are ovoid, uninucleate and thick-walled. Once the

Ichthyophonae mainly consists of parasitic organisms, although a small number of commensals and putative saprotrophs have also been described (Cafaro 2005; Kimura et al. 2002; Mendoza et al. 2002). The taxon is a phylogenetically distinct sister-group to the Rhinosporideacae (Fig. 1.3; Cafaro 2005) and exhibits a number of morphological differences (Mendoza et al. 2002). The Ichthyophonae and Rhinosporideacae differ in the morphology of their dispersal zoospores, with most described ichthyophonids producing amoeboid dispersal cells (Mendoza et al. 2002). Flagellated zoospores have only been reported in one ichthyophonid, Pseudoperkinsus tapetis (Ords and Figueras 1998), however this has yet to be verified. Hosts of the ichthyophonids are known to include arthropods, molluscs, fish, amphibians, reptiles and birds (Baker et al. 1999; Herman 1984; Mendoza et al. 2002; Rand 1994; Spanggaard et al. 1996). Not all ichthyophonids are believed to be parasitic; despite its name, the arthropodophilous

12

Martin Carr and Sandra L. Baldauf

parasite is established in its new host, it produces elongated, multinucleate, thin-walled sporangiospores (Cafaro 2005). Ichthyophonus hoferi has a particularly unusual morphology among known ichthyosporeans. It is the only known taxon for which tubular cristae have been reported (Ragan et al. 1998), but it is probably the dispersal cells that are its most distinctive character. In culture it has been shown that hyphae are formed at acidic pH, whereas at higher pH levels amoeboid cells are formed (Baker et al. 1999). Despite showing great variation in their ecology and morphology, there are a number of characters which are seen across Ichthyosporea. The formation of a cyst-like cell with a hard wall, along with production of multiple spores within the cyst, is common to many species. A parasitic/ commensal lifestyle is prevalent within the group. However, given their basal position in Holozoa and the fact that their hosts are mostly Metazoa, this life style is probably derived. Chitin within cell walls is only known in one ichthyosporean, Ichthyophonus hoferi, but given its widespread nature in the opisthokonts and the relative ease of losing complex traits compared to gaining them, there is a possibility that the ancestral ichthyosporean also had a chitin-containing cell wall.

VII. Choanoflagellida The choanoflagellates are an almost ubiquitously distributed group of aquatic protists. They play a profoundly important role in aquatic ecosystems, as they are one of the predominant predators of bacteria (King 2005; Leadbeater et al. 2009). There are approximately 250 described species of choanoflagellate (Carr et al. 2008a) present in either freshwater or the marine environment. Recent molecular phylogenies suggest that choanoflagellates may have only invaded freshwater from the marine environment on a single occasion (Carr et al. 2008a). However, most of the ~50 described freshwater species remain to be sampled. All described choanoflagellates possess a very similar looking protoplast (Karpov and Leadbeater 1997) and their mitochondrial cristae are uniformly flat (Hibberd 1975; Maldonado 2004). Cells are typically spherical or ovoid, with a posterior

flagellum surrounded by a collar of actin-based microvilli. The beating of the flagellum draws water through the collar; bacterial cells and other food particles adhere to the surface of the collar and are transported along the cell surface to the cell body where they are ingested (Boenigk and Arndt 2000). In unrestrained protoplasts, the beating flagellum causes a locomotory force which reduces the efficiency of water flow through the collar (Pettitt et al. 2002). Therefore many species produce a peduncle in order to attach the protoplast to a substratum. The peduncle is not an extension of the cell itself, but is formed from an external coat (periplast) that covers the anterior part of the cell. All choanoflagellate cells posses a periplast. This takes one of three major forms and is the basis for the traditional division of the group into three families (Codosigidae, Salpingoecidae, Acanthoecidae). However, molecular phylogeny has now shown that periplast morphology is an unstable character among two of the families, which are no longer valid. Instead, using molecular data, Carr et al. (2008a) identify three robust clades of choanoflagellate, one of which corresponds to the traditional Acanthoecidae and two of which are a mixture of traditional codosigid and salpingoecid taxa. This indicates that the choanoflagellates require considerable taxonomic revision, as well as the molecular characterization of the remaining >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.

The Protistan Origins of Animals and Fungi

13

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.

14

Martin Carr and Sandra L. Baldauf

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

The Protistan Origins of Animals and Fungi

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.

16

Martin Carr and Sandra L. Baldauf

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 mm) with 20 evenly spaced, unbranched microvilli radiating symmetrically around the protoplast (Patterson 1999; Adl et al. 2005). Although originally thought to have discoidal mitochondrial cristae (Cavalier-Smith 2000), Ministeria cristae are in fact flat (Patterson 1999). Cavalier-Smith and Chao (2003) noted that in both choanoflagellates and Ministeria, the microvilli are made up of microfilaments and lack a microtubule skeleton. Like choanoflagellates, it appears that Ministeria uses its microvilli to capture bacterial cells, which are then engulfed by phagocytosis (Cavalier-Smith and Chao 2003).

However, it is at yet unknown whether the microvilli in the two holozoan lineages are homologous or a case of convergent evolution. Although molecular phylogeny clearly shows that Ministeria is placed within Holozoa, its exact position within the group is not clear (CavalierSmith and Chao 2003; Ruiz-Trillo et al. 2006; Steenkamp et al. 2006). However, two recent studies strongly placed M vibrans together with Capsaspora owczarzaki, in the newly erected Filasterea (Burger et al. 2009; Shalchian-Tabrizi et al. 2008).

D. Corallochytrea As in the case of Capsaspora, Corallochytrea is only represented by a single species. Corallochytrium limacisporum is a marine protist isolated from coral reef lagoons in the Indian Ocean (Raghu-Kumar et al. 1987) and was initially described as a thraustochytrid. As with most holozoans, the cristae of mitochondria are flat (Cavalier-Smith and Allsopp 1996). Protoplasts are spherical and show considerable variation in size (4.5–20.0 mm in diameter). Mature cells undergo several rounds of binary division and up to 32 daughter cells are released through pores in the cell wall. Daughter cells have an elongated, amoeboid morphology and no flagellated cells have been observed in any stages of the lifecycle (Mendoza et al. 2002). C. limacisporum is a saprotroph and has not been found associated with any host organism (Mendoza et al. 2002). This nutritional mode allows it to be cultured axenically, which is useful for studying the organism in the laboratory, (Sumathi et al. 2006). Phylogenetic analyses robustly place C. limacisporum within Holozoa, but its exact placement has yet to be confirmed. Small-subunit RNA sequences weakly place C. limacisporum as a sister-group to the choanoflagellates (Cavalier-Smith and Chao 2003; Hertel et al. 2002), but multigene analyses place it with Ichthyosporea (Fig. 1.1; Carr et al. 2008a; Steenkamp et al. 2006). In all phylogenies C. limacisporum tends to be positioned on a long branch, which contributes to the difficulty in determining its true position. The sporangium formation followed by endospore release in C. limacisporum shows some similarity to that in the ichthyosporean R. seeberi, suggesting that they may be related. However endospores are released

The Protistan Origins of Animals and Fungi

from a single pore in R. seeberi, whereas there are multiple pores in Corallochytrium; in addition endospores are amoeboid in Corallochytrium and flagellated in Rhinosporidium.

IX. Conclusions A. Phylogeny of Opisthokonta As we have shown in this chapter, a number of the major branches within Opisthokonta have now been resolved (see Fig. 1.1). However resolution of the deeper branches is still heavily dependent on data from just a few taxa. There are also possibly whole groups that have yet to be characterized in molecular terms, such as the Aphelidea, which are so far placed into Opisthokonta based on morphological and ecological characters alone. In general, simply sequencing more taxa should be very useful by acting to break up long branches within the group. Within Holozoa we are now confident that the choanoflagellates are the sister-group to Metazoa. The Ichthyosporea may be the sister group to this metazoan + choanoflagellate clade, depending on the phylogenetic positions of various incertae sedis protists. Within Holofungi, the Nucleariida appear to be the deepest branch, with the possible exception of the Microsporidia. However, despite the publication of a number of multigene phylogenies of deep fungal branches (James et al. 2006a, b; Liu et al. 2006; White et al. 2006), none as yet include both microsporidians and nucleariids.

B. Sister-Group to Opisthokonta One of the outstanding questions in opisthokont evolution is the identity of their sister taxon. Large multigene phylogenies support the supergroup Amoebozoa as the closest group to opisthokonts (Burki et al. 2007; Hackett et al. 2007; Hampl et al. 2009). However, there is an increasing body of evidence to suggest that Apusozoa may in fact be a closer relation. Apusozoa are gliding soil zooflagellates which possess an anterior locomotory flagellum and a posterior feeding flagellum (Cavalier-Smith and Chao 1995). Their phylogenetic position has long been enigmatic, having previously been placed in Rhizaria (CavalierSmith 2002) or Excavata (Cavalier-Smith 2003),

17

whilst both Adl et al. (2005) and Bass et al. (2005) considered Apusozoa as Eukaryota incertae sedis. Several recently multigene studies tentatively place Apusozoa as a sister-group to Opisthokonta (Kim et al. 2006; Moreira et al. 2007; Ruiz-Trillo et al. 2007). Although each of these studies used only a single species, these were different species. Therefore, it seems less likely that the observed positioning could be due to stochastic effects such as long branch attraction. A genome project is currently underway for the apusozoan Amastigomonas sp. at the Broad Institute, Massachusetts.

C. Determining Opisthokont Diversity Like many, possibly even most protists, the true diversity of opisthokont protists is probably largely unknown. This is highlighted by the presence of Capsaspora owczarzaki, Corallochytrium limacisporum and Ministeria vibrans on long isolated branches, suggesting that each could represent a substantial unexplored lineage. Related to this is the observation that most ichthyosporeans are parasites, normally of either man or animals associated with man (such as farmed fish). This could reflect a bias in observed nutrition mode due to the fact that parasites associated with human activities are more likely to be discovered than unobtrusive saprotrophs such C. limacisporum. PCR amplification of rDNA from total environmental DNA has begun to highlight major gaps in our knowledge of eukaryotic diversity. Opisthokont sequences from undescribed taxa have been identified within such environmental datasets (Bass et al. 2007; Chen et al. 2008; Lara et al. 2010; Lepe`re et al. 2008; Takashita et al. 2007) and are yielding new insights into the diversity of both unicellular and multicellular opisthokonts.

D. Reconstructing Opisthokont Evolution and the Origins of Multicellularity The origins of multicellularity in Metazoa and Fungi are longstanding evolutionary questions of broad and deep interest. Studying their sistergroups, Nucleariida and Choanoflagellida, is key to understanding early events in the evolution of this complexity. To this end, it is important to examine a taxonomically broad set of species, to determine which characteristics are likely to

18

Martin Carr and Sandra L. Baldauf

have been present in their last common ancestor, versus which are species specific novelties. Fewer than ten nucleariid species have been described and one nucleariid genome sequencing project (Nuclearia simplex) is in progress (Broad Institute, Massachusetts). Over 200 species of choanoflagellate have been described and there are three choanoflagellate whole genome projects (Monosiga brevicollis, Monosiga ovata, Proterospongia sp. ATCC culture 50818). In addition to those species, there is one nucleariid (Genome Canada PEP Database; http://megasun.bch.umontreal.ca/ pepdb/pep.html) EST project (http://tbestdb.bcm. umontreal.ca/searches/login.php). The greater volume of choanoflagellate data means that comparative genomics studies involving choanoflagellates are possible (Carr et al. 2008b, 2010; King and Carroll 2001; King et al. 2003; Sez et al. 2009), whilst similar studies have yet to be performed on the nucleariids. It will be of great interest to see if nucleariids, as well as the unicellular basal fungi, possess the genes with functions necessary for multicellularity in fungi (albeit employed differently), or whether these genes arose later in the evolution of Fungi. Within the next few years a large body of genome data will become available for opisthokont protists, as well as for other eukaryotic groups. This should allow more accurate determination of the closest opisthokont sister taxon. With this information, it will also be possible to begin to reconstruct ancestral genomes of the group. Comparative genomics across opisthokont lineages will make it possible to estimate the complexity and gene families present in the last common ancestor of the group. In addition, both morphological and ecological reconstructions are possible when sister relationships are known (Carr et al. 2008a). Comparative gene expression and function studies may provide insights into the original function of such genes and how the genes have been differently employed in different opisthokont lineages. Thus we expect to see major changes in our understanding of early opisthokont evolution in the next few years.

References Abedin M and King N (2008) The premetazoan ancestry of cadherins. Science 319:946–948 Adl SM, Simpson AGB, Farmer MA, Andersen RA, Anderson OR, et al. (2005) The new higher level of

classification of eukaryotes with emphasis on the taxonomy of protists. J Euk Microbiol 52:399–451 Aguinaldo AA, Turbeville JM, Linford LS, Rivera MC, Garey JR, Raff RA, Lake JA (1997) Evidence for a clade of nematodes, arthropods and other moulting animals. Nature 387:489–493 Ahluwalia KBN, Maheshwari N, Deka RC (1997) Rhinosporidiosis: a study that resolves etiologic controversies. Am J Rhinol 11:479–483 Amaral-Zettler LA, Nerad TA, O’Kelly J, Sogin ML (2001) The nucleariid amoebae: more protists at the animal– fungal boundary. J Euk Microbiol 48:293–297 Amaral-Zettler LA, Go´mez F, Zettler E, Keenan BG, Amils R, Sogin ML (2002) Eukaryotic diversity in Spain’s River of Fire. Nature 417:417 Anderson CL, Canning EU, Okamura B (1998) A triploblasts origin for Myxozoa? Nature 392:346–347 Anderson FE, Co´rdoba AJ, Thollesson M (2004) Bilaterian phylogeny based on analyses of a region of the sodium–potassium ATPase a-subunit gene. J Mol Evol 58:252–268 Arkush, KD, Frasca S Jr, Hedrick RP (1998) Pathology associated with the rosette agent, a systemic protist infecting salmonid fishes. J Aquat Anim Health 10: 1–11 Baker GC, Beebee TJ, Ragan MA (1999) Prototheca richardsi, a pathogen of anuran larvae, is related to a clade of protistan parasites near the animal–fungal divergence. Microbiology 145:1777–1784 Baldauf SL, Palmer JD (1993) Animals and fungi are each others closest relatives: congruent evidence from multiple proteins. Proc Natl Acad Sci USA 90:11558–11562 Bass D, Howe A, Brown N, Barton H, Demidova M, Michelle H, Li L, Sanders H, Watkinson SC, Willcock S, Richards TA (2007) Yeast forms dominant fungal diversity in the deep oceans. Proc R Soc B 274: 3069–3077 Bass D, Moreira D, Lo´pez-Garcı´a P, Polet S, Chao EE, von der Heyden S, Pawlowski J, Cavalier-Smith T (2005) Polyubiquitin insertions and the phylogeny of Cercozoa and Rhizaria. Protist 156:149–161 Baurain D, Brinkmann H, Philippe H (2007) Lack of resolution in the animal phylogeny: Closely spaced cladogeneses or undetected systematic errors? Mol Biol Evol 24:6–9 Benny GL, O’Donnell K (2000) Amoebidium parasiticum is a protozoan, not a Trichomycete. Mycologica 92: 1133–1137 Boenigk J, Arndt H (2000) Comparative studies on the feeding behavior of two heterotrophic nanoflagellates: the filter-feeding choanoflagellate Monosiga ovata and the raptorial-feeding kinetoplastid Rhynchomonas nasuta. Aqua Micro Eco 22:243–249 Borchiellini C, Manuel M, Alivon E, Boury-Esnault N, Vacelet J, Le Parco Y (2001) Sponge paraphyly and the origin of Metazoa. J Evol Biol 14:171–179 Bourlat SJ, Juliusdottir T, Lowe CJ, Freeman R, Aronowicz J, Kirschner M, Lander ES, Thorndyke M, Nakano H, Kohn AB, Heyland A, Moroz LL, Copley RR, Telford MJ (2006) Deuterostome phylogeny reveals monophyletic chordates and the new phylum Xenoturbellida. Nature 444:85–88

The Protistan Origins of Animals and Fungi Bourlat SJ, Nielsen C, Economou AD, Telford M (2008) Testing the new animal phylogeny: a phylum level molecular analysis of the animal kingdom. Mol Phylogenet Evol 49:23–31 Bourrelly P (1968) Les algues d’eau douce. Tome II: Les algues jaunes et brunes. Paris: Boube´e et Cie Bozarth RF (1972) Mycoviruses: a new dimension in microbiology. Environ Health Perspect 2:23–39 Brown MW, Spiegel FW, Silberman JD (2009) Phylogeny of the “forgotten” cellular slime mold, Fonticula alba, reveals a key evolutionary branch within Opisthokonta. Mol Biol Evol 26:2699–2709 Brown TA, Waring RB, Scazzocchio C, Davies RW (1985) The Aspergillus nidulans mitochondrial genome. Curr Genet 9:113–117 Bullerwell CE, Leigh J, Forget L, Lang BF (2003) A comparison of three fission yeast mitochondrial genomes. Nucleic Acids Research 31:759–768 Burger G, Forget L, Zhu Y, Gray MW, Lang BF (2003) Unique mitochondrial genome architecture in unicellular relatives of animals. Proc Natl Acad Sci USA 100: 892–897 Burger G, Yan Y, Javadi P, Lang BF (2009) Group I-intron trans-splicing and mRNA editing in the mitochondria of placozoan animals. Trends Genet 25:381–386 ˚, Burki F, Shalchian-Tabrizi K, Minge M, Skjæveland A Nikolaev SI, Jakobsen KS, Pawlowski J (2007) Phylogenomics reshuffles the eukaryotic supergroups. PLoS ONE 8:e790 Burki F, Shalchian-Tabrizi K, Pawlowski J (2008) Phylogenomics reveals a new ‘megagroup’ including most photosynthetic eukaryotes. Biol Lett 23:366–369 Buss KW (1987) The evolution of individuality. Princeton University Press, Princeton. 203 pp Cafaro MJ (2005) Eccrinales (Trichomycetes) are not fungi, but a clade of protists at the early divergence of animals and fungi. Mol Phy Evol 35:21–34 Cann JP (1986) The feeding behavior and structure of Nuclearia delicatula (Filosea: Aconchulinida) J Euk Microbiol 33:392–396 Cardoso MAG, Tambor JHM, Nobrega FG (2007) The mitochondrial genome from the thermal dimorphic fungus Paracoccidioides brasiliensis. Yeast 24: 607–616 Carr M, Leadbeater BSC, Baldauf SL (2010). Conserved meiotic genes point to sex in the choanoflagellates. J Euk Microbiol 57:56–62 Carr M, Leadbeater BSC, Hassan R, Nelson M, Baldauf SL (2008a). Molecular phylogeny of choanoflagellates, the sister group to Metazoa. Proc Natl Acad Sci USA 105:16641–16646 Carr M, Nelson M, Leadbeater BSC, Baldauf SL (2008b) Three families of LTR retrotransposons are present in the genome of the choanoflagellate Monosiga brevicollis. Protist 159:579–590 Cavalier-Smith T (1987) The origin of fungi and pseudofungi in: Rayner ADM, Brasierand M and Moore D (eds.) Evolutionary biology of fungi, pp. 339–353, Cambridge: Cambridge University Press Cavalier-Smith T (1993) Kingdom Protozoa and its 18 phyla. Microbiological Reviews 57:953–994 Cavalier-Smith T (1998) A revised six kingdom system of life. Biol Rev 73:203–266

19

Cavalier-Smith T (2000) Flagellate megaevolution: The basis for eukaryote diversification. In: Green J.R., Leadbeater, B.S.C. (eds) The flagellates. Taylor and Francis, London, pp 361–390 Cavalier-Smith T (2001) What are fungi?, in J.W. McLaughlin, J.W. McLaughlin and P.A. Lemke (eds.) The Mycota VII Part A, Systematics and evolution, pp. 3–37, Berlin Heidelberg: Springer Verlag Cavalier-Smith T (2002) The phagotrophic origin of eukaryotes and phylogenetic classification of Protozoa. Int J Syst Evol Microbiol 52:297–354 Cavalier-Smith T (2003) Protist phylogeny and the highlevel classification of Protozoa. Europ J Protistol 39: 338–349 Cavalier-Smith T (2009) Megaphylogeny, cell body plans, adaptive zones: causes and timing of eukaryotic basal radiations. J Eukaryot Microbiol 56:26–33 Cavalier-Smith T, Allsopp MTEP (1996) Corallochytrium, an enigmatic non-flagellate protozoan related to choanoflagellates. Europ J Protistol 32:306–310 Cavalier-Smith, T, Chao EEY (1995) The opalozoan Apusomonas is related to the common ancestor of animals, fungi and choanoflagellates. Proc R Soc B 261:1 Cavalier-Smith, T, Chao EEY (1997) Sarcomonad ribosomal RNA sequences, rhizopod phylogeny, and the origin of euglyphid amoebae. Arch Protistenkd 147:227–236 Cavalier-Smith, T, Chao EEY (2003) Phylogeny of Choanozoa, Apusozoa, and other protozoa and early eukaryotic megaevolution. J Mol Evol 56:540–563 Cavalier-Smith, T, Chao EEY, Oates B (2004) Molecular phylogeny of Amoebozoa and the evolutionary significance of the unikont Phalansterium. Europ J Protistol 40:21–48 Chadefaud M (1960) Les ve´ge´taux non vasulaires (Cryptogamie). Traite´ de botanique syste´matique, vol. 3. Chadeauf and Emberger. pp. 1018. Paris: Massor et Cie Chen M, Chen F, Yu Y, Ji J, Kong F (2008) Genetic diversity of eukaryotic microorganisms in Lake Taihu, a large shallow subtropical lake in China. Microb Ecol 56: 572–583 Cummings DJ, Mcnally KL, Domenico JM, Matsuura ET (1990) The complete DNA sequence of the mitochondrial genome of Podospora anserina. Curr Genet 17: 375–402 Deasey MC (1982) Aspects of sorogenesis in the cellular slime mold Fonticula alba. PhD thesis, University of North Carolina, Chapel Hill Dellaporta SL, Xu A, Sagasser S, Jakob W, Moreno MA, Buss LW, Schierwater B (2006) Mitochondrial genome of Trichoplax adhaerens support Placozoa as the basal lower metazoan phylum. Proc Natl Acad Sci USA 103:8751–8756 Dunn CW, Hejnol A, Matus DQ, Pang K, Browne WE, et al. (2008) Broad phylogenomic sampling improves resolution of the animal tree of life. Nature 452:745–749 Dykov I, Veverkov M, Fiala I, Machckov B, Peckov H (2003) Nuclearia pattersoni sp. n. (Filosea), a new species of amphizoic amoeba isolated from gills of roach (Rutilus rutilus), and its rickettsial endosymbiont. Folia Parasit 50:161–170 Erpenbeck D, Voigt O, Adamski M, Adamska M, Hooper JNA, Wo¨rheide G, Degnan BM (2007) Mitochondrial

20

Martin Carr and Sandra L. Baldauf

diversity of early-branching Metazoa is revealed by the complete mt genome of a Haplosclerid Demosponge. Mol Biol Evol 24:19–22 Farabee MJ (2002). Biological diversity: animals I, II and III, in: Online biology book. Available at http://www. emc.maricopa.edu/faculty/farabee/BIOBK/ BioBookTOC.HTML Feldman SH, Wimsatt JH, Green DE (2005) Phylogenetic classification of the frog pathogen Amphibiothecum (Dermosporidium) penneri based on small ribosomal subunit sequencing. J Wildl Dis 41:701–706 Gill EE, Fast NM (2006) Assessing the microsporidia–fungi relationship: combined phylogenetic analysis of eight genes. Gene 375:103–109 Gromov BV (2000) Algal parasites of the genera Aphelidium, Amoeboaphelidium, and Pseudoaphelidium from the Cienkovski’s “Monadinea” group as representatives of a new class. Zool Zhurn 79:517–525 Hackett JD, Yoon HS, Li S, Reyes-Prieto A, Ru¨mmele SE, Bhattacharya D (2007) Phylogenomic analysis supports the monophyly of cryptophytes and haptophytes and the association of Rhizaria with chromoalveolates. Mol Biol Evol 24:1702–1713 Haeckel E (1874) Die Gastraea-Theorie, die phylogenetischen Classification des Thierreichs und die Homologie der Keimbla¨tter. Z Naturwiss 8:1–55 Haen KM, Lang BF, Pomponi SA, Lavrov DV (2007) Glass sponges and bilaterian animals share derived mitochondrial genomic features: A common ancestry or parallel evolution? Mol Biol Evol 24:1518–1527 Halanych KM (2004) The new view of animal phylogeny. Annu Rev Ecol Evol Syst 35:229–256 Halanych KM, Bacheller JD, Aguinaldo AMA, Liva SM, Hillis DM, Lake JA (1995) Evidence from 18S ribosomal DNA that the lophophorates are protostome animals. Science 267:1641–1643 Hampl V, Hug L, Leigh JW, Dacks JB, Lang BF, Simpson AGB and Roger AJ (2009) Phylogenomic analyses support the monophyly of Excavata and resolve relationships among eukaryotic “supergroups”. Proc Natl Acad Sci USA 106:3859–3864 Harrell LW, Elston RA, Scott TM, Wilkinson MT (1986) A significant new systemic disease of net-pen reared chinook salmon (Oncorhyncus tshawytscha) brook stock. Aquaculture 55:249–262 Hejnol A, Martindale MQ (2008) Acoel development indicates the independent evolution of the bilaterian mouth and anus. Nature 456:382–386 Hendy MD, Penny D (1989) A framework for the quantitative study of evolutionary trees. Syst Zool 38: 297–309 Herman RL (1984) Ichthyophonus-like infection in newts (Notophthalmus viridescens Rafinesque) J Wildlife Dis 20:55–56 Herr RA, Ajello L, Taylor JW, Arseculeratne SN, Mendoza L (1999) Phylogenetic analysis of Rhinosporidium seeberis 18S small-subunit ribosomal DNA groups this pathogen among members of the protoctistan Mesomycetozoa clade. J Clin Microbiol 37:2750–2754 Hertel LA, Bayne C, Loker ES (2002) The symbiont Capsaspora owczarzaki, nov. gen. sp., isolated from three strains of the pulmonate snail Biomphalaria glabrata

is related to members of the Mesomycetozoa. Int J Parasit 32:1183–1191 Hibberd DJ (1975) Observations on the ultrastructure of the choanoflagellate Codosiga botrytis (ehr.) Saville– Kent with special reference to the flagellar apparatus. J Cell Sci 17:191–219 Hibbett DS, Binder M, Bischoff JF, Blackwell M, Cannon PF, et al. (2007) A higher-level phylogenetic classification of the Fungi. Mycol Res 111:509–547 Hyman LH (1940) The Invertebrates, vol. 1, New York: McGraw-Hill James TY, Kauff F, Schoch L, Matheny PB, Hofstetter V, et al. (2006a) Reconstructing the early evolution of fungi using a six-gene phylogeny. Nature 443:818–822 James TY, Letcher PM, Longcore JE, Mozley-Standridge SE, Porter D, Powell MJ, Grffith GW, Vilgalys R (2006b) A molecular phylogeny of the flagellated fungi (Chytridiomycota) and description of a new phylum (Blastocladiomycota) Mycologia 98:860–871 James-Clark H (1866) Note on the infusoria flagellata and the spongiae ciliatae. Am J Sci 1:113–114 James-Clark H (1868) On the spongiae ciliatae as infusoria flagellata; or observations on the structure, animality and relationship of Leucosolenia botryoides, Bowerbank. Ann Mag Nat Hist 1:133–142 Jay JM, Pohley WJ (1981) Dermosporidium penneri sp. n. from the skin of the American toad, Bufo americanus (Amphibia: Bufonidae). J Parasitology 67:108–110 Jime´nez-Guri E, Philippe H, Okamura B, Holland PWH (2007) Buddenbrockia is a cnidarian worm. Science 317:116–118 Jones MD, Forn I, Gadelha C, Egan MJ, Bass D, Massana R, Richards TA (2011) Discovery of novel intermediate forms redefines the fungal tree of life. Nature (in press) Karpov SA and Leadbeater BSC (1997) Cytoskeleton structure and composition in choanoflagellates. Europ J Protistol 33:323–334 Keeling PJ (2003) Congruent evidence from a-tubulin and b-tubulin gene phylogenies for a zygomycete origin of microsporidia. Fungal Genet Biol 38:298–309 Kent SW (1880) A manual of the Infusoria, vols 1–3 (1880– 1882). Kent, London Kim E, Simpson AGB, Graham LE (2006) Evolutionary relationships of Apusomonads inferred from taxonrich analyses of 6 nuclear encoded genes. Mol Biol Evol 23:2455–2466 Kimura H, Harada K, Hara K, Tamaki A (2002) Enzymatic approach to fungal association with arthropod guts: a case study for the crustacean host, Nihonotrypaea harmandi, and its foregut fungus, Enteromyces callianassae. Mar Ecol 23:157–183 King N (2005) Choanoflagellates. Curr Biol 15:R113–R114 King N and Carroll SB (2001) A receptor tyrosine kinase from choanoflagellates: molecular insights into early animal evolution. Proc Natl Acad Sci USA 98: 15032–15037 King N, Hittinger CT, Carroll SB (2003) Evolution of key cell signaling and adhesion protein families predates animal origins. Science 301:361–363 King N, Westbrook MJ, Young SL, Kuo A, Abedin M, et al. (2008) The genome of the choanoflagellate Monosiga

The Protistan Origins of Animals and Fungi brevicollis and the origin of metazoans. Nature 451: 783–788 Kodner RB, Summons RE, Pearson A, King N, Knoll AH (2008) Sterols in a unicellular relative of the metazoans. Proc Natl Acad Sci USA 105:9897–9902 Kruger W (1894) Kurze Charakteristik einiget niederer Organismen in Saftlusse der Laubbaume. Hedwigia 33: 241–266 Kirk PM, Cannon PF, David JC, Stalpers JA (2001) Ainsworths & Bisbys dictionary of the fungi, 9th edn, Wallingford: CABI publishing Lang BF, Gray MW, Burger G (1999) Mitochondrial genome evolution and the origin of eukaryotes. Annu Rev Genet 33:351–397 Lang BF, O’Kelly C, Nerad T, Gray MW, Burger G (2002) The closest unicellular relatives of Animals. Current Biology 12:1773–1778 Lara E, Moreira D, Lo´pez-Garcı´a P (2010) The environmental clade LKM11 and Rozella form the deepest branching clade of fungi. Protist 161:116–121 Lavrov DV, Forget L, Kelly M, Lang BF (2005) Mitochondrial genomes of two demosponges provide insights into an early stage of animal evolution. Mol Biol Evol 22:1231–1239 Leadbeater BSC (1979) Developmental studies in the loricate choanoflagellate Stephanoeca diplocostata. II. Cell division and lorica assembly. Protoplasma 98:311–328 Leadbeater BSC, Hassan R, Nelson M, Carr M, Baldauf SL (2008) A new genus, Helgoeca gen. nov., for a nudiform choanoflagellate. Europ J Protistol 44:227–237 Leadbeater BSC, Morton C (1974) A microscopical study of a marine species of Codosiga James-Clark (Choanoflagellata) with special reference to the ingestion of bacteria. Biol J Linn Soc 6:337–347 Leadbeater BSC, Yu Q, Kent J, Stekel DJ (2009) Threedimensional images of choanoflagellate loricae. Proc. R. Soc. B 276:3–11 Lee J, Young JPW (2009) The mitochondrial genome sequence of the arbuscular mycorrhizal fungus Glomus intraradices isolate 494 and implications for the phylogenetic placement of Glomus. New Phytol. 183: 200–211 Lee SC, Corradi N, Byrnes III EJ, Torres-Martinez S, Dietrich FS, Keeling PJ, Heitman J (2008) Microsporidia evolved from ancestral sexual fungi. Current Biology 18:1675–1679 Lepe`re C, Domaizon I, Debroas D (2008) Unexpected importance of potential parasites in the composition of the freshwater small-eukaryote community. Appl Environ Microbiol 74:2940–2949 Lichtwardt RW (1986) The Trichomycetes: Fungal associates of arthropods. Springer-Verlag, New York pp 343 Liu YJ, Hodson MC, Hall BD (2006) Loss of the flagellum happened only once in the fungal lineage: phylogenetic structure of Kingdom Fungi inferred from RNA polymerase II subunit genes. BMC Evolutionary Biology 6:74 Liu Y, Leigh JW, Brinkmann H, Cushion MT, RodriguezEzpeleta N, Philippe H, Lang BF (2009) Phylogenomic analyses support the monophyly of Taphrinomycotina, including Schizosaccharomyces fission yeasts. Mol Biol Evol 26:27–34

21

Loytynoja A, Milikovitch MC (2001) Molecular phylogenetic analyses of the mitochondrial ADP–ATP carriers: the Plantae/Fungi/Metazoa trichotomy revisited. Proc Natl Acad Sci USA 98:10202–10207 Maldonado M (2004) Choanoflagellates, choanocytes, and animal multicellularity. Invert Bio 123:1–22 Mallatt JM, Garey JR, Schultz JW (2004) Ecdysozoan phylogeny and Bayesian inference: first use of nearly complete 28S and 18S rRNA gene sequences to classify the arthropods and their kin. Mol Phy Evol 31: 178–191 Manning G, Young SL, Miller WT, Zhai Y (2008) The protist, Monosiga brevicollis, has a tyrosine kinase signaling network more elaborate and diverse than found in any known metazoan. Proc Natl Acad Sci USA 105:9674–9679 Matus DQ, Pang K, Marlow H, Dunn CW, Thomsen GH, Martindale MQ (2006) Molecular evidence for deep evolutionary roots of bilaterality in animal development. Proc Natl Acad Sci USA 103:11195–11200 Medina M, Collins AG, Silberman JD, Sogin ML (2001) Evaluating hypotheses of basal animal phylogeny using complete sequences of large and small subunit rRNA. Proc Natl Acad Sci USA 98:9707–9712 Medina M, Collins AG, Taylor JW, Valentine JW, Lipps JH, Amaral-Zettler L, Sogin ML (2003) Phylogeny of Opisthokonta and the evolution of multicellularity and complexity in Fungi and Metazoa. Int J Astro 2: 203–211 Mendoza L, Ajello L, Taylor JW (2001) The taxonomic status of Lacazia loboi and Rhinosporidium seeberi has been finally resolved with the use of molecular tools. Rev Iberoam Micol 18:95–98 Mendoza L, Taylor JW, Ajello L (2002) The Class Mesomycetozoa: a heterogeneous group of microorganisms at the animal–fungal boundary. Annu Rev Microbiol 56:315–344 Minge MA, Silberman JD, Orr RJS, Cavalier-Smith T, Shal˚ , Jakobsen KS chian-Tabrizi K, Burki F, Skjæveland A (2009) Evolutionary position of breviate amoebae and the primary eukaryote divergence. Proc R Soc B 276: 597–604 Monteiro AS, Okamura B, Holland PWH (2002) Orphan worm finds a home: Buddenbrockia is a myxozoan. Mol Biol Evol 19:968–971 Moreira D, von der Heyden S, Bass D, Lo´pez-Garcı´a P, Chao EEY, Cavalier-Smith T (2007) Global eukaryote phylogeny: Combined small- and large-subunit ribosomal DNA trees support monophyly of Rhizaria, Retaria and Excavata. Mol Phylogenet Evol 44:255–266 Norris RE (1965) Neustonic marine Craspedomonadales (Choanoflagellata) from Washington and California. J Protozool 12:589–612 Ords MC, Figueras A (1998) In vitro culture of Perkinsus atlanticus, a parasite of the carpet shell clam Ruditapes decussates. Dis Aquat Org 33:129–136 Owczarzak A, Stibbs HH, Bayne CJ (1980) The destruction of Schistosoma mansoni mother sporocysts in vitro by amoebae isolated from Biomphalaria glabrata: an ultrastructural study. J Inver Path 35:26–33 Page FC (1987) The classification of ‘naked’ amoebae (Phylum Rhizopoda). Arch Protistenk 133:199–217

22

Martin Carr and Sandra L. Baldauf

Patron NJ, Inagaki Y, Keeling PJ (2007) Multiple gene phylogenies support the monophyly of cryptomonad and haptophyte host lineages. Current Biology 17: 887–891 Patterson DJ (1984) The genus Nuclearia (Sarcodina, Filosea): species composition and characteristics of the taxa. Arch Protistenk 128:127–139 Patterson DJ (1999) The diversity of eukaryotes. Am Nat 154(Suppl):S86–S124. Patterson DJ, Nygaard K, Steinberg G, Turley C (1993) Heterotrophic flagellates and other protists associated with organic detritus throughout the water column in the mid North Atlantic. J Mar Biol Assoc UK 73:67–95 Perry RP, Cheng TY, Freed JJ, Greenberg JR, Kelley DE, Tartof KD (1970) Evolution of the transcription unit of ribosomal RNA. Proc Natl Acad Sci USA 65: 609–616 Pettitt ME, Orme BAA, Blake JR, Leadbeater BSC (2002) The hydrodynamics of filter feeding in choanoflagellates. Europ J Protistol 38:313–332 Philippe H, Derelle R, Lopez P, Pick K, Borchiellini C, Boury-Esnault N, Vacelet J, Renard E, Houliston E, Que´innec E, Da Silva C, Wincker P, Le Guyader H, Leys S, Jackson DJ, Schreiber F, Erpenbeck D, Morgenstern B, Wo¨rheide G, Manuel M (2009) Phylogenomics revives traditional views on deep animal relationships. Curr Biol 19:706–712 Philippe H, Snell EA, Bapteste E, Lopez P, Holland PWH, Casane D (2004) Phylogenomics of eukaryotes: Impact of missing data on large alignments. Mol Biol Evol 21:1740–1752 Philippe H, Telford MJ (2006) Large-scale sequencing and the new animal phylogeny. Trends in Ecology and Evolution 21:614–620 Pirozynski KA, Malloch DW (1975) The origin of land plants: a matter of mycotrophism. Biosystems 5: 153–164 Powell MJ (1984) Fine structure of the unwalled thallus of Rozella polyphagi in its host Polyphagus euglenea. Mycologia 76:1039–1048 Ragan MA, Goggins CL, Cawthorn RJ, Cerenius L, Jamienson AVC, et al. (1998) A novel clade of protistan parasites near the animal–fungal divergence. Proc Natl Acad Sci USA 93:11907–11912 Raghu-Kumar S, Chandramohan D, Ramaiah N (1987) Contribution of the thraustochytrid Corallochytrium limacisporum Raghu-Kumar to microbial biomass in coral reef lagoons. Ind J Marine Sciences 16: 122–125 Rand TG (1994) An unusual form of Ichthyophonus hoferi from yellowtail flounder Limanda Ferruginea from the Nova Scotia shelf. Dis Aquat Org 18:21–28 Ronquist F, Huelsenbeck JP (2003) MrBayes 3: Bayesian phylogenetic inference under mixed models. Bioinformatics 19:1572–1574 Ruiz-Trillo I, Inagaki Y, Davis LA, Sperstad S, Landfald B, Roger AJ (2004) Capsaspora owczarzaki is an independent opisthokont lineage. Curr Biol 14:R946 Ruiz-Trillo I, Lane CE, Archibald JM, Roger AJ (2006) Insights into the evolutionary origin and genome architecture of the unicellular Opisthokonts Capsas-

pora owczarzaki and Sphaeroforma arctica. J Eukaryot Microbiol 53:379–384 Ruiz-Trillo I, Burger G, Holland PWH, King N, Lang BF, Roger AJ, Gray MW (2007) The origins of multicellularity: a multi-taxon genome initiative. Trends Genet 23:113–118 Ruiz-Trillo I, Roger AJ, Burger G, Gray MW, Lang BF (2008) A phylogenomic investigation into the origin of Metazoa. Mol Biol Evol 25:664–672 Sez AG, Lozano E, Zaldı´var-Rivero´n A (2009) Evolutionary history of Na,K-ATPases and their osmoregulatory role. Genetica 136:479–490 Schierwater B (2005) My favorite animal, Trichoplax adhaerens. Bioessays 27:1294–1302 Shalchian-Tabrizi K, Minge MA, Espelund M, Orr R, Ruden T, Jakobsen KS, Cavalier-Smith T (2008) Multigene phylogeny of Choanozoa and the origin of animals. PLoS One 3:e2098 Signorovitch AY, Buss LW, Dellaporta SL (2007) Comparative genomics of large mitochondria in placozoans. PLoS Genetics 3:44–50 Smothers JF, von Dohlen CD, Smith Jr LH, Spall RD (1994) Molecular evidence that the myxozoan protists are metazoans. Science 265:1719–1721 Spanggaard B, Skouboe P, Rossen L, Taylor JW (1996) Phylogeneti relationships of the intercellular fish pathogen Ichthyophonus hoferi, and fungi, choanoflagellates and the rosette agent. Mar Biol 126:109–115 Srivastava M, Begovic E, Chapman J, Putnam NH, Hellsten U, et al. (2008) The Trichoplax genome and the nature of placozoans. Nature 454:955–960 Stamatakis A, Ludwig T, Meier H (2005) RAxML-III: a fast program for maximum likelihood-based inference of large phylogenetic trees. Bioinformatics 21:456–463 Stechmann A, Cavalier-Smith T (2003) Phylogenetic analysis of eukaryotes using heat–shock protein Hsp90. J Mol Evol 57:408–419 Steenkamp ET, Wright J, Baldauf SL (2006) The protistan origins of animals and fungi. Mol Biol Evol 23:93–106 Stibbs HH, Owczarzak A, Bayne CJ, DeWan P (1979) Schistosome sporocyst-killing amoebae isolated from Biomphalaria glabrata. J Invert Pathol 33:159–170 Sumathi JC, Raghukumar S, Kasbekar DP, Raghukumar C (2006) Molecular evidence of fungal signatures in the marine protist Corallochytrium limacisporum and its implications in the evolution of animals and fungi. Protist 157:363–376 Tanabe Y, Watanabe MM, Sugiyama J (2005) Evolutionary relationships among basal fungi (Chytridiomycota and Zygomycota): Insights from molecular phylogenetics. J Gen Appl Microbiol 51:267–276 Takashita K Tsuchiya M, Kawato M, Oguri K, Kitazato H, Maruyama T (2007) Genetic diversity of microbial eukaryotes in anoxic sediment of the saline meromictic Lake Namako-ike (Japan): on the detection of anaerobic or anoxic-tolerant lineages of eukaryotes. Protist 158:51–64 Tehler A (1988) A cladistic outline of the Eumycota. Cladistics 4:227–277 Tehler A, Farris JS, Lipscomb DL and Ka¨llersjo¨ M (2000) Phylogenetic analyses of the fungi based on large rDNA data sets. Mycologia 92:459–474

The Protistan Origins of Animals and Fungi Telford MJ (2006) Animal phylogeny. Curr Biol 16:R981– R985 Trotter MJ, Whisler HC (1965) Chemical composition of the cell wall of Amoebidium parasiticum. Can J Bot 43:869–876 van Etten JL, Lane LC, Meints RH (1991) Viruses and viruslike particles of eukaryotic algae. Microbiol Rev 55:586–620 van Hannen EJ, Mooij W, van Agterveld MP, Gons HJ, van Laanbroek P (1999) Detritus-dependent development of the microbial community in an experimental system: quality analysis by denaturing gradient gel electrophoresis. Appl Env Micro 65:2478–2484 Voigt K, Wo¨stemeyer J (2001) Phylogeny and origin of 82 zygomycetes from all 54 genera of the Mucorales and Mortierellales based on combined analysis of actin and translation elongation factor EF-1a genes. Gene 270:113–120 Voigt O, Collins AG, Buchsbaum Pearse V, Pearse JS, Ender A, Hadrys H, Schierwater B (2004) Placozoa – no longer a phylum of one. Curr Biol 14: R944–R945 Wainright PO, Hinkle G, Sogin ML, Stickel SK (1993) Monophyletic origins of the Metazoa: an evolutionary link with fungi. Science 260:340–342

23

Wallberg A, Thollesson M, Farris JS, Jondelius U (2004) The phylogenetic position of the comb jellies (Ctenophora) and the importance of taxonomic sampling. Cladistics 20:558–578 Wang X, Lavrov DV (2007) Mitochondrial genome of the homoscleromorph Oscarella carmela (Porifera, Demospongiae) reveals unexpected complexity in the common ancestor of sponges and other animals. Mol Biol Evol 24:363–373 White MM, James TY, O’Donnell, Cafaro MJ, Tanabe Y, Sugiyama J (2006) Phylogeny of the Zygomycota based on nuclear ribosomal sequence data. Mycologia 98:872–884 Woo PC, Zhen H, Cai JJ, Yu J, Lau SK, Wang J, Teng JL, Wong SS, Tse RH, Chen R, Yang H, Liu B, Yuen KY (2003) The mitochondrial genome of the thermal dimorphic fungus Penicillium marneffei is more closely related to those of moulds than yeasts. FEBS Lett 555:469–477 Worley AC, Raper KB, Hohl M (1979) Fonticula alba: a new cellular slime mold (Acrasiomycetes). Mycologia 71:746–76 Yoshida M, Nakayama T, Inouye I (2009) Nuclearia thermophila sp. nov. (Nucleariidae), a new nucleariid species isolated from Yunoko Lake in Nikko (Japan). Eur J Protistol 45:147–155.

.

2

Microsporidia – Highly Reduced and Derived Relatives of Fungi

BRYONY A.P. WILLIAMS1, PATRICK J. KEELING2

CONTENTS I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. History of the Classification of Microsporidia. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . III. Microsporidia in or out of the Fungi?. . . . . . . . . . . . IV. Cellular and Genomic Reduction in the Microsporidia. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Genome Reduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Reduced Microsporidian Biochemistry and Molecular Biology . . . . . . . . . . . . . . . . . . . . . B. Simplified Microsporidian Cell Structure: Variations on the Eukaryotic Theme. . . . . . . . . . V. Diversity of Microsporidia. . . . . . . . . . . . . . . . . . . . . . . . VI. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

25 26 27 29 30 30 31 33 33 33

I. Introduction Microsporidia are a diverse group of eukaryotic parasites with over 1200 described species. Most species infect animals, though some species infect other single-celled eukaryotes, for example the gregarine Selenidium and the amoeba Vanella (Leger and Dubosq 1909; Scheid 2007). However, microsporidia are increasingly important humans pathogens with 14 species now recognized as causing illness in the immunocompromised; and given the diversity of microsporidia there are likely many other species extant with the potential to infect humans given the right opportunity (Mathis et al. 2005). The spores of microsporidia are highly environmentally resistant and in humans are generally transmitted via the fecal– oral route, frequently causing intestinal infections. Other sites of infections are also possible 1

Biosciences, Geoffrey Pope Building, College of Life and Environmental Sciences, University of Exeter, Stocker Road, Exeter, EX4 4QD, UK; e-mail: [email protected] 2 Canadian Institute for Advanced Research, Botany Department, University of British Columbia, 3529–6270 University Boulevard, Vancouver, British Columbia, V6T 1Z4, Canada; e-mail: [email protected]

and once in the body the parasites often migrate and can infect virtually any tissue of the body. These disseminated infections are rare, but generally far more serious and may result in death (Coyle et al. 2004). The microsporidia have an interesting but not fully understood mechanism of cell invasion. The infective stage of microsporidia is the dormant spore, a characteristic structure dominated by a tightly coiled filament called the polar filament (Fig. 2.1; Kudo 1918). The germination of spores is triggered by a variety of environmental cues, which leads to expulsion of the polar filament, and its eversion to become a tube. The polar vacuole expands, forcing the contents of the spore through the tube into the host cell where it forms a rather amorphous cell, at this stage called a meront (Va´vra and Larsson 1999). Expulsion of the polar tube is thought to be precipitated by an increase in pressure within the spore. This increase in pressure is likely produced by a rapid influx of water into the cell. Aquaporin proteins encoded in the Encephalitozoon cuniculi genome are suggested to provide the means of entrance of water in the cell across the plasma membrane (Frixione et al. 1997; Ghosh et al. 2006), the permeability of which may be controlled by changes in ion concentrations in the environment. It is also hypothesized that osmotic pressure may be increased inside the spore at the time of germination by the breakdown of the disaccharide sugar trehalose into less complex sugars, driving an influx of water. In line with this, it has been shown in some aquatic species of microsporidia that there is a rapid disappearance of trehalose and the appearance of smaller sugars at germination (Undeen and Vander Meer 1999). Conventional theories suggest that triggers for germination are chemical and that the polar tube physically pierces through the host cell, and the spore contents pass directly into the host cell cytoplasm, where it can directly interact with Evolution of Fungi and Fungal-Like Organisms, The Mycota XIV S. Po¨ggeler and J. Wo¨stemeyer (Eds.) © Springer-Verlag Berlin Heidelberg 2011

26

Bryony A.P. Williams and Patrick J. Keeling

Fig. 2.1. Microsporidian generalized life cycle. The infection process starts with the expulsion of the polar tube forced out by expansion of the posterior vacuole. The tube pierces the host cell and the microsporidian passes its contents into the host cell cytoplasm to form a meront. These develop (sometimes within a vacuole – as shown here) into more spores that are released into the environment for further infection

the host cell environment. However, recent data also suggest that infection may be mediated by a molecular anchoring of the spore to the host cell surface glycosaminoglycans prior to germination (Southern et al. 2006). This discovery of a molecular interaction between microsporidia and host before infection may provide a basis for understanding the host specificity of different microsporidia species. It has also been observed that the end of the everted polar tube, and subsequently the spore contents, can be endocytosed by the host after germination rather than being passed through the membrane via the polar tube (Schottelius et al. 2000). Once inside the cell, microsporidia life cycles are variable from species to species but generally there is a phase of meront cell division, followed by sporogony, in which the infection apparatus and spore wall develop, followed by maturation and release of the spores through the rupture of the host cell (Schottelius et al. 2000).

II. History of the Classification of Microsporidia Microsporidia have been studied for over 150 years, but like many single-celled eukaryotic lineages, they have a long history of phylogenetic misplacement (see Chapter 1 in this volume). Suprisingly, when Na¨geli first examined them in the 1800s (Na¨geli 1857), he grouped microsporidia with the Schizomycete fungi, though at this time there was little information on the untrastructural differences between single-celled

organisms. The Schizomycete fungi since turned out to be a collection of many unrelated species of eukaryotes and prokaryotes, and microsporidia were soon were re-assigned to their own group within the Sporozoa class of Protozoa (Balbiani 1882) and later grouped with the Myxozoa in the Cnidosporidia (Do¨flein 1901). This was based on the fact that both taxa share the characters of a polar filament and an ability to form spores, which were thought to be homologous traits. This grouping persisted for over 70 years until a greater understanding of the ultrastructure and biochemistry of the Cnidosporidia revealed fundamental differences in the two characters between various subgroups (Lom 1962), eventually leading to the microsporidia being moved into their own phylum (Sprague 1969, 1977). In 1983, Cavalier-Smith placed microspora in the subkingdom Archezoa based on the perceived absence of mitochondria in the group. He hypothesized that microsporidia and other Archezoa (diplomonads, Parabasalia, Archamoebae) were primitively amitochondriate, having branched away from the main eukaryotic lineage before the mitochondrial endosymbiosis (Cavalier-Smith 1983). With the advent of molecular phylogeny, the source of characters for reconstructing protozoon phylogeny switched from ultrastructure and biochemistry to gene and amino acid sequences. The sequencing and phylogenetic analysis of the small subunit ribosomal RNA of the microsporidian Vairimorpha necatrix appeared to provide a stunning confirmation of

Microsporidia – Highly Reduced and Derived Relatives of Fungi

the hypothesis that microsporidia were very earlybranching eukaryotes, because the phylogenies not only placed microsporidia at the base of eukaryotic tree (Vossbrinck et al. 1987), but also revealed the presence of fused 5.8S-LSU rRNA, a character shared with prokaryotes (Vossbrinck and Woese 1986). Subsequent phylogenies of EF1-a and EF-2 reinforced this, also showing microsporidia to branch early (Kamaishi et al. 1996). Indeed, microsporidia often branched even earlier than the other Archezoa, which together with other characters led to the idea that they may be the first branch of extant eukaryotes (Patterson 1994). In 1996, the phylogenetic analysis of a- and b-tubulin sequences from Antonospora (Nosema) locustae, Spraguea lophii, Encephalitozoon cuniculi, and Encephalitozoon hellem suggested an alternative position for microsporidia as relatives of the Fungi (Edlind et al. 1996; Keeling and Doolittle 1996). This prompted re-analysis of the previously analysed data sets using methods correcting for potential artefacts and the sequencing of new phylogenetic markers in microsporidia (Fast et al. 1999; Hirt et al. 1999; Van de Peer et al. 2000). One particular source of artefact pervasive to molecular phylogenies of microsporidia is long-branch attraction. If two taxa in a phylogeny have significantly longer branches than the other taxa, they have an increasing probability of being grouped together irrespective of their phylogenetic relationships as more characters are added to the data set (Felsenstein 1978). Another issue that has confounded microsporidian phylogenies is the problem of unequal site-rate variation. Not all sites vary freely or at the same rate across sequence alignments and some sites may not vary in any of the taxa. These sites can be hypothesised to be invariable, particularly in an alignment, which includes sequences representing highly diverse taxa, including eubacteria and diverse eukaryote lineages. If invariable sites are included in an analysis and among-site rate variation is not taken into account, the number of changes that have occurred at the fast evolving sites may be underestimated.

This can cause fast evolving sequences to branch together (Yang 1996). After these sources of artefacts were corrected for, microsporidia were consistently found to branch within or at the base of the Fungi in most molecular markers previously used to support their basal position, except in the phylogeny of SSU rRNA where they remained basal to eukar-

27

yotes (Fast et al. 1999; Hirt et al. 1999; Van de Peer et al. 2000). The importance of long-branch attraction in analysis of microsporidian genes was recently highlighted by an analysis of 99 genes from the E. cuniculi genome. This showed a clear inverse correlation between the rate of substitution in a gene and its propensity to unite microsporidia and fungi in its phylogeny (Thomarat et al. 2004), which reinforces the conclusion that early phylogenies were hampered by long-branch attraction problems.

III. Microsporidia in or out of the Fungi? Given that microsporidia are phylogenetically allied to the Fungi, the question is still open as to whether microsporidia evolved from a lineage within the Fungi or are a distinct lineage of eukaryotes branching at the base of the Fungi, therefore representing a sister group (see Chapter 1 in this volume). Apart from phylogenies, which may of course be vulnerable to artefactual relationships caused by poor taxon sampling, long-branch attraction, or lack of fit of the characters to the model of evolutionary change used to infer the phylogeny, are there any reliable synapomorphic characters that link microsporidia to the Fungi? There are three characters that solidly place microsporidia within the opisthokonts (animals, fungi, and their closest protist relatives). One is a 10-amino-acid insertion within the EF1A protein that is unique to opisthokonts and is also present within microsporidia (Baldauf et al. 1996; Kamaishi et al. 1996). The second is the presence of two separate genes for dihydrofolate reductase and thymidylate synthase (Duffieux et al. 1998), which are fused in non-opithokont lineages (Stechmann and Cavalier-Smith 2002). Lastly, microsporidia and other opisthokonts also share a related holoarchaeal-type tyrosyltRNA synthetase, presumably laterally transferred into the ancestral lineages that gave rise to animals and fungi (Huang et al. 2005). These place microsporidia within the opithokonts but not specifically within the Fungi. More specifically, one character that has been used to place microsporidia specifically outside or at the base of the Fungi is a two amino acid deletion, again in EF1A (Tanabe et al. 2002, 2005). This deletion exits in all fungi where EF1A has

28

Bryony A.P. Williams and Patrick J. Keeling

been sequenced, but generally not in animals, nor does it occur in any microsporidia studied so far. This would suggest that microsporidia be placed toward the base of the Fungi, if not outside the group. However this character will probably never be definitive because some entomophagous fungi and many chytrid and fungi, including the important genus Rozella (see below), have lost the conventional EF1A gene in favour of a distantly related member of the same GTPase family, named EF-like or EFL (James et al. 2006a; Keeling and Inagaki 2004). Without EF1A in these lineages, we cannot determine whether this deletion was present in the most basal fungal lineages, and therefore whether any of these lineages branched before or concurrently with microsporidia. None of these molecular characters place microsporidia specifically within the Fungi, however there are some characteristics of microsporidia that are typically fungal, for example the presence of closed intranuclear mitosis and the presence of chitin in the cell wall. Furthermore several relatively conserved homeotic and TALE genes have been identified in the E. cuniculi genome (Burglin 2003). These are characteristically found in plants, animals and fungi, controlling polarity, morphology and mating types. Two of these genes are adjacent in the E. cuniculi genome, reminiscent of a mating-type loci found in other fungi (Fraser and Heitman 2004). This does open an intriguing question about why a mating-type locus would occur in E. cuniculi, as sex has never been documented in this species, though molecular markers for meiosis are present (Gill and Fast 2007). In at least some zygomycete fungi, the mating type locus occurs as a syntenic cluster consisting of a high mobility group transcription factor flanked by a triose phosphate transporter and an RNA helicase. This gene cluster also occurs in both E. cuniculi and E. bieneusi and is partially conserved in A. locustae (Lee et al. 2008). In general the conservation of synteny between these microsporidia and zygomycetes is much higher than it is between microsporidia and other fungi (Lee et al. 2008). This provides a convincing character to place microsporidia within the Fungi and specifically suggests a relationship between microsporidia and zygomycetes. This is interesting as re-analysis of a- and b-tubulin protein sequences with representation from several chytridiomycete, ascomycete, basio-

diomycete and zygomycete lineages also suggested some affinity to zygomycetes, and specifically a clade of entomophagous zygomycetes (Keeling 2003; Keeling et al. 2000). The analysis of tubulins is problematic, however, since evolutionary rates correlate to the presence of flagella. Both the microsporidia and most fungi lack flagella, and share long-branch tubulins, which could easily lead to an erroneous placement of microsporidia within such flagella-lacking fungi just as other genes erroneously placed microsporidia at the base of eukaryotes (Keeling 2003; Keeling et al. 2000). A concatenated phylogeny of eight protein-coding genes using a smaller taxon sample (but nevertheless including representatives of all four major fungal lineages) found microsporidia to branch at the base of the ascomycetes and basdiomycetes (Gill and Fast 2006). The authors of this study also suggest a possibility of long-branch attraction between the representative organisms within this clade and microsporidia, and that there are problems of taxon sampling as just a single representative for the chytrids was available for the eight-gene concatenation. More recently, the Fungal Tree of Life project attempted to resolve some of the major issues confounding fungal phylogenetics and classification by producing a multigene phylogeny with an unprecedented number of fungal lineages, including the microsporidia (James et al. 2006a). Here the microsporidia branch near the base of the fungal tree, and fascinatingly they form a clade with the parasite Rozella allomycis (James et al. 2006a). Rozella is not present in any other analysis including microsporidia, and indeed few genes have been sampled from both lineages, so this remains an intriguing avenue to follow. This relationship between microsporidia and Rozella was proposed on the basis a phylogeny of two protein coding genes: RPB1 and RPB2. It is not well supported statistically, and several equally likely branching positions for microsporidia within the fungal tree of life were also proposed, one of which placed microsporidia basal to all fungi (James et al. 2006a). There are, however, a couple of conspicuous similarities between R. allomycis and microsporidia. Both are obligate intracellular parasites and therefore both require a mechanism for entering the host cell. In Rozella it is reported that the zoospore stage forms an attachment to the Allomyces host cell, which is followed by retraction of the flagellum, encystment,

Microsporidia – Highly Reduced and Derived Relatives of Fungi

proceeded by the formation of a germ tube. The parasite then injects its protoplast into the host cell, forced out of the cyst by a rapidly expanding vacuole (Held 1972). Of course, there is no evidence of a flagellum in microsporidia, or even any other 9+ 2 microtubular structures. However the need for an expanding vacuole is a somewhat reminiscent of the polar vacuole in microsporidia which forces out the sporoplasm (see above) and provides an interesting possible homology between these two unique fungal groups. As with molecular data, there are few hints from morphology about the exact relationship between microsporidia and fungi. The relationship between microsporidia and entomophagous zygomycetes prompted some speculation that the microsporidian infection mechanism might be related to the mechanism used for zygospore dispersal in this group of insect parasites (Keeling 2003; Keeling et al. 2000), and interestingly a similar position among the Fungi was previously suggested based on possible homologies in cell biology and SSU ribosomal RNA phylogenies (Cavalier-Smith 2001). Cavalier-Smith (2001) hypothesized that there could be homology between the filamentous appendages in the infective trichospores of the harpellalean fungi and the polar tube of the microsporidia. In the spore stage of the harpellalean fungi, filamentous appendages develop and are coiled up within the cell. These are expelled as long filaments during germination (Lichtwar 1967). However their function is thought to be to allow entanglement of the fungus with the substrate and, although there are some morphological similarities to the microsporidian polar tubes, the two do not share functional similarity. There are two main impediments to any attempt to clarify the relationship between microsporidia and fungi. First, data representing the full diversity of fungi and microsporidia are generally lacking (Bass et al. 2007; Gill and Fast 2006; James et al. 2006b; Keeling 2003). Even if the presently available phylogenetic markers were sufficient and the methods to extract phylogenetic signal from the data were adequate, we probably have not yet sampled some major fungal lineages and so may not yet have sampled from the fungal lineage most closely related to the microsporidia. Second, the microsporidia are so derived at all levels, from molecular sequences, to biochemistry,

29

to morphology that it is possible that available markers and methods will not be sufficient to address the problem simply because their rapid adaptation and evolution have erased too much of their history. If this is the case, then one tack to take must be to identify the least divergent lineages of microsporidia. It is possible that early-diverging lineages of microsporidia were not subject to the same massive levels of change (similar to the recent discovery of an algal relative of apicomplexan parasites that retains many characters believed to be ancestral; Moore et al. 2008), and if so perhaps these lineages could be linked to their closest relatives more clearly. Further exploration of the diversity of both fungi and microsporidia is needed in order to know.

IV. Cellular and Genomic Reduction in the Microsporidia Microsporidia were once regarded as ‘primitive’, but when they are re-evaluated as opithokonts, and perhaps fungi, we now see microsporidia as ‘reduced’ instead. Indeed, reduction and loss are two prevailing themes in just about every aspect of their biology. Reduction is seen in the simplification of many molecular systems (e.g., the ribosomes and ribosomal RNAs are both similar in size to those of prokaryotes rather than other eukaryotes), the absence of many components of some cellular and biochemical pathways, and the degeneration of organelles, such as the mitochondrion. Loss too is seen at all levels from the loss of genes, loss of introns, loss of complete biochemical pathways and perhaps even the complete loss of organelles (e.g., the peroxisome and flagella). It is interesting to ask whether this drastic reorganization of the cell and it molecular biology is really unique to the microsporidia, or are aspects of such reduction evident in other fungal lineages? In fact, many fungal lineages do exhibit many of the same characteristics that reduction and loss have generated in microsporidia, but the difference is that we do not see them altogether in one cell. Below, we will review some of the levels where microsporidian reduction is best studied and most severe, and we will compare a few such traits to corresponding characteristics of fungi.

30

Bryony A.P. Williams and Patrick J. Keeling

A. Genome Reduction Members of the microsporidia bear the smallest known eukaryotic nuclear genomes and are now model organisms for understanding genome compaction and reduction. The smallest documented microsporidian genome is 2.3 Mb and the largest estimated at 23 Mb, with a range of sizes in between (Belkorchia et al. 2008; Biderre et al. 1994; Me´te´nier and Vivare`s 2001). However, before the E. cuniculi genome was sequenced, Saccharomyces cerevisiae provided this model of genome compaction. Whereas E. cuniculi has a genome of just 2.9 Mb with an estimated 1997 protein coding genes (Katinka et al. 2001), baker’s yeast has a 12-Mb genome with around 6000 genes (Dujon 1996): a proteome three times the size in a genome six times the size. In other fungi moderate genome reductions is also well known: the Pneumocystis carinii genome is just 6.5 Mb (Cushion and Smulian 2001) and the genome of the basiodiomycete Malassezia is around 9 Mb with 4285 genes (Xu et al. 2007). Although these are larger than the smallest microsporidian genome, they are within the range of microsporidian genome sizes, and with eukaryotic genomes ranging from 2.3 to 670000 Mb, they are also certainly at the small end of the spectrum in general. Moreover, smaller fungal genomes are still very likely to exist, given the number of species that have been investigated is a small proportion of what exists, so it is not inconceivable that a fungal genome smaller than the smallest microsporidian genome will be described. Overall, genome reduction appears to be a phenomenon that occurs in several lineages throughout the Fungi, though perhaps not the same extent as we see in the microsporidia, and perhaps not as consistently. One of the notable traits of microsporidian genomes that arises as a consequence of this reduction is a high level of conservation of gene order between distantly related species.

undergo a gene rearrangement event without disrupting a gene (Hurst et al. 2002; Slamovits et al. 2004). This is not unique to the microsporidia but also studied in yeast, which also have short distances between genes (Poyatos and Hurst 2007). However the number of shared syntenic genes between different species is exaggerated in the microsporidia where the spaces between genes are shortened to the extreme. Microsporidia have also been shown to have a messy system of transcription, apparently owing again to the fact that intergenic spaces are small, in this case so small that promoters and termination signals can no longer be contained within them (Corradi et al. 2008; Williams et al. 2005). Again this system is not exclusive to microsporidia and is found in other fungi with instances of overlapping transcription being found in mRNA transcripts from S. cerevisiae (Hurowitz and Brown 2003), but as with genome rearrangements, the effect is exaggerated in the extremely reduced genomes of microsporidia. 1. Reduced Microsporidian Biochemistry and Molecular Biology

This was initially observed between A. locustae and E. cuniculi and later between A. locustae, E. cuniculi and Enterocytozoon bieneusi (Corradi et al. 2007; Slamovits et al. 2004).

Microsporidia have lost many metabolic pathways and molecular systems compared to their freeliving relatives. The most obvious are the loss of the electron transport chain and the Krebs cycle. There are no components of the heme biosynthesis pathway encoded in the E. cuniculi genome. In the absence of mitochondrial cytochromes there are fewer heme-containing proteins and it is possible that heme is scavenged from the host rather than being synthesized within the microsporidia (Vivare`s et al. 2002). There are no fatty acid or cholesterol synthesis pathway components, also suggesting a reliance on the host (Vivare`s et al. 2002). There are also many pathways that are not completely lost, but merely downsized, and whittled down to a core set of proteins. One well described example is the various DNA repair systems that exist in eukaryotic cells, which are of course essential systems in cell maintenance.

This is hypothesized to be a consequence of the difficulty in rearranging genes when intergenic spaces between genes are very small; the larger the gap between gene pairs the more likely they are to

Reviewing the extent of loss of components the major DNA repairs systems in E. cuniculi compared to S. cerevisiae, including non-homologous end joining, homologous recombination repair, mismatch repair

Microsporidia – Highly Reduced and Derived Relatives of Fungi (MMR), nucleotide excision repair, base excision repair and methyltransferase repair, showed all pathways were reduced to some extent in microsporidia, but some components of each individual process were also retained, suggesting the retention of a minimal DNA repair system (Gill and Fast 2007).

The network of protein kinases involved in signaling processes within the cell seems to be similarly reduced. Here there is a bias towards signaling pathways involved in cell maintenance, whereas signaling processes involved in sensing environmental change are more pared down, probably reflecting the protective and stable intracellular in which the microsporidia reside (Miranda-Saavedra et al. 2007).

B. Simplified Microsporidian Cell Structure: Variations on the Eukaryotic Theme Microsporidia were long thought to be an example of the primitive cellular state of the eukaryotic cell. They were suggested to be primitively lacking mitochondria, Golgi and peroxisomes. Of course, in the context of microsporidia being closely related to fungi it makes no sense for them to be primitively lacking any of these organelles; instead they have been lost or transformed, and here we will discuss their evolution. The mitochondrion has attracted much attention, in part due to the Archezoa hypothesis described earlier. A homologous organelle is now known to exist in microsporidia, but it is so highly reduced it was only identified through the immunolocalisation of an endosymbiont-derived mitochondrial Hsp70 protein (Williams et al. 2002). The complete genome of E. cuniculi (Katinka et al. 2001) revealed that the mitochondrion, termed mitosome, had lost its genome, lost its electron transport chain and the Krebs cycle and was predicted to exist mainly to allow iron–sulfur cluster assembly for export and integration into cytosolic iron–sulfur proteins, which is an essential function of mitochondria in yeast cells. This conclusion was substantiated by work that has localized proteins involved in iron–sulfur cluster in both E. cuniculi and Trachipleistophora hominis (Goldberg et al. 2008; Williams et al. 2008b). It has also been shown that some E. cuniculi proteins involved in iron–sulfur cluster assembly can complement yeast homologs. Interestingly, however, some of these predicted mitochondrial

31

proteins do not seem to be localized to the mitosomes in all species of microsporidia. Goldberg et al. (2008) show that, in E. cuniculi, both the iron–sulfur cluster proteins ISCU1p and cysteine desulfurase localize to a mitosome whereas, in T. hominis, they appear to be cytoplasmic. Similarly, in E. cuniculi, ferredoxin, a typically mitochondrial protein in yeast cells, has a punctate distribution, consistent with a mitosomal location whereas the mitochondrial glycerol-3-phoshate dehydrogenase appears to have a cytoplasmic distribution (Williams et al. 2008b), but there is evidence that this protein is more likely mitosomal in A. locustae (Burri et al. 2006). This may indicate that mitosomes in different species of microsporidia may differ and be in different states of degeneration. This is also is reflected in a different complement of mitochondrial gene homologs in different microsporidia (Burri et al. 2006; Williams et al. 2008a). Various levels of mitochondrial reduction also occur in other fungal mitochondria. For example, mitochondrial genomes have been completely lost in the anaerobic chytrid fungi in the family Neocallimastigaceae. Here the mitochondria have been transformed into hydrogenosomes: mitochondrion-derived organelles that produce ATP via decarboxylation of pyruvate and generate hydrogen gas and, as in microsporidia, rely exclusively on proteins encoded in the nuclear genome (Yarlett et al. 1986; van der Giezen et al. 2002). Diverse yeast lineages, the so-called petite-positive yeast species, can also exist with a genome-less mitochondrion, and the ability to survive without mitochondrial DNA is a widespread trait across the yeasts (Fekete et al. 2007) and in some basiodiomycetes (Kuscera et al. 2000). One specific factor that may have allowed the degeneration of the mitochondria in the microsporidia is the capacity of the parasite cell to import ATP from the host cell. This ability has known of for quite some time, and was identified through the measuring relative levels of ATP consumption in blue crabs, infected or uninfected with microsporidia (Weidner and Trager 1973). The genetic basis for this trait was identified in the E. cuniculi genome as a family of ATP/ADP translocases that otherwise exist in only in the eubacterial energy parasites of the Rickettsia and Chlamydia lineages, and in plastids where they import ATP into storage plastids to drive biosynthetic processes (Katinka et al. 2001; Neuhaus and

32

Eu ka o ry te s

The peroxisome is an organelle that has not been seen in the microsporidian cell, though this may not be so surprising since peroxisomes are

ia

Microsporidia genomes have a reduced complement of Rabs, SNAREs and vesicle coat proteins (Dacks and Field 2007; Katinka et al. 2001). Unlike a typical Golgi, the microsporidian Golgi is also suggested not to have COPI and COPII vesicles, and therefore require a direct connection somewhere between the ER and the Golgi (Beznoussenko et al. 2007). So again we see a typical eukaryotic system present in microsporidia, but very much pared down.

i Fung

Emes 2000; Tsaousis et al. 2008). Homologous genes have now been found in the genomes of E. cuniculi, E. bieneusi, S. lophii (Corradi et al. 2007; Hinkle et al. 1997; Katinka et al. 2001) and A. locustae (http://gmod.mbl.edu/perl/site/antonospora01). This gene is thought to have moved between lineages by lateral gene transfer (Tyra et al. 2007), and it appears possible that the microsporidia proteins may also have been acquired by lateral gene transfer, perhaps from either a rickettsial or chlamydial parasite (Fig. 2.2; Richards et al. 2003). Another organelle of note that is absent from microsporidia is a classical stacked Golgi dictyostome. This is not to say there is no requirement for protein sorting in microsporidia; indeed, proteins inserted into the spore wall and into the polar tube appear to be trafficked much like secreted proteins in other eukaryotes. The microsporidian Golgi exists as a mass of vesicular tubules, which is present throughout the life cycle, but is particularly apparent during sporogony when the spore wall and polar tube are being formed.

orid

ea

ha

c Ar

p ros mic

Eubacteria

Rickettsiales

Fig. 2.2. Lateral transfer of the energy parasite ATP/ADP translocase. The ability to exploit host ATP may have been acquired by microsporidia through the lateral gene transfer of the gene encoding an ATP/ ADP translocase from the lineage of proteobacteria containing Rickettia parasites (adapted from Richards et al. 2003)

Bryony A.P. Williams and Patrick J. Keeling

Hypothesised transfer of genes allowing ATP uptake from host

small organelles and might be easily overlooked in cells as small as those of microsporidia. However, a gene for one of one of the hallmark enzymes of the organelle, catalase, is definitely absent from the complete E. cuniculi genome. An alternative catalase is present in the genome of the insect parasite A. locustae (Fast et al. 2003), however this is a kind of catalase that is not normally localized to a peroxisome in other organisms, but rather can be secreted into the spore wall in certain fungi, possibly to protect the spore from an oxidative burst from the host (Paris et al. 2003). Furthermore it appears to be derived by lateral gene transfer from an, obviously, organelle lacking proteobacterium (Fast et al. 2003). Catalase-like activity has been observed within the polar vacuole in the fish-infecting microsporidian S. lophii. In the presence of catalase, diaminobenzidine (DAB) reacts with H2O2 to form a dark precipitate that can be visualized with by electron microscopy. Such a reaction was observed around the polar vacuole in S. lophii, and based on this it was suggested that the polar vacuole is a highly derived peroxisome (Findley et al. 2005), though there is not yet any evidence for a gene encoding a peroxisomal catalase in microsporidia (Katinka et al. 2001). An alternative possibility is that this vacuole is related to the vacuoles of S. cerevisiae and other fungi, which are considered analogs of mammalian lysosomes (Weisman 2003). Similarly, in the infection strategy of the rice blast fungus Magnaporthe grisea, the vacuole plays a key role and acts as the site of breakdown of lipid reserves to create the high turgor pressure needed for the fungus to penetrate leaf surfaces (Weber et al. 2001).

Microsporidia – Highly Reduced and Derived Relatives of Fungi

V. Diversity of Microsporidia Most of what we know of the molecular biology of microsporidia has been inferred from two species where extensive genomic data have been available for several years, A. locustae and E. cuniculi. While it is always tempting to generalize from model species, it is likely that these two species represent only a small fraction of the variability within microsporidia. Indeed, a genome sequence survey of two other species, Brachiola algerae and Edhazardia aedis, shows that the gene complement of these is likely similar to the better-studied E. cuniculi and A. locustae, but the overall architecture of these gene-sparse genome is completely different (Williams et al. 2008c). Now, two further draft genomes for N. ceranae and E. bieneusi are available (Akiyoshi et al. 2009; Cornman et al. 2009). In the case of E. bieneusi, it has demonstrated even further metabolic reduction than E. cuniculi with some key pathways, such as glycolysis and trehalose metabolism seemingly degenerated. Additional new genome projects for microsporidia with larger genomes are underway (e.g., Belkorchia et al. 2008; Xu et al. 2006) and as these data emerge they will undoubtedly reveal still more diversity within microsporidia. Given the large number of described species, we are likely to see a degree of variation in a wide range of different cellular characteristics, and perhaps we may even pinpoint characteristics that link the microsporidia to other fungi.

VI. Conclusion The exact phylogenetic position of the microsporidia remains an open question: it is clear they are not a basal eukaryotic lineage and are somehow related to fungi. Current data is converging on the idea that that they are a true fungal group and potentially allied to a zygomycete lineage. However the discovery of their true sister group requires further analysis and more data. The E. cuniculi genome not only provided a wealth of direct evidence for many conclusions on various aspects of microsporidian cell and molecular biology, but it also acted as a catalyst for other research questions, which are still transforming our view of many characteristics of this parasite. At the same time, however, greater

33

sampling from across the taxonomic breadth of the group is and will continue to challenge generalizations which are based on E. cuniculi or any other single species; so one of the major lessons from recent microsporidian research is to expect the unexpected. Much of what we know about the biology of these parasites relates to their bending or breaking the ‘rules’ of biology, so perhaps it is not surprising that the rules of microsporidian biology are not easily predictable.

References Akiyoshi DE, Morrison HG, Lei S, Feng X, Zhang Q, Corradi N, Mayanja H, Tumwine JK, Keeling PJ, Weiss LM et al. (2009) Genomic survey of the noncultivatable opportunistic human pathogen, Enterocytozoon bieneusi. PLoS Pathog 5:e1000261 Balbiani G (1882) Sur les microsporidies ou sporospermies des articules. C R Acad Sci 95:1168–1171 Baldauf SL, Palmer JD and Doolittle WF (1996) The root of the universal tree and the origin of eukaryotes based on elongation factor phylogeny. Proc Natl Acad Sci USA 93:7749–7754 Bass D, Howe A, Brown N, Barton H, Demidova M, Michelle H, Li L, Sanders H, Watkinson SC, Willcock S et al. (2007) Yeast forms dominate fungal diversity in the deep oceans. Proc Biol Sci 274:3069–3077 Belkorchia A, Biderre C, Militon C, Polonais V, Wincker P, Jubin C, Delbac F, Peyretaillade E and Peyret P (2008) In vitro propagation of the microsporidian pathogen Brachiola algerae and studies of its chromosome and ribosomal DNA organization in the context of the complete genome sequencing project. Parasitol Int 57:62–71 Beznoussenko GV, Dolgikh VV, Seliverstova EV, Semenov PB, Tokarev YS, Trucco A, Micaroni M, Di Giandomenico D, Auinger P, Senderskiy IV et al. (2007) Analogs of the Golgi complex in microsporidia: structure and avesicular mechanisms of function. J Cell Sci 120:1288–1298 Biderre C, Pages M, Metenier G, David D, Bata J, Prensier G and Vivares CP (1994) On small genomes in eukaryotic organisms: molecular karyotypes of two microsporidian species (Protozoa) parasites of vertebrates. C R Acad Sci III 317:399–404 Burglin TR (2003) The homeobox genes of Encephalitozoon cuniculi (Microsporidia) reveal a putative mating-type locus. Dev Genes Evol 213:50–52 Burri L, Williams BA, Bursac D, Lithgow T and Keeling PJ (2006) Microsporidian mitosomes retain elements of the general mitochondrial targeting system. Proc Natl Acad Sci USA 103:15916–15920 Cavalier-Smith T (1983) A 6-kingdom classification and a unified phylogeny. In: (ed) Endocytobiology Vol II. Walter de Gruyter & co., Berlin New York, pp 1027–1034

34

Bryony A.P. Williams and Patrick J. Keeling

Cavalier-Smith T (2001) What are Fungi? In: G. L. McLaughlin EGM, and Lempke P. A., (ed) The Mycota VII Part A. Springer-Verlag, Berlin, pp 1–37 Cornman RS, Chen YP, Schatz MC, Street C, Zhao Y, Desany B, Egholm M, Hutchison S, Pettis JS, Lipkin WI et al. (2009) Genomic analyses of the microsporidian Nosema ceranae, an emergent pathogen of honey bees. PLoS Pathog 5:e1000466 Corradi N, Akiyoshi DE, Morrison HG, Feng X, Weiss LM, Tzipori S and Keeling PJ (2007) Patterns of genome evolution among the microsporidian parasites Encephalitozoon cuniculi, Antonospora locustae and Enterocytozoon bieneusi. PLoS ONE 2:e1277 Corradi N, Gangaeva A and Keeling PJ (2008) Comparative profiling of overlapping transcription in the compacted genomes of microsporidia Antonospora locustae and Encephalitozoon cuniculi. Genomics 91:388–393 Coyle CM, Weiss LM, Rhodes LV, 3rd, Cali A, Takvorian PM, Brown DF, Visvesvara GS, Xiao L, Naktin J, Young E et al. (2004) Fatal myositis due to the microsporidian Brachiola algerae, a mosquito pathogen. N Engl J Med 351:42–47 Cushion MT and Smulian AG (2001) The Pneumocystis genome project: update and issues. J Eukaryot Microbiol Suppl:182S–183S Dacks JB and Field MC (2007) Evolution of the eukaryotic membrane-trafficking system: origin, tempo and mode. J Cell Sci 120:2977–2985 Do¨flein F (1901) Die Protozoen als Parasiten und Krankheitserreger nach biologischen Gesichtspunkten dargestellt. Fisher, G, Jena Duffieux F, Peyret P, Roe BA and Vivares CP (1998) First report on the systematic sequencing of the small genome of Encephalitozoon cuniculi (Protozoa, Microspora): gene organization of a 4.3 kbp region on chromosome I. Microb Comp Genomics 3:1–11 Dujon B (1996) The yeast genome project: What did we learn? Trends Genet 12:263–270 Edlind TD, Li J, Visversvara GS, Vodkin MH, McLaughlin GL and Katiyar SK (1996) Phylogenetic analysis of the b-tubulin sequences from amitochondriate protozoa. Mol Phylogenet Evol 5:359–367 Fast NM, Law JS, Williams BA and Keeling PJ (2003) Bacterial catalase in the microsporidian Nosema locustae: implications for microsporidian metabolism and genome evolution. Eukaryot Cell 2:1069–1075 Fast NM, Logsdon JM, Jr. and Doolittle WF (1999) Phylogenetic analysis of the TATA box binding protein (TBP) gene from Nosema locustae: evidence for a microsporidia–fungi relationship and spliceosomal intron loss. Mol Biol Evol 16:1415–1419 Fekete V, Cierna M, Polakova S, Piskur J and Sulo P (2007) Transition of the ability to generate petites in the Saccharomyces/Kluyveromyces complex. FEMS Yeast Res 7:1237–1247 Felsenstein J (1978) Cases in which parsimony or compatibility methods will be positively misleading. Syst Zool 27:401–410 Findley AM, Weidner EH, Carman KR, Xu Z and Godbar JS (2005) Role of the posterior vacuole in Spraguea lophii (Microsporidia) spore hatching. Folia Parasitol (Praha) 52:111–117

Fraser JA and Heitman J (2004) Evolution of fungal sex chromosomes. Mol Microbiol 51:299–306 Frixione E, Ruiz L, Cerbon J and Undeen AH (1997) Germination of Nosema algerae (Microspora) spores: conditional inhibition by D2O, ethanol and Hg2+ suggests dependence of water influx upon membrane hydration and specific transmembrane pathways. J Eukaryot Microbiol 44:109–116 Ghosh K, Cappiello CD, McBride SM, Occi JL, Cali A, Takvorian PM, McDonald TV and Weiss LM (2006) Functional characterization of a putative aquaporin from Encephalitozoon cuniculi, a microsporidia pathogenic to humans. Int J Parasitol 36:57–62 Gill EE and Fast NM (2006) Assessing the microsporidia– fungi relationship: Combined phylogenetic analysis of eight genes. Gene 375:103–109 Gill EE and Fast NM (2007) Stripped-down DNA repair in a highly reduced parasite. BMC Mol Biol 8:24 Goldberg AV, Molik S, Tsaousis AD, Neumann K, Kuhnke G, Delbac F, Vivares CP, Hirt RP, Lill R and Embley TM (2008) Localization and functionality of microsporidian iron–sulphur cluster assembly proteins. Nature 452:624–628 Held AA (1972) Host–parasite relations between Allomyces and Rozella – parasite penetration depends on growth response of host cell-wall. Arch Microbiol 82:128–139 Hinkle G, Morrison HG and Sogin ML (1997) Genes coding for reverse transcriptase, DNA-directed RNA polymerase, and chitin synthase from the microsporidian Spraguea lophii. Biol Bull 193:250–251 Hirt RP, Logsdon JM, Jr., Healy B, Dorey MW, Doolittle WF and Embley TM (1999) Microsporidia are related to Fungi: evidence from the largest subunit of RNA polymerase II and other proteins. Proc Natl Acad Sci USA 96:580–585 Huang J, Xu Y and Gogarten JP (2005) The presence of a haloarchaeal type tyrosyl-tRNA synthetase marks the opisthokonts as monophyletic. Mol Biol Evol 22:2142–2146 Hurowitz EH and Brown PO (2003) Genome-wide analysis of mRNA lengths in Saccharomyces cerevisiae. Genome Biol 5:R2 Hurst LD, Williams EJ and Pal C (2002) Natural selection promotes the conservation of linkage of co-expressed genes. Trends Genet 18:604–606. James TY, Kauff F, Schoch CL, Matheny PB, Hofstetter V, Cox CJ, Celio G, Gueidan C, Fraker E, Miadlikowska J et al. (2006a) Reconstructing the early evolution of Fungi using a six-gene phylogeny. Nature 443: 818–822 James TY, Letcher PM, Longcore JE, Mozley-Standridge SE, Porter D, Powell MJ, Griffith GW and Vilgalys R (2006b) A molecular phylogeny of the flagellated fungi (Chytridiomycota) and description of a new phylum (Blastocladiomycota). Mycologia 98:860–871 Kamaishi T, Hashimoto T, Nakamura Y, Nakamura F, Murata S, Okada N, Okamoto K-I, Shimzu M and Hasegawa M (1996) Protein phylogeny of translation elongation factor EF-1a suggests Microsporidians are extremely ancient eukaryotes. J Mol Evol 42: 257–263

Microsporidia – Highly Reduced and Derived Relatives of Fungi Katinka MD, Duprat S, Cornillot E, Me´te´nier G, Thomarat F, Prensier G, Barbe V, Peyretaillade E, Brottier P, Wincker P et al. (2001) Genome sequence and gene compaction of the eukaryote parasite Encephalitozoon cuniculi. Nature 414:450–453 Keeling PJ (2003) Congruent evidence from alpha-tubulin and beta-tubulin gene phylogenies for a zygomycete origin of microsporidia. Fungal Genet Biol 38:298– 309 Keeling PJ and Doolittle WF (1996) Alpha-tubulin from early-diverging eukaryotic lineages and the evolution of the tubulin family. Mol Biol Evol 13:1297–1305 Keeling PJ and Inagaki Y (2004) A class of eukaryotic GTPase with a punctate distribution suggesting multiple functional replacements of translation elongation factor 1 alpha. Proc Natl Acad Sci USA 101:15380–15385 Keeling PJ, Luker MA and Palmer JD (2000) Evidence from beta-tubulin phylogeny that microsporidia evolved from within the fungi. Mol Biol Evol 17:23–31 Kudo R (1918) Experiments on the Extrusion of Polar Filaments of Cnidosporidian Spores. J Parasitol 4:141–147 Kuscera J, Pfeiffer I and Takeo K (2000) Biology of the red yeast Xanthophyllomyces dendrorhous (Phaffia rhodozyma). Mycoscience Volume 41:195–199 Lee SC, Corradi N, Byrnes EJ, 3rd, Torres-Martinez S, Dietrich FS, Keeling PJ and Heitman J (2008) Microsporidia evolved from ancestral sexual fungi. Curr Biol 18:1675–1679 Leger L and Dubosq O (1909) Zoology. A parasite microsporidia of a gregarine. Cr Hebd Acad Sci 148:733–734 Lichtwar RW (1967) Zygospores and spore appendages of Harpella (Trichomycetes) from larvae of Simuliidae. Mycologia 59:482–491 Lom J, and Va´vra, J. (1962) A proposal to the classification within the subphylum CNIDOSPORA. Syst Zool 11:172–175 Mathis A, Weber R and Deplazes P (2005) Zoonotic potential of the microsporidia. Clin Microbiol Rev 18:423–445 Me´te´nier G and Vivare`s CP (2001) Molecular characteristics and physiology of microsporidia. Microbes Infect 3:407–415 Miranda-Saavedra D, Stark MJ, Packer JC, Vivares CP, Doerig C and Barton GJ (2007) The complement of protein kinases of the microsporidium Encephalitozoon cuniculi in relation to those of Saccharomyces cerevisiae and Schizosaccharomyces pombe. BMC Genomics 8:309 Moore RB, Obornik M, Janouskovec J, Chrudimsky T, Vancova M, Green DH, Wright SW, Davies NW, Bolch CJ, Heimann K et al. (2008) A photosynthetic alveolate closely related to apicomplexan parasites. Nature 451:959–963 Na¨geli K (1857) Uber die neue Krankheit der Seidenraupe und verwandte Organistem. Botanische Zeitung 15:760–761 Neuhaus HE and Emes MJ (2000) Nonphotosynthetic metabolism in plastids. Annu Rev Plant Physiol Plant Mol Biol 51:111–140

35

Paris S, Wysong D, Debeaupuis JP, Shibuya K, Philippe B, Diamond RD and Latge JP (2003) Catalases of Aspergillus fumigatus. Infect Immun 71:3551–3562 Patterson DJ (1994) Protozoa: evolution and systematics. In: Hausmann K and Hulsmann N (ed) Progress in Protozoology. Fischer-Verlag, Stuttgart, pp Poyatos JF and Hurst LD (2007) The determinants of gene order conservation in yeasts. Genome Biol 8:R233 Richards TA, Hirt RP, Williams BA and Embley TM (2003) Horizontal gene transfer and the evolution of parasitic protozoa. Protist 154:17–32 Scheid P (2007) Mechanism of intrusion of a microspordian-like organism into the nucleus of host amoebae (Vannella sp.) isolated from a keratitis patient. Parasitol Res 101:1097–1102 Schottelius J, Schmetz C, Kock NP, Schuler T, Sobottka I and Fleischer B (2000) Presentation by scanning electron microscopy of the life cycle of microsporidia of the genus Encephalitozoon. Microbes Infect 2:1401–1406 Slamovits CH, Fast NM, Law JS and Keeling PJ (2004) Genome compaction and stability in microsporidian intracellular parasites. Curr Biol 14:891–896 Southern TR, Jolly CE, Lester ME and Hayman JR (2006) Identification of a microsporidia protein potentially involved in spore adherence to host cells. J Eukaryot Microbiol 53 [Suppl 1]:S68–S69 Sprague V (1969) Need for drastic revision of the classification of the subphylum Amoebogena. Prog Protozool Proc Int Congr Protozool 3:372 Sprague V (1977) Classification and Phylogeny of the Microsporidia. In: Bulla LAaC, T. C. (ed) Comparative Pathobiology. Plenum Press, New York, pp 1–30 Stechmann A and Cavalier-Smith T (2002) Rooting the eukaryote tree by using a derived gene fusion. Science 297:89–91 Tanabe Y, Watanabe MM and Sugiyama J (2002) Are Microsporidia really related to Fungi?: a reappraisal based on additional gene sequences from basal fungi. Mycol Res 106:1380–1391 Tanabe Y, Watanabe MM and Sugiyama J (2005) Evolutionary relationships among basal fungi (Chytridiomycota and Zygomycota): Insights from molecular phylogenetics. J Gen Appl Microbiol 51:267–276 Thomarat F, Vivare`s CP and Gouy M (2004) Phylogenetic analysis of the complete genome sequence of Encephalitozoon cuniculi supports the fungal origin of microsporidia and reveals a high frequency of fastevolving genes. J Mol Evol 59:780–791 Tsaousis AD, Kunji ER, Goldberg AV, Lucocq JM, Hirt RP and Embley TM (2008) A novel route for ATP acquisition by the remnant mitochondria of Encephalitozoon cuniculi. Nature 453:553–556 Tyra HM, Linka M, Weber AP and Bhattacharya D (2007) Host origin of plastid solute transporters in the first photosynthetic eukaryotes. Genome Biol 8:R212 Undeen AH and Vander Meer RK (1999) Microsporidian intrasporal sugars and their role in germination. J Invertebr Pathol 73:294–302 Van de Peer Y, Ben Ali A and Meyer A (2000) Microsporidia: accumulating molecular evidence that a group of

36

Bryony A.P. Williams and Patrick J. Keeling

amitochondriate and suspectedly primitive eukaryotes are just curious fungi. Gene 246:1–8 van der Giezen M, Slotboom DJ, Horner DS, Dyal PL, Harding M, Xue GP, Embley TM, and Kunji ER (2002) Conserved properties of hydrogenosomal and mitochondrial ADP/ATP carriers: a common origin for both organelles. EMBO J 21:572–579 Va´vra J and Larsson JIR (1999) Structure of the Microsporidia. In: Wittner M and Weiss LM (ed) The Microsporidia and Microsporidiosis. American Society for Microbiology, Washington B. C., pp 7–84 Vivare`s CP, Gouy M, Thomarat F and Me´te´nier G (2002) Functional and evolutionary analysis of a eukaryotic parasitic genome. Curr Opin Microbiol 5:499–505 Vossbrinck CR, Maddox JV, Friedman S, DebrunnerVossbrinck BA and Woese CR (1987) Ribosomal RNA sequence suggests microsporidia are extremely ancient eukaryotes. Nature 326:411–414 Vossbrinck CR and Woese CR (1986) Eukaryotic ribosomes that lack a 5.8S RNA. Nature 320:287–288 Weber RW, Wakley GE, Thines E and Talbot NJ (2001) The vacuole as central element of the lytic system and sink for lipid droplets in maturing appressoria of Magnaporthe grisea. Protoplasma 216:101–112 Weidner E and Trager W (1973) Adenosine triphosphate in the extracellular survival of an intracellular parasite (Nosema michaelis, Microsporidia). J Cell Biol 57:586–591 Weisman LS (2003) Yeast vacuole inheritance and dynamics. Annu Rev Genet 37:435–460 Williams BA, Haferkamp I and Keeling PJ (2008a) An ADP/ATP-specific mitochondrial carrier protein in the microsporidian Antonospora locustae. J Mol Biol 375:1249–1257

Williams BA, Hirt RP, Lucocq JM and Embley TM (2002) A mitochondrial remnant in the microsporidian Trachipleistophora hominis. Nature 418:865–869 Williams BA, Slamovits CH, Patron NJ, Fast NM and Keeling PJ (2005) A high frequency of overlapping gene expression in compacted eukaryotic genomes. Proc Natl Acad Sci USA 102:10936–10941 Williams BAP, Cali A, Takvorian PM and Keeling PJ (2008b) Distinct localization patterns of two putative mitochondrial proteins in the microsporidian Encephalitozoon cuniculi. J Eukaryot Microbiol 55: 131–133 Williams BAP, Lee RH, Becnel JJ, Weiss LM, Fast NM and Keeling PJ (2008c) Genome sequence surveys of Brachiola algerae and Edhazardia aedis reveal microsporidia with low gene densities. BMC Genomics 9:200 Xu J, Pan G, Fang L, Li J, Tian X, Li T, Zhou Z and Xiang Z (2006) The varying microsporidian genome: existence of long-terminal repeat retrotransposon in domesticated silkworm parasite Nosema bombycis. Int J Parasitol 36:1049–1056 Xu J, Saunders CW, Hu P, Grant RA, Boekhout T, Kuramae EE, Kronstad JW, Deangelis YM, Reeder NL, Johnstone KR et al. (2007) Dandruff-associated Malassezia genomes reveal convergent and divergent virulence traits shared with plant and human fungal pathogens. Proc Natl Acad Sci USA 104:18730–18735 Yang ZH (1996) Among-site rate variation and its impact on phylogenetic analyses. Trends Ecol Evol 11:367–372 Yarlett N, Orpin CG, Munn EA, Yarlett NC and Greenwood CA (1986) Hydrogenosomes in the rumen fungus Neocallimastix patriciarum. Biochem J 236:729–739

3

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

MEREDITH D.M. JONES1, THOMAS A. RICHARDS1

CONTENTS I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Environmental DNA Analysis, the Clone Library Approach. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Fungal Molecular Diversity in Aquatic Environments using a Clone Library Approach . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Marine Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Deep-Sea Environments. . . . . . . . . . . . . . . . . . . . 3. Freshwater Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . 4. Novel Basal Branches in the Fungal Radiation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Fungal Diversity in Soil and Plant Environments using the Clone Library Approach . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Limitations of the Clone Library Approach . . III. Alternative Environmental DNA Methods. . . . . . . A. 454 Sequencing of Taxon Identification Sequence Tags from Environmental DNA . . . . B. Fluorescence In Situ Hybridisation of Uncultured Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Conclusions and Future Perspectives . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

37 38 42 42 43 44 44 45 46 47 47 48 49 50

I. Introduction Fungi play pivotal roles in global environments; they form mycorrhizal associations with land plants, symbioses with algae as lichens, are comensal microbes of animals, are the causal agents of numerous diseases of animal and plant species, and perform vital functions as decomposers to enable carbon and nutrient cycling. As such, the Fungi represent a significant proportion of the microbial diversity (Hawksworth 2001; Hibbett et al. 2007; Mueller et al. 2007), biomass and genomic variations found on Earth and include a number of very useful organisms. For example, fungi are the primary source for a number of leading pharmaceuticals (e.g., cholesterol-lowering 1

Natural History Museum, Cromwell Road, London, SW7 5BD, UK; e-mail: [email protected]

statins, the immunosuppressant cyclosporinA, several antibiotics including penicillin and cephalosporin). The evolutionary diversification of the Fungi has included several radical changes in cell biology, including loss of true mitochondria (Embley et al. 2003; van der Giezen et al. 2002), loss of phagotrophy and gain of osmotrophic mechanisms, loss or losses of a cilium (flagellum) (James et al. 2006a, b), gain of a rigid chitin rich cell wall (Bartnicki-Garcia 1987), development of filamentous structures, and the acquisition of saprotrophic and pathogenic lifestyles (James et al. 2006a). However, it is currently unclear when these major cellular changes occurred on the fungal phylogeny, and furthermore, the true extent and complexity of the fungal tree of life (James et al. 2006a; Liu et al. 2009). Previously, the diversity of the kingdom Fungi was loosely classified as four major groups: (i) the ascomycetes of the subkingdom Dikarya (filamentous or yeast forms that reproduce sexually with internal spore maturation within an ascus), (ii) basidiomycetes also of the subkingdom Dikarya (filamentous or yeast forms that reproduce sexually with external spore maturation on a basidium), (iii) zygomycetes (multinucleated cells which produce filaments and lack complex fruiting bodies), and (iv) chytrids (an informal name given to species which produce motile flagellated spores during their lifecycle) (Hibbett et al. 2007). However, this simplistic model of fungal taxonomy has been challenged at a number of levels. These revisions include the placement of the microsporidia with (Hirt et al. 1997; Hirt et al. 1999) and potentially within the Fungi (Adl et al. 2005; James et al. 2006a; Keeling 2003), division of the ‘chytrids’ and zygomycetes into multiple paraphyletic clades and subsequent taxonomic reclassifications (Hibbett et al. 2007; James et al. 2006b; Liu et al. 2009). All progress made in understanding the fungal tree of life and its taxonomy has, in Evolution of Fungi and Fungal-Like Organisms, The Mycota XIV S. Po¨ggeler and J. Wo¨stemeyer (Eds.) © Springer-Verlag Berlin Heidelberg 2011

38

Meredith D.M. Jones and Thomas A. Richards

the most part, been made by studying fungi that have been isolated and cultured. This, however, represents only a narrow perspective on the true evolutionary complexity of the Fungi. The number of fungal species present on Earth is unknown, however one estimate suggests a minimum of 1.5 million species (Hawksworth 1991). This estimate was primarily based on the diversity of fungi associated with land plants in specimens from Western Europe, which was then extrapolated to a global number using estimates of global plant diversity (Hawksworth 1991, 2001). This figure is controversial because it makes several risky assumptions and is primarily based only upon observations of fungal diversity associated with vascular plants in temperate regions. Such assumptions include the premise that the diversity of plant and fungal species are positively correlated and that fungal and plant species have comparable range sizes (May 1991, 1994; Muller and Schmit 2007). This has called some authors to suggest this figure may represent an overestimation of fungal diversity (May 1991, 1994). Yet in contrast, this figure is also suggested to be an underestimate of fungal diversity, because it is based on morphological identification and therefore does not account for cryptic species diversity (Hawksworth 2001). Furthermore, this estimate does not include the numerous alternative ecosystems where fungi are both abundant and diverse, suggesting that global fungal diversity may in fact be much greater than this estimate suggests. For example, a study by Suh et al. (2005) isolated 650 yeast species from a variety of beetles from different geographical regions. Sequencing of a 600 bp fragment of the large subunit (LSU) ribosomal RNA (rRNA) gene (D1/D2 loop) showed a dominance of ascomycete yeast among the isolated strains, 68% of which differed by >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

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

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-Garcı´a 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-Garcı´a 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.

41

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

42

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.

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

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

43

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).

44

Meredith D.M. Jones and Thomas A. Richards

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

50

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

40

“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

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

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

45

(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

46

Meredith D.M. Jones and Thomas A. Richards

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 <500 clones. Consequently, to our knowledge, no eukaryote-wide study, or fungus-specific study, has demonstrated sampling saturation (e.g., Edgcomb et al. 2002;

Jebaraj et al. 2009; Porter et al. 2008; Stoeck et al. 2006). This implies further sampling from the same environments would reveal additional diversity (Moreira and Lo´pez-Garcia 2002; Sogin et al. 2006; Stoeck et al. 2006). For example, a recent study of the Cariaco Basin, Venezuela, demonstrated that, in a comparison of three clone libraries, each derived from the same environmental DNA but using different primer combinations, and with a sampling of 241, 119, and 137 clones respectively, only three operational taxonomic units were present in all three clone libraries (Stoeck et al. 2006). This result demonstrates a very low overlap between the clone libraries and suggests that molecular diversity remains largely undersampled using these approaches. These problems mean that lineages which are locally rare or patchily distributed are likely to remain undetected; consequently, we currently know very little about organisms belonging to either category. Artefacts produced by incomplete sampling are further compounded, as generally only a single set of primers are chosen to amplify the targeted gene region, or indeed the DNA preparation methodology may favour certain species and exclude other groups. Furthermore, standard practice is to amplify as much of the SSU and/or ITS regions in order to acquire as many characters as possible for molecular phylogeny. However, it is known that the targeting of longer amplicons can reduce the recovery of diversity profiles in clone libraries (Huber et al. 2009). Additionally, ligation of amplicons and transformation protocols often result in the exclusion of rare amplicon types. The net effect of these limitations is that most environments remain under-sampled and meaningful comparisons of species diversity across differing environment samples are not possible. Environmental gene libraries generated from multiple group-specific primers provide a possible means of detecting low-abundance microbes, improving sampling efficiency and enabling systematic comparisons of microbial diversity between and within environments (e.g., Bass and Cavalier-Smith 2004; von der Heyden et al. 2004). However, it is important to note that environmental clone library methods are limited when trying to understand microbial community dynamics including activity, habitat distribution, and complete diversity profiles.

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

III. Alternative Environmental DNA Methods A. 454 Sequencing of Taxon Identification Sequence Tags from Environmental DNA Recently, Sogin and co-workers (Sogin et al. 2006) employed DNA sequencing using micro-fabricated high-density picolitre reactors (454 sequencing) to investigate the molecular diversity of prokaryotes in marine environments. This uses a relatively new technology where millions of nucleotide base pairs are sequenced with ‘99.5%’ accuracy in 100–500 bp fragments (Margulies et al. 2005) depending on which version of the technology is applied (e.g., GS-FLX or GS-FLX titanium). Consequently, the level of sequence sampling achieved is much greater than can be contemplated for standard environmental clone library analysis. The method begins as with clone library generation, by the amplification of a gene sequence of interest from environmental DNA (Fig. 3.1, right column) but steps 3–6 of the environmental clone library analysis are eliminated and replaced by the 454 protocol. Briefly: adapter sequences are incorporated on the 5’ region of the primer to enable the attachment of the PCR products to DNA capture beads. The adapters can be ligated to the PCR products or incorporated as part of the PCR primer. We recommend the latter to minimise sampling artefacts. The 454 method incorporates a one fragment per bead association and isolation process, followed by a PCR reaction mixture in oil emulsion, which in effect isolates single templates on each bead and then generates millions of copies per bead. The DNA strands are denatured and the beads, now carrying millions of identical single-stranded DNA molecules, are centrifuged into individual picolitre-sized wells (formed by a fibre-optic slide) and sequenced using pyrophosphate-based sequencing (Margulies et al. 2005). This method avoids the need to use a cloning step (Fig. 3.1), enabling PCR amplicons to be effectively sequenced directly, without cloning and transformation biases. This factor, combined with the very high number of sequence reads generated, increases the probability that low-abundance amplicons can be recovered, thus removing a strong potential source of artefact between environment sampling and sequence sampling.

47

In the first experiment of this kind Sogin and co-workers (Huber et al. 2007; Sogin et al. 2006) recovered 118778 prokaryote V6 SSU rDNA sequences demonstrating an ‘unprecedented level of bacterial complexity within marine samples’. This method has now been adapted for the analysis of eukaryotic microbial communities (Amaral-Zettler et al. 2009; Stoeck et al. 2009, 2010). For example, Stoeck et al. (2010) used a multiple marker approach targeting both the V9 and V4 variable regions of the SSU rDNA sequence in order to investigate the complexity of microbial eukaryotic communities in anoxic water from a Norwegian coastal fjord. This experiment demonstrated that eukaryote DNA diversity profiles, like prokaryotes, appear to be composed of a small pool of highly abundant genotypes, while the majority of the molecular diversity detected is present in very low numbers. This suggests the greater part of the microbial diversity present in these environments is at low abundance (Huber et al. 2007; Sogin et al. 2006). The idea that microbial communities are arranged in this manner is a popular concept in microbial ecology and is often termed the ‘rare biosphere’ (e.g., Pedro´sAlio´ 2006). The pattern of diversity observed for both prokaryote and eukaryote 454 datasets has been compared with this ecological concept (e.g., Sogin et al. 2006; Stoeck et al. 2010). The significance of the large diversity of sequence tags both in eukaryotes (Stoeck et al. 2010) and prokaryotes (Huber et al. 2007; Sogin et al. 2006) awaits further investigation and correction for factors that potentially artificially inflate the diversity profile detected (e.g., PCR and sequencing errors, levels of intragenomic SSU rDNA polymorphism, PCR chimaera formation). The eukaryotic V4 and V9 SSU 454 sequencing approach on the Norwegian coastal fjord samples recovered a number of sequences which cluster with or within the fungi (Stoeck et al. 2010). When the dataset was clustered at 98% similarity values, 0.71% of 3993 unique V4 sequence clusters and 3.8% of 2633 V9 sequence clusters were classified as fungi. As part of the study Stoeck et al. (2010) also compare this data with 1000 environmental SSU rDNA clones sequenced from the same environment. The clone library approach indicated an absence of the fungi within the costal marine water columns, consistent with the pattern identified from the review of

48

Meredith D.M. Jones and Thomas A. Richards

marine SSU rDNA literature discussed earlier (Massana and Pedro´s-Alio´ 2008). However, the 454 approach revealed the presence of a significant fungal community and suggests that deep sequencing initiatives, potentially in combination with fungal specific primers, will demonstrate an increasingly complex fungal community in marine environments. This hidden diversity may prove important for understanding the fungal tree of life. Similar methods have also been applied to investigate the diversity of fungal communities in soil and phyllosphere environments using fungal specific ITS PCR amplicon 454 sequencing. These studies used conservative sequence clustering of 95% (Jumpponen and Jones 2009) and 97% (Bue´e et al. 2009), but still demonstrated that both environment types harbour complex fungal communities. Both analyses note that accurate taxonomic classification was dominated by the Dikarya, with the phyloplane environments dominated by ascomycetes (93.4%; Jumpponen and Jones 2009), while the soil environment was dominated by basidiomycetes (44%; Bue´e et al. 2009). Interestingly, the analysis of soil environments using a filtered database containing only identified fungal strains and excluding uncultured environmental sequences enabled improved taxonomic classification and identified that 20% of the 454 ITS sequence tags recovered grouped as ‘unclassified Dikarya’, while 11% were grouped as ‘unclassified fungi’, with many of these novel sequences clustering with the soil clone group 1 investigated by Schadt et al. (2003) and Porter et al. (2008). This data again demonstrated that relatively well-sampled environments like forest soils harbour numerous unexplored fungal forms.

While the 454 approach enables us to better understand the breath of fungal diversity, and hints at the existence of a vast number of undetected species, classification of much of this novel diversity remains elusive due to the small size of the DNA sequence regions amplified and a lack of understanding of how this relates to wider genome/phenotype variation, intra-genomic rDNA variation, and inconsistent understanding of how measures of rDNA variation relate to species boundaries (Nilsson et al. 2006, 2008). Whilst such studies greatly enhance the discovery of novel species, drawbacks have been found. The generation of sequencing artefacts including ¨ pik et al. 2009) has homo-polymer read errors (O been reported as a problem for pyrosequencing and stringent sequence analyses have to be performed using conservative sequence clustering approaches (Kunin et al. 2009). Sequencing both strands and the incorporation of only sequences containing both primer sites is one approach that increases the reliability of the sequence data and can help with correction of homo-polymer read errors. The application of this technique to reveal the same novel sequences from multiple samples and multiple locations, and combining this technique with fluorescence in situ hybridisation (FISH) represent important additional experiments for the validation of novel phylogenetic groups.

Similar 454 methods have also been applied to target arbuscular mycorrhizal glomeromycete communities of forests where the method again proved itself as an effective technique for sampling increasing levels of fungal diversity. However, the vast majority of sequences detected could be classified as known taxa using 97% clustering thresholds. The majority of the remaining sequences were putatively novel sequences belonging to Glomus group A, however the authors suggest the novelty of these sequences could be the product of putative errors mainly in single nucleotide repeat tracks (homopolymer ¨ pik et al. 2009). 454 errors) of the recovered sequences (O The authors could identify this source of artefact because they were sampling within the confines of a relatively well-sampled group and were using strict sequences clustering criteria. This approach is not always possible

This method works through the illumination of target cells by the use of a fluorophore-labelled DNA oligonucleotide probe designed to be specific to the target group. As such the FISH method requires the prior discovery of rDNA sequences of the target group in the environment sampled, and has been successfully combined with SSU rDNA environmental clone library analysis. The FISH approach has been used to reveal the cellular forms represented by novel environmental sequences and has proved successful in identifying cells from the previously unknown MAST groups (Massana et al. 2002) discussed earlier, a marine

at wider taxonomic sampling scales but these results dem¨ pik onstrate an important concern relating to 454 data (O et al. 2009).

B. Fluorescence In Situ Hybridisation of Uncultured Fungi

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

picoplanktonic group – the ‘picobiliphytes’ (Not et al. 2007), and a highly diverse group of alveolate parasites present in marine environments (Chambouvet et al. 2008). This method is especially effective on aquatic environments where microbial communities of specific size ranges can be isolated using filtration. Briefly a unique 15–20 bp sequence motif is identified for the target group using comprehensive DNA alignment. These are often compared to rRNA accessibility models (e.g., Behrens et al. 2003) to identify a region with greater accessibility in the ribosome. A reverse complement 18 bp oligonucleotide is synthesised with a 5’ fluorophore attached. The target microbial community is usually fixed using formaldehyde (although other fixative approaches can be used and can minimise some autofluorescence signals), filtered using specific size selection onto filters (often polycarbonate filters), dried, cut into small sections, placed on glass slides, hybridised with FISH probes under a range of differing chemical or thermal stringency conditions, washed to remove unbound probes, mounted, and then imaged using epifluorescence microscopy (as described by Massana et al. 2002; Moter and Go¨bel 2000; Pernthaler et al. 2001). We note that the robust chitin cell wall often associated with the fungi can block FISH probes from entry into the cell and therefore prevent detection, therefore some researchers when looking at fungi often add a chitinase incubation wash step prior to probe hybridisation (Baschien et al. 2001). It is important also to add a range of control experiments, especially when targeting uncultured microbes. These can include testing for cross reactivity with nontarget cultured microbes that branch near the target group on phylogenetic trees, and repeating the hybridisation experiments using a range of stringency conditions. The FISH method is primarily useful for identifying the abundance of target communities within given environments, but it can also be combined with a range of additional cell stains to identify morphological and cellular characteristics. These include standard DNA stains such as 4’,6-diamidino-2-phenylindole (DAPI) and probidium iodide which are useful to confirm the target cells possess a nucleus and therefore demonstrate the probe is illuminating an eukaryotic cell rather than cross reacting with a prokaryote, or trapped probe within environmental debris. Furthermore, cell wall stains can be used to identify the presence

49

of chitin or other cell wall components (Jones et al. 2011). Subsequent hybridisation of probes to environmental samples enables visualisation of target cells under fluorescent light and reveals cell morphology. The approach has also been used to identify the ecological aspects of a target group. For example, FISH can be used to identify if the target group is an intracellular parasite of other organisms (Chambouvet et al. 2008). Furthermore, incubation of samples with fluorescently labelled bacteria, followed by fixation and FISH has been used to identify targeted cells as phagotrophic bacterial grazers (Massana et al. 2002). Whilst the presence of the chitin rich cell wall found surrounding most fungal cells might be a hindrance (Baschien et al. 2001), FISH has proven successful at targeting fungi from natural environments (Baschien et al. 2001; Kosse et al. 1997; Li et al. 1996). Excitingly these methods have been used to demonstrate subgroups of cryptomycota (discussed above) represented 4.5% of the eukaryotic community in a lake water environment, suggesting this novel group represents a significant component of this aquatic ecosystems. However, this technique is rarely applied when investigating novel fungal lineages identified using environmental sequencing and therefore these techniques await exploitation for investigating the ecology and cell biology of novel uncultured fungal forms.

IV. Conclusions and Future Perspectives Our knowledge of fungal diversity is expanding, facilitated by the use of environmental DNA methods. The detection of novel forms from habitats such as freshwater, marine, and soil environments has been discussed in this chapter. However, in many cases the detection of novel fungal diversity was part of a larger eukaryotic diversity study. There are of course exceptions, such as the recovery of novel yeasts from beetle guts (Suh et al. 2005), and the identification of novel ascomycete groups from soils (e.g., Porter et al. 2008), but such directed studies appear to be in the minority. These studies greatly enhance our knowledge of both diversity and fungal evolution. Of particular note is the cryptomycota clade (Jones et al. 2011), the putative primary branch in the fungal radiation. The discovery of such a diverse group is rare, and its exciting phylogenetic placement could

50

Meredith D.M. Jones and Thomas A. Richards

yield much information regarding the evolutionary history of the Fungi. The development of new sequencing technologies, such as 454 sequencing combined with environmental DNA PCR, enables sequencing of community diversity to a far greater depth than was previously possible. This methodology has enabled detection of large sequence diversity within both prokaryotic and eukaryotic communities (e.g., Sogin et al. 2006; Stoeck et al. 2010) and has revealed greater fungal diversity in marine environments than detected using standard clone library approaches. These results suggest fungal diversity is much greater than previously identified in many environments; hinting at the possibility of a large diversity of low abundance fungal forms which are yet to be explored (e.g., Bue´e et al. 2009; Jumpponen and Jones 2009; Stoeck et al. 2010). With the decrease in cost, large-scale sequencing efforts are becoming more financially viable and more readily implemented. Numerous primers for the direct amplification of fungal DNA from the environment have been developed and successfully applied; yet, to the best of our knowledge no large scale sequencing efforts based purely upon determining fungal diversity from multiple environments using multiple primer strategies have been undertaken. With the advancement in sequencing technologies this approach is now viable. Furthermore, until this approach has been applied, our understanding of the evolutionary complexity of the Fungi remains primarily based on cultured representatives, and therefore is likely to remain both biased and incomplete.

References Adl SM, Simpson AG, Farmer MA, Andersen RA, Anderson OR, Barta JR, Bowser SS, Brugerolle G, Fensome RA, Fredericq S et al. (2005) The new higher level classification of eukaryotes with emphasis on the taxonomy of protists. J Eukaryot Microbiol 52:399–451 Amaral Zettler LA, Go´mez F, Zettler E, Keenan BG, Amils R and Sogin ML (2002) Microbiology: eukaryotic diversity in Spain’s river of fire. Nature 417:137 Amaral-Zettler LA, McCliment EA, Ducklow HW and Huse SM (2009) A method for studying protistan diversity using massively parallel sequencing of V9 hypervariable regions of small-subunit ribosomal RNA genes. PLoS ONE 4:e6372 Anderson IC and Cairney JW (2004) Diversity and ecology of soil fungal communities: increased understanding

through the application of molecular techniques. Environ Microbiol 6:769–779 Anderson IC, Campbell CD and Prosser JI (2003) Potential bias of fungal 18S rDNA and internal transcribed spacer polymerase chain reaction primers for estimating fungal biodiversity in soil. Environ Microbiol 5:36–47 Arnold AE, Maynard Z, Gilbert GS, Coley PD and Kursar TA (2000) Are tropical fungal endophytes hyperdiverse? Ecology Letters 3:267–274 Bartnicki-Garcia S (1987) The cell wall: a crucial structure in fungal evolution. In: Rayner ADM, Brasier CM and Moore D (ed) Evolutionary Biology of the Fungi. Cambridge University Press, Cambridge, pp Baschien C, Manz W, Neu TR and Szewzyk U (2001) Fluorescence in situ hybrization of freshwater fungi. Internat. Rev. Hydrobiol. 86:371–381 Bass D and Cavalier-Smith T (2004) Phylum-specific environmental DNA analysis reveals remarkably high global biodiversity of Cercozoa (Protozoa). Int J Syst Evol Microbiol 54:2393–2404 Bass D, Howe A, Brown N, Barton H, Demidova M, Michelle H, Li L, Sanders H, Watkinson SC, Willcock S et al. (2007) Yeast forms dominate fungal diversity in the deep oceans. Proc Biol Sci 274:3069–3077 Behrens S, Ru¨hland C, Incio J, Huber H, Fonseca A, SpencerMartins I, Fuchs BM and Amann R (2003) In situ accessibility of small-subunit rRNA of members of the domains Bacteria, Archaea, and Eucarya to Cy3-labeled oligonucleotide probes. Appl Environ Microbiol 69:1748–1758 Berney C, Fahrni J and Pawlowski J (2004) How many novel eukaryotic ’kingdoms’? Pitfalls and limitations of environmental DNA surveys. BMC Biol 2:13 Brock PM, Do¨ring H and Bidartondo MI (2009) How to know unknown fungi: the role of a herbarium. New Phytol 181:719–724 Bruns TD and Gardes M (1993) Molecular tools for the identification of ectomycorrhizal fungi-taxon-specific oligonucleotide probes for suilloid fungi. Mol Ecol 2:233–242 Buchan A, Newell SY, Moreta JI and Moran MA (2002) Analysis of internal transcribed spacer regions of rRNA genes in fungal communities in a southeastern U.S. salt marsh. Microb Ecol 43:329–340 Bue´e M, Reich M, Murat C, Morin E, Nilsson RH, Uroz S and Martin F (2009) 454 Pyrosequencing analyses of forest soils reveal an unexpectedly high fungal diversity. New Phytol 184:449–456 Burgaud G, Le Calvez T, Arzur D, Vandenkoornhuyse P and Barbier G (2009) Diversity of culturable marine filamentous fungi from deep-sea hydrothermal vents. Environ Microbiol 11:1588–1600 Cavalier-Smith T (2004) Only six kingdoms of life. Proc R Soc Lond B Biol Sci 271:1251–1262 Chambouvet A, Morin P, Marie D and Guillou L (2008) Control of toxic marine dinoflagellate blooms by serial parasitic killers. Science 322:1254–1257 Cole JR, Chai B, Marsh TL, Farris RJ, Wang Q, Kulam SA, Chandra S, McGarrell DM, Schmidt TM, Garrity GM et al. (2003) The Ribosomal Database Project (RDP-II): previewing a new autoaligner that allows regular

Environmental DNA Analysis and the Expansion of the Fungal Tree of Life updates and the new prokaryotic taxonomy. Nucleic Acids Res 31:442–443 Damare S and Raghukumar C (2008) Fungi and macroaggregation in deep-sea sediments. Microb Ecol 56: 168–177 Damare S and Raghukumar C (2008) Fungi and macroaggregation in deep-sea sediments. Microb Ecol 56: 168–177 Dawson SC and Pace NR (2002) Novel kingdom-level eukaryotic diversity in anoxic environments. Proc Natl Acad Sci USA 99:8324–8329 DeSantis TZ, Hugenholtz P, Larsen N, Rojas M, Brodie EL, Keller K, Huber T, Dalevi D, Hu P and Andersen GL (2006) Greengenes, a chimera-checked 16S rRNA gene database and workbench compatible with ARB. Appl Environ Microbiol 72:5069–5072 Edgcomb VP, Kysela DT, Teske A, de Vera Gomez A and Sogin ML (2002) Benthic eukaryotic diversity in the Guaymas Basin hydrothermal vent environment. Proc Natl Acad Sci USA 99:7658–7662 Embley TM, van der Giezen M, Horner DS, Dyal PL and Foster P (2003) Mitochondria and hydrogenosomes are two forms of the same fundamental organelle. Philos Trans R Soc Lond B Biol Sci 358:191–201 Fisher CR, Takai K and Le Bris N (2007) Hydrothermal vent ecosystems. Oceanography 20:14–23 Gage JD and Tyler PA (1991) Deep-sea biology: a natural history of organisms at the deep-sea floor. Cambridge University Press, Cambridge Gardes M and Bruns TD (1993) ITS primers with enhanced specificity for basidiomycetes – application to the identification of mycorrhizae and rusts. Mol Ecol 2: 113–118 Giovannoni SJ, Britschgi TB, Moyer CL and Field KG (1990) Genetic diversity in Sargasso Sea bacterioplankton. Nature 345:60–63 Guillou L, Viprey M, Chambouvet A, Welsh RM, Kirkham AR, Massana R, Scanlan DJ and Worden AZ (2008) Widespread occurrence and genetic diversity of marine parasitoids belonging to Syndiniales (Alveolata). Environ Microbiol 10:3349–3365 Harrison KE (1990) The role of nutrition in maturation, reproduction and embryonic development of decapod crustaceans: a review. J Shellfish Res 9:1–28 Hawksworth DL (1991) The fungal dimension of biodiversity: magnitude, significance and conservation. Mycol Res 95:641–655 Hawksworth DL (2001) The magnitude of fungal diversity: the 1.5 million species estimate revisited. Mycol Res 105:1422–1432 Held AA (1981) Rozella and Rozellopsis: naked endoparasitic fungi which dress up as their hosts. The Botanical Review 47:451–515 Hibbett DS, Binder M, Bischoff JF, Blackwell M, Cannon PF, Eriksson OE, Huhndorf S, James T, Kirk PM, Lu¨cking R et al. (2007) A higher-level phylogenetic classification of the Fungi. Mycol Res 111:509–547 Hirt RP, Healy B, Vossbrinck CR, Canning EU and Embley TM (1997) A mitochondrial HSP70 orthologue in Vairimorpha necatrix: molecular evidence that microsporidia once contained mitochondria. Curr Biol 7:995–998

51

Hirt RP, Logsdon JM, Jr., Healy B, Dorey MW, Doolittle WF and Embley TM (1999) Microsporidia are related to Fungi: evidence from the largest subunit of RNA polymerase II and other proteins. Proc Natl Acad Sci USA 96:580–585 Horton TR and Bruns TD (2001) The molecular revolution in ectomycorrhizal ecology: peeking into the blackbox. Mol Ecol 10:1855–1871 Huber JA, Mark Welch DB, Morrison HG, Huse SM, Neal PR, Butterfield DA and Sogin ML (2007) Microbial population structures in the deep marine biosphere. Science 318:97–100 Huber JA, Morrison HG, Huse SM, Neal PR, Sogin ML and Mark Welch DB (2009) Effect of PCR amplicon size on assessments of clone library microbial diversity and community structure. Environ Microbiol 11:1292–1302 Huber T, Faulkner G and Hugenholtz P (2004) Bellerophon: a program to detect chimeric sequences in multiple sequence alignments. Bioinformatics 20:2317–2319 Hugenholtz P, Goebel BM and Pace NR (1998) Impact of culture-independent studies on the emerging phylogenetic view of bacterial diversity. J Bacteriol 180:4765–4774 Hugenholtz P and Huber T (2003) Chimeric 16S rDNA sequences of diverse origin are accumulating in the public databases. Int J Syst Evol Microbiol 53:289–293 James TY, Kauff F, Schoch CL, Matheny PB, Hofstetter V, Cox CJ, Celio G, Gueidan C, Fraker E, Miadlikowska J et al. (2006a) Reconstructing the early evolution of Fungi using a six-gene phylogeny. Nature 443:818–822 James TY, Letcher PM, Longcore JE, Mozley-Standridge SE, Porter D, Powell MJ, Griffith GW and Vilgalys R (2006b) A molecular phylogeny of the flagellated fungi (Chytridiomycota) and description of a new phylum (Blastocladiomycota). Mycologia 98:860–871 Jebaraj CS, Raghukumar C, Behnke A and Stoeck T (2009) Fungal diversity in oxygen-depleted regions of the Arabian Sea revealed by targeted environmental sequencing combined with cultivation. FEMS Microbiol Ecol 71:399–412 Jones MDM, Forn I, Gadelha C, Egan MJ, Bass D, Massana R and Richards TA (2011) Discovery of novel intermediate forms redefines the fungal tree of life. Nature: Advance Online Publication, doi:10.1038/ nature09984 Jumpponen A and Jones KL (2009) Massively parallel 454 sequencing indicates hyperdiverse fungal communities in temperate Quercus macrocarpa phyllosphere. New Phytol 184:438–448 Keeling PJ (2003) Congruent evidence from alpha-tubulin and beta-tubulin gene phylogenies for a zygomycete origin of microsporidia. Fungal Genet Biol 38: 298–309 Kis-Papo T (2005) Marine fungal communities. In: Dighton J, White JF and Oudemans P (ed) The fungal community: Its organisation and role in the ecosystem. Taylor and Francis Group, Boca Raton, pp Kohlmeyer J and Kohlmeyer E (1979) Marine mycology: the higher fungi. Academic Press, New york

52

Meredith D.M. Jones and Thomas A. Richards

Kosse D, Amann RI, Ludwig W and Scherer S (1997) Identification of yoghurt-spoiling yeasts with 18S rRNA-targeted oligonucleotide probes. Syst Appl Microbiol 20:468–480 Kunin V, Engelbrektson A, Ochman H and Hugenholtz P (2009) Wrinkles in the rare biosphere: pyrosequencing errors can lead to artificial inflation of diversity estimates. Environ Microbiol 12:118–123 Lara E, Moreira D and Lo´pez-Garcia P (2010) The environmental clade LKM11 and Rozella form the deepest branching clade of Fungi. Protist 161:116–121 Lawley B, Ripley S, Bridge P and Convey P (2004) Molecular analysis of geographic patterns of eukaryotic diversity in Antarctic soils. Appl Environ Microbiol 70:5963–5972 Le Calvez T, Burgaud G, Mahe´ S, Barbier G and Vandenkoornhuyse P (2009) Fungal diversity in deep-sea hydrothermal ecosystems. Appl Environ Microbiol 75: 6415–6421 Lefe`vre E, Bardot C, Noe¨l C, Carrias JF, Viscogliosi E, Amblard C and Sime-Ngando T (2007) Unveiling fungal zooflagellates as members of freshwater picoeukaryotes: evidence from a molecular diversity study in a deep meromictic lake. Environ Microbiol 9: 61–71 Lefe`vre E, Roussel B, Amblard C and Sime-Ngando T (2008) The molecular diversity of freshwater picoeukaryotes reveals high occurrence of putative parasitoids in the plankton. PLoS ONE 3:e2324 Lefranc M, The´not A, Lepe`re C and Debroas D (2005) Genetic diversity of small eukaryotes in lakes differing by their trophic status. Appl Environ Microbiol 71: 5935–5942 Lepe`re C, Boucher D, Jardillier L, Domaizon I and Debroas D (2006) Succession and regulation factors of small eukaryote community composition in a lacustrine ecosystem (Lake Pavin). Appl Environ Microbiol 72: 2971–2981 Lesaulnier C, Papamichail D, McCorkle S, Ollivier B, Skiena S, Taghavi S, Zak D and van der Lelie D (2008) Elevated atmospheric CO2 affects soil microbial diversity associated with trembling aspen. Environ Microbiol 10:926–941 Li S, Cullen D, Hjort M, Spear R and Andrews JH (1996) Development of an oligonucleotide probe for Aureobasidium pullulans based on the small-subunit rRNA gene. Appl Environ Microbiol 62:1514–1518 Liu Y, Steenkamp ET, Brinkmann H, Forget L, Philippe H and Lang BF (2009) Phylogenomic analyses predict sistergroup relationship of nucleariids and fungi and paraphyly of zygomycetes with significant support. BMC Evol Biol 9:272 Lo´pez-Garcı´a P, Rodriguez-Valera F, Pedro´s-Alio´ C and Moreira D (2001) Unexpected diversity of small eukaryotes in deep-sea Antarctic plankton. Nature 409:603–607 Lo´pez-Garcia P, Vereshchaka A and Moreira D (2007) Eukaryotic diversity associated with carbonates and fluid-seawater interface in Lost City hydrothermal field. Environ Microbiol 9:546–454 Luo Q, Krumholz LR, Najar FZ, Peacock AD, Roe BA, White DC and Elshahed M (2005) Diversity of the

microeukaryotic community in sulfide-rich Zodletone Spring (Oklahoma). Appl Environ Microbiol. 71: 6175–6184 Maidak BL, Olsen GJ, Larsen N, Overbeek R, McCaughey MJ and Woese CR (1996) The Ribosomal Database Project (RDP). Nucleic Acids Res 24:82–85 Mann KH (1988) Production and use of detritus in various freshwater, estuarine, and coastal marine ecosystems. Limnology and Oceanography 33:910–930 Margulies M, Egholm M, Altman WE, Attiya S, Bader JS, Bemben LA, Berka J, Braverman MS, Chen YJ, Chen Z et al. (2005) Genome sequencing in microfabricated high-density picolitre reactors. Nature 437:376–380 Massana R, Castresana J, Balague´ V, Guillou L, Romari K, Groisillier A, Valentin K and Pedro´s-Alio´ C (2004) Phylogenetic and ecological analysis of novel marine stramenopiles. Appl Environ Microbiol 70:3528–3534 Massana R, Guillou L, Diez B and Pedro´s-Alio´ C (2002) Unveiling the organisms behind novel eukaryotic ribosomal DNA sequences from the ocean. Appl Environ Microbiol 68:4554–4558 Massana R and Pedro´s-Alio´ C (2008) Unveiling new microbial eukaryotes in the surface ocean. Curr Opin Microbiol 11:213–218 May RM (1991) Fondness for Fungi. Nature 352:475–476 May RM (1994) Conceptual aspects of quantification of the extent of biological diversity. Philos Trans R Soc Lond B Biol Sci 345:13–20 Moon-van der Staay SY, De Wachter R and Vaulot D (2001) Oceanic 18S rDNA sequences from picoplankton reveal unsuspected eukaryotic diversity. Nature 409:607–610 Moreira D and Lo´pez-Garcia P (2002) The molecular ecology of microbial eukaryotes unveils a hidden world. Trends Microbiol 10:31–38 Moter A and Go¨bel UB (2000) Fluorescence in situ hybridization (FISH) for direct visualization of microorganisms. J Microbiol Methods 41:85–112 Mueller GM, Schmit JP, Leacock PR, Buyck B, Cifuentes J, Desjardin DE, Halling RE, Hjortstam K, Iturriaga T. Larsson KH, et al. (2007) Global diversity and distribution of macrofungi. Biodivers. Conserv. 16:37–48 Muller GM and Schmit JP (2007) Fungal biodiversity: what do we know? what we can we predict. Biodivers.Conserv. 16:1–5 Newell SY (1996) Established and potential impacts of eukaryotic mycelial decomposers in marine/terrestrial ecotones. J Exp Mar Biol Ecol 200:187–206 Nilsson RH, Kristiansson E, Ryberg M, Hallenberg N and Larsson KH (2008) Intraspecific its variability in the kingdom fungi as expressed in the international sequence databases and its implications for molecular species identification. Evol Bioinform Online 4: 193–201 Nilsson RH, Ryberg M, Kristiansson E, Abarenkov K, Larsson KH and Ko˜ljalg U (2006) Taxonomic reliability of DNA sequences in public sequence databases: a fungal perspective. PLoS ONE 1:e59 Not F, Valentin K, Romari K, Lovejoy C, Massana R, To¨be K, Vaulot D and Medlin LK (2007) Picobiliphytes: a marine picoplanktonic algal group with unknown affinities to other eukaryotes. Science 315:253–255

Environmental DNA Analysis and the Expansion of the Fungal Tree of Life O’brien HE, Parrent JL, Jackson JA, Moncalvo J-M and Vilgalys R (2005) Fungal community analysis by large-scale sequencing of environmental samples. Appl Environ Microbiol 71:5544–5550 Olsen GJ, Lane DJ, Giovannoni SJ, Pace NR and Stahl DA (1986) Microbial ecology and evolution: a ribosomal RNA approach. Annu Rev Microbiol 40:337–365 Olsen GJ and Woese CR (1993) Ribosomal RNA: a key to phylogeny. Faseb J. 7:113–123 ¨ pik M, Metsis M, Daniell TJ, Zobel M and Moora M O (2009) Large-scale parallel 454 sequencing reveals host ecological group specificity of arbuscular mycorrhizal fungi in a boreonemoral forest. New Phytol 184:424–437 Pace NR (1997) A molecular view of microbial diversity and the biosphere. Science 276:734–740 Pedro´s-Alio´ C (2006) Marine microbial diversity: can it be determined? Trends Microbiol 14:257–263 Pernthaler J, Glo¨ckner FO, Scho¨nhuber W and Rudolf A (2001) Fluorescence in situ hybridization (FISH) with rRNA-targeted oligonucleotide probes. In: Paul JH (ed) Marine Microbiology. Academic Press, London, pp Porter TM, Schadt CW, Rizvi L, Martin AP, Schmidt SK, Scott-Denton L, Vilgalys R and Moncalvo JM (2008) Widespread occurrence and phylogenetic placement of a soil clone group adds a prominent new branch to the fungal tree of life. Mol Phylogenet Evol 46:635–644 Raghukumar S (2004) The role of fungi in marine detrital processes. In: Ramaiah N (ed) Marine Microbiology: Facets & Opportunities. National Institute of Oceanography Goa, pp 91–101 Richards TA and Bass D (2005) Molecular screening of free-living microbial eukaryotes: diversity and distribution using a meta-analysis. Curr Opin Microbiol 8: 240–252 Richards TA, Vepritskiy AA, Gouliamova D and Nierzwicki-Bauer SA (2005) The molecular diversity of freshwater picoeukaryotes from an oligotrophic lake reveals diverse, distinctive and globally dispersed lineages. Environ Microbiol 7:1413–1425 Robison-Cox JF, Bateson MM and Ward DM (1995) Evaluation of nearest-neighbor methods for detection of chimeric small-subunit rRNA sequences. Appl Environ Microbiol 61:1240–1245 Schadt CW, Martin AP, Lipson DA and Schmidt SK (2003) Seasonal dynamics of previously unknown fungal lineages in tundra soils. Science 301:1359–1361 Seifert KA, Samson RA, Dewaard JR, Houbraken J, Le´vesque CA, Moncalvo JM, Louis-Seize G and Hebert PD (2007) Prospects for fungus identification using CO1 DNA barcodes, with Penicillium as a test case. Proc Natl Acad Sci USA 104:3901–3906 Slapeta J, Moreira D and P. L-G (2005) The extent of protist diversity: insights from molecular ecology of freshwater eukaryotes. Proc R Soc B 272:2073–2081 Sogin ML, Elwood HJ and Gunderson JH (1986) Evolutionary diversity of eukaryotic small subunit rRNA genes. Proc Natl Acad Sci USA 83:1383–1387 Sogin ML, Morrison HG, Huber JA, Mark Welch D, Huse SM, Neal PR, Arrieta JM and Herndl GJ (2006) Micro-

53

bial diversity in the deep-sea and the underexplored “rare biosphere”. Proc Natl Acad Sci USA 103: 12115–12120 Stoeck T, Bass D, Nebel M, Christen R, Jones MD, Breiner HW and Richards TA (2010) Multiple marker parallel tag environmental DNA sequencing reveals a highly complex eukaryotic community in marine anoxic water. Mol Ecol 19 Suppl 1:21–31 Stoeck T, Behnke A, Christen R, Amaral-Zettler L, Rodriguez-Mora MJ, Chistoserdov A, Orsi W and Edgcomb VP (2009) Massively parallel tag sequencing reveals the complexity of anaerobic marine protistan communities. BMC Biol 7:72 Stoeck T and Epstein S (2003) Novel eukaryotic lineages inferred from small-subunit rRNA analyses of oxygendepleted marine environments. Appl Environ Microbiol 69:2657–2663 Stoeck T, Hayward B, Taylor GT, Varela R and Epstein SS (2006) A multiple PCR-primer approach to access the microeukaryotic diversity in environmental samples. Protist 157:31–43 Suh SO, McHugh JV, Pollock DD and Blackwell M (2005) The beetle gut: a hyperdiverse source of novel yeasts. Mycol Res 109:261–265 Takishita K, Tsuchiya M, Reimer JD and Maruyama T (2006) Molecular evidence demonstrating the basidiomycetous fungus Cryptococcus curvatus is the dominant microbial eukaryote in sediment at the Kuroshima Knoll methane seep. Extremophiles 10: 165–169 Van de Peer Y, Jansen J, De Rijk P and De Wachter R (1997) Database on the structure of small ribosomal subunit RNA. Nucleic Acids Res 25:111–116 van der Giezen M, Slotboom DJ, Horner DS, Dyal PL, Harding M, Xue GP, Embley TM and Kunji ER (2002) Conserved properties of hydrogenosomal and mitochondrial ADP/ATP carriers: a common origin for both organelles. Embo J 21:572–579 Van Dover CL, Ward ME, Scott JL, Underdown J, Andersen B, Gustafson C, Whalen M and Carnegia RB (2007) A fungal epizootic in mussels at a deep-sea hydrothermal vent. Marine Ecology 28:54–62 van Hannen EJ, Mooij W, van Agterveld MP, Gons HJ and Laanbroek HJ (1999) Detritus-dependent development of the microbial community in an experimental system: qualitative analysis by denaturing gradient gel electrophoresis. Appl Environ Microbiol 65:2478–2484 Vandenkoornhuyse P, Baldauf SL, Leyval C, Straczek J and Young JPW (2002) Extensive fungal diversity in plant roots. Science 295:2051 Vilgalys R (2003) Taxonomic misidentification in public DNA databases. New Phyt. 160:4–5 von der Heyden S, Chao EE and Cavalier-Smith T (2004) Genetic diversity of goniomonads: an ancient divergence between marine and freshwater species. Eur J Phycol 39:343–350 White TJ, Bruns TD, Lee IY and Taylor JW (1990) Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenies. In: Innis MA, Gelfand DH, Sninsky JJ and White T (ed) PCR protocols:

54

Meredith D.M. Jones and Thomas A. Richards

a guide to methods and amplifications. Academic Press, San Diego, pp Winker S and Woese CR (1991) A definition of the domains Archaea, Bacteria and Eucarya in terms of small subunit ribosomal RNA characteristics. Syst Appl Microbiol 14:305–310 Winker S and Woese CR (1991) A definition of the domains Archaea, Bacteria and Eucarya in terms of small subunit ribosomal RNA characteristics. Syst Appl Microbiol 14:305–310

Woese CR, Kandler O and Wheelis ML (1990) Towards a natural system of organisms: proposal for the domains Archaea, Bacteria, and Eucarya. Proc Natl Acad Sci USA 87:4576–4579 Wuyts J, De Rijk P, Van de Peer Y, Pison G, Rousseeuw P and De Wachter R (2000) Comparative analysis of more than 3000 sequences reveals the existence of two pseudoknots in area V4 of eukaryotic small subunit ribosomal RNA. Nucleic Acids Res 28: 4698–4708

Evolution of Signalling in Fungi and Fungal-Like Organisms

.

4

Evolution of Signalling and Morphogenesis in the Dictyostelids

PAULINE SCHAAP1

CONTENTS I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Development of Dictyostelium discoideum . . . . . . A. Cell-Type Specialization and Morphogenesis 1. Aggregation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Formation and Migration of Slugs . . . . . . . . 3. Fruiting Body Construction. . . . . . . . . . . . . . . . B. Signals that Control D. discoideum Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Signalling before and during Aggregation 2. Signalling in Mounds and Slugs . . . . . . . . . . . 3. Signalling during Fruiting Body Formation 4. Signalling in the Spore Head. . . . . . . . . . . . . . . III. Evolution of Morphogenesis and Developmental Signalling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Evolution of Morphology. . . . . . . . . . . . . . . . . . . . . . B. Evolution of Developmental Signalling . . . . . . . IV. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

57 58 58 58 59 60 61 61 62 62 64 64 64 66 68 68

I. Introduction Similar to fungi, dictyostelids build fruiting bodies for easy dispersal of spores. However, they are only distant relatives. The dictyostelids or social amoebas are members of the amoebozoans, a sister group to opisthokonts, the group that contains the animals and fungi (Baldauf et al. 2000; see Chapter 1 in this volume). Unlike fungi, which feed by adsorption of low molecular weight compounds, dictyostelids are predators that engulf bacteria. Other differences are the absence of a cell wall during most of the dictyostelid life cycle and the fact that multicellularity is achieved not by cell division but by cell aggregation. In fact, multicellularity and morphogenesis in dictyostelids are highly dependent on the fact that their unwalled amoeboid cells remain highly motile until the fruiting body is completed. 1

College of Life Sciences, University of Dundee, MSI/WTB/JBC Complex, Dow Street, Dundee DD1 5EH, UK; e-mail: p.schaap@ dundee.ac.uk

Dictyostelids are found in all types of soils including those in arctic regions and above the tree line. However, species diversity is higher in forest soils and particularly those at tropical latitudes (Swanson et al. 1999). As single-celled amoebae they feed on bacteria that live on leaf detritus or animal waste. They have a highly efficient machinery for engulfing and digesting bacteria, at a rate of about 300 bacteria cell–1 h–1 (Peracino et al. 1998). Amoebas feed on many different bacterial species, but can also be invaded by and serve as host for bacterial pathogens (Steinert and Heuner 2005). Around hundred dictyostelid species have been isolated that grow under laboratory conditions on agar substrata. Some species, such as Dictyostelium deminutivum (Anderson et al. 1968) and D. menorah (Vadell and Cavender 2007) only form fruiting bodies efficiently when co-cultured with a fungus, indicating that symbiotic relationships between fungi and dictyostelids may exist. Multicellularity is a response to nutrient depletion, which triggers regulated secretion of a chemoattractant by the amoebas. Cells aggregate to form a mound, which can consist of up to a million amoebas. Sophisticated cell–cell signalling mechanisms between the amoebas orchestrate the differentiation of up to five different cell types and coordinate an intricate progression of cell movements. In combination with the synthesis of a flexible skin-like matrix, cell differentiation and cell movement first generate the formation of a motile structure, called the “slug”. The slug is attracted by light and warmth, which cause it to move to the soil’s top layer. Here, the slug cell mass projects upwards and forms the fruiting body, which consists of a column of stalk cells topped by a globular mass of spores. Depending on the species, the stalk can show different patterns of side branches and/or be decorated with disc-, root- or cup-shaped support structures (Raper 1984). Evolution of Fungi and Fungal-Like Organisms, The Mycota XIV S. Po¨ggeler and J. Wo¨stemeyer (Eds.) © Springer-Verlag Berlin Heidelberg 2011

58

Pauline Schaap

Many dictyostelid species show two other survival strategies: the microcyst, where individual amoebae encapsulate without aggregation, and the macrocyst, a sexual structure. The microcyst is an evolutionary relic from the dictyostelid ancestors, the solitary amoebas. Together with many other protist phyla, solitary amoebas usually encyst individually when faced with food depletion or other forms of stress (Gutierrez et al. 1990; Eichinger 2001; Lauwaet et al. 2007). Macrocyst formation starts with the fusion of two haploid amoebas to form a diploid zygote, which then attracts other amoebas by chemotaxis. Most of these cells are cannibalized by the zygote and their resources are used to build a thick wall around the structure. The zygote goes through sexual recombination and rounds of meiotic and mitotic division and eventually numerous haploid cells emerge (Raper 1984). Although the macrocyst stage should allow application of traditional genetic approaches to Dictyostelium research, its usefulness is hampered by its slow and inefficient germination. The developmental process of Dictyostelium with its strong dependence on coordinated cell migration is more reminiscent to animal development than fungal or plant development, which depend mostly on coordinated outgrowth. The ease of culture and genetic manipulation of D. discoideum, combined with its excellent accessibility to biochemical and cell biological approaches has therefore encouraged many workers to use this organism as a model for studies of chemotaxis, cell migration, developmental cell signalling and cell-type specialization, and cellular processes such as vesicle trafficking, phagocytosis and cytokinesis. More recently, dictyostelids have also become popular to resolve evolutionary questions. Most modern multicellular organisms start life as a fertilized egg, which through coordinated cell division, cell differentiation and cell migration, generates structures, organs and tissues that consist of millions of cells. Although these cells may vary greatly in form and function, they are all genetically identical. Social amoeba become multicellular by aggregating to form colonies. Here individuals cooperate to build a structure that serves to propagate only a fraction of their number. Since the individual cells need not necessarily be genetically identical, the genes of the nonpropagating cells are lost. This raises the question

whether such cells are either being altruistic or being exploited, and how cells avoid being the losers of the colony (Strassmann et al. 2000; Castillo et al. 2005). This is an important issue since modern multicellular organisms may have arisen through colony formation. Sponges, the earliest animals, arose through colony formation of unicellular filter-feeding choanoflagellates, which as choanocytes still line their internal cavities (King 2004). The dicytostelids with their combined singleand multi-celled life style are also very suited to answer another fundamental question. What caused multicellularity to evolve in the first place? Multicellular organisms are typically dependent on highly orchestrated signalling between cells, whereas unicellular organisms respond mainly to environmental signals. A major aspect of the evolution of multicellularity is therefore how the sensory systems of unicellular organisms were adapted to allow for more sophisticated signalling between cells. In this chapter, I will first present an overview of the signalling mechanisms that control development of the model organism D. discoideum. Second, I will summarize how phenotypic complexity evolved in the dictyostelids and highlight changes in developmental signalling processes that may have caused novel forms and behaviours to appear.

II. Development of Dictyostelium discoideum A. Cell-Type Specialization and Morphogenesis 1. Aggregation Upon nutrient depletion many dictyostelid species have a choice of three alternative pathways. They can aggregate to form fruiting bodies, encyst individually, or fuse with a mate and then attract cells to form a macrocyst. Apart from starvation there are apparently other cues to guide their choice. In general terms, light and low humidity favour fruiting body formation, while macrocyst formation occurs under dark and humid conditions, using ethylene as a specific trigger (Chang et al. 1983; Amagai 1984). Microcyst formation is promoted by high osmolyte or high ammonia levels (Toama and Raper 1967; Lonski 1976).

Evolution of Signalling and Morphogenesis in the Dictyostelids

59

D. discoideum has lost the microcyst pathway, but nevertheless depends on other signals besides starvation to initiate development. Two of these signals are the glycoproteins prestarvation factor (PSF) and conditioned medium factor (CMF). They act as quorum sensors; cells sense that a cell density appropriate for aggregation has been reached, when the PSF and CMF concentrations reach a specific threshold. The response to PSF is inhibited by bacteria and allows cells to monitor their own density relative to food. CMF acts later and is secreted by starving cells (Clarke and Gomer 1995). PSF stimulates basal expression of genes that encode proteins required for aggregation, such as the G protein-coupled cAMP receptor, cAR1, that detects the chemoattractant cyclic AMP (cAMP), and the extracellular cAMP phosphodiesterase PdsA (Rathi et al. 1991). CMF contributes to this response, but also directly facilitates cAMP signalling. Once basal levels of cAMP signalling genes are expressed, some cells start to spontaneously

secrete pulses of cAMP. In surrounding cells this evokes two responses: (i) extension of a pseudopod and movement towards the cAMP source and (ii) synthesis and secretion of a cAMP pulse. The latter response causes the original pulse to travel outward through the population and the former causes the cells to move inwards and collect at the oscillating centre (Fig. 4.1). In addition to mediating aggregation, the cAMP pulses also serve another function. They trigger rapid upregulation of all proteins that are involved in the aggregation process, and induce competence for the subsequent stages of development (Gerisch et al. 1975; Schaap et al. 1986).

Fig. 4.1. Life cycle of Dictyostelium discoideum. D. discoideum amoebas emerge from spores and start feeding on bacteria. When food runs out, amoebas secrete cAMP pulses, which trigger chemotactic movement and aggregation of cells. Once aggregated, the amoebas differentiate into prestalk and prespore cells in a regulated ratio. The organizing tip continues to emit cAMP pulses, which shape the cell mass by coordinating cell movement. The cAMP pulses also cause the prestalk cells, which are che-

motactically most responsive, to move towards the front. At the posterior, prespore cells dedifferentiate into anterior-like cells, which replenish the prestalk cells when needed. At the onset of culmination the cells synthesize a cellulose tube, the apical prestalk cells move into the tube and mature into stalk cells, the remaining prestalk cells form support structures, such as the upper and lower cups and the basal disk. The prespore cells move up the stalk and mature into spores

2. Formation and Migration of Slugs The formation of well proportioned fruiting bodies from loose heaps of cells depends on an interplay between cell differentiation and cell movement. Cell movement remains under control of cAMP signalling. The top of the aggregate takes over the

60

Pauline Schaap

function of the aggregation centre and emits waves of cAMP (Dormann et al. 1998). Cells become more adhesive and secrete matrix proteins and cellulose, which form an elastic sheath around the aggregate (Wilkins and Williams 1995). The cells, which are now confined, can only go upwards in response to cAMP waves and form the sausage-shaped “slug”. Gravity causes the slugs to topple over and they start migrating towards light and warmth (Bonner et al. 1950). In nature, this serves to reach the top layer of the soil, the most propitious spot for fruiting body formation and spore dispersal (Bonner and Lamont 2005). Meanwhile, cells are differentiating into spore and stalk precursors. In the mound, these cells types appear at first intermixed (Fig. 4.1). Intrinsic factors, such as glycogen content and cell cycle phase at the onset of starvation influence the initial choice in such a manner that the prespore cells differentiate from the largest, best fed cells and the prestalk cells from their leaner brethren (Leach et al. 1973; Weijer et al. 1984). In a starving population the leaner cells are also the first to initiate aggregation and to express aggregationassociated genes (Wang et al. 1988a). The prespore cells largely loose the properties of aggregating cells, but the prestalk cells retain the ability to sense and propagate cAMP waves (Matsukuma and Durston 1979; Schaap and Spek 1984; Verkerke-van Wijk et al. 2001). They therefore move with greatest efficiency towards the oscillating tip, thus creating an anterior–posterior prestalk–prespore pattern (Sternfeld and David 1981). The differentiation of the prespore cells involves the expression of many spore-coat genes (Fosnaugh and Loomis 1991); the cells fill up with Golgi-derived vesicles that bear a thick internal coating of spore-coat glycoproteins and contain precursors and enzymes for complete spore coat synthesis (West 2003). The prestalk cells express genes encoding sheath and stalk matrix proteins (McRobbie et al. 1988) and are characterized by autophagic vacuoles (Schaap 1983; Yamamoto and Takeuchi 1983). These vacuoles will later fuse to form the large central vacuole of the stalk cells. The prestalk cells also provide most of the motive force in slugs (Inouye 1984). This and synthesis of the cellulose stalk cell wall require extensive energy and materials resources, making autophagy an essential process for development (Otto et al. 2003). Many prestalk

cells are used up during slug migration and their corpses are discarded from the slug’s rear (Smith and Williams 1981; Fig. 4.1). The prestalk region is replenished by the forward movement of anterior-like cells (ALCs) that are generated by dedifferentiation of the leanest prespore cells (Sternfeld and David 1982; Thompson and Kay 2000a). In early slugs, the ALCs are inhibited to move forward by a signal secreted by prestalk cells. There is good evidence that this signal is ammonia, an end-product of protein degradation (Feit et al. 1990, 2001). When the prestalk region becomes depleted, less ammonia is being produced and ALCs move forward to replace the lost prestalk cells. This process ensures that individual prespore cells retain their resources for spore survival, while appropriate proportions of prestalk and prespore cells for fruiting body construction are maintained as slugs become progressively smaller during prolonged migration.

3. Fruiting Body Construction Incident light and low local ammonia levels trigger fruiting body formation (Schindler and Sussman 1977). The slug contracts and points its tip upwards (Fig. 4.1). Prestalk cells start to synthesize a central cellulose tube (Raper and Fennell 1952). They continue their movement towards the cAMP waves emitted by the tip, but now move into the upper open end of the stalk tube and differentiate into stalk cells. This involves synthesis of a cellulose cell wall, uptake of water, formation of one large central vacuole and eventually cell death. The prespore cells move up the newly formed stalk and mature into spores, starting from the prestalk boundary downwards. Anterior-like cells sort both downwards and upwards from the prespore region (Fig. 4.1). The downward moving cells form a basal disc surrounding the base of the stalk and a lower cup that supports the spore head, while the upward moving cells cap the spore head from above as the upper cup. Spore maturation is a rapid process that involves fusion of prespore vesicles with the plasma membrane. This brings the first layer of the spore coat in situ at the surface, while the remaining two layers are rapidly being produced by ejected enzymes and prefabricated materials (West 2003).

Evolution of Signalling and Morphogenesis in the Dictyostelids

B. Signals that Control D. discoideum Development 1. Signalling before and during Aggregation

61

Schaap 1995). The second quorum signal CMF affects early gene expression through another route, which involves the receptor CMFR1 (Deery et al. 2002). The biochemical network that enables pulsatile cAMP signalling involves the following sequence of events (Fig. 4.2):

The most remarkable aspect of D. discoideum development is the very dominant role of cAMP. As a secreted signal it controls cell movement and differentiation throughout the developmental programme, but in its more common role as intracellular messenger, it mediates the effect of many other developmental signals. The main intracellular target for cAMP is cAMP-dependent protein kinase (PKA), which like the fungal PKA consists of a single catalytic (PKA-C) and a single regulatory subunit (PKA-R; Mutzel et al. 1987). During most of the life cycle PKA is activated in the manner similar to other eukaryotes. cAMP binds to PKA-R, which causes PKA-R to dissociate from PKA-C, leaving PKA-C in its active form. However, in early development PKA-C is regulated at the translational level. In feeding cells PKA-C translation is inhibited by binding of the translational repressor PufA to the 3’ region of PKA-C mRNA (Souza et al. 1999). Upon starvation this repression is relieved by YakA, a member of a deeply conserved protein kinase family that also regulates the decision between growth and differentiation in animals and fungi (Hartley et al. 1994; Souza et al. 1998; Mercer and Friedman 2006). Active PKA is essential for expression of early genes such as discoidin I, and for basal expression of the aggregation genes, cAR1, PdsA and adenylate cyclase A (ACA; Schulkes and

A population of cells or other entities that alternate a period of excitation with a desensitized refractory period is known as an excitable medium. Such media can generate spontaneous oscillations and waves of excitation. The geometry of the waves depends on local physical or chemical conditions. In a field of starving cells, randomly pulsing cells appear first, followed by circular waves that break up to form single- or multi-armed spirals. Local high concentrations of the PdsA inhibitor, PDI that

Fig. 4.2. Production of cAMP pulses. Pulsatile cAMP signalling results from positive and negative feedback loops acting on cAMP production. Cells secrete a low basal level of cAMP, which binds to to the cAMP receptor cAR1 to activate the the heterotrimeric G protein G2, which splits into its a and bg subunits. The latter activates PI3 kinase, which phosphorylates the membrane phospholipid PIP2

to form PIP3. PIP3 brings the cytosolic regulator of adenylate cyclase (CRAC) to the membrane, where it activates the adenylate cyclase ACA. This positive feedback loop terminates because cAMP also causes desensitization of cAR1, which blocks further cAMP accumulation. Removal of extracellular cAMP by the phosphodiesterase PdsA resets the system for the next pulse

1. ACA produces a low basal level of cAMP, which is rapidly secreted. 2. Secreted cAMP binds to the cAMP receptor cAR1 and upregulates ACA activity. This involves activation of the heterotrimeric Gprotein, G2, and a series of intermediate steps. 3. The cAMP pulse reaches its peak when persistent stimulation with cAMP brings cAR1 in an inactive desensitized state, blocking further activation of ACA. 4. Hydrolysis of extracellular cAMP by the cAMP phosphodiesterase PdsA frees up cAR1 and resets the system for the next pulse (Devreotes 1994; Saran et al. 2002; Kriebel and Parent 2004).

62

Pauline Schaap

is secreted during starvation firstly promote spontaneous pulsing and secondly cause the transformation of circular into spiral waves (Palsson and Cox 1996). In the absence of PDI, spiral waves never form and aggregation territories remain small (Palsson et al. 1997). Spiral cAMP waves persist in mounds once aggregation is completed. They evolve into scroll waves in the front of the slug and standing waves in the rear, the three-dimensional equivalents of spiral and concentric waves, respectively (Dormann et al. 1998; Dormann and Weijer 2001). cAMP waves originating from the tip shape the organism during slug and fruiting body formation by coordinating the movement of its component cells. Extracellular cAMP also gains other roles in cell type-specification, and new layers of complexity in cAMP signalling become apparent after aggregation. 2. Signalling in Mounds and Slugs Once collected in mounds, overt phenotypic differences between cells become apparent. First, the proteins that are involved in oscillatory cAMP signalling, such as ACA, PdsA and cAR1, become specifically localized or enriched in prestalk and anterior-like cells. Three new cARs appear: cAR3 in all cells, and cAR2 and cAR4 in prestalk cells (Ginsburg et al. 1995). Two new adenylate cyclases appear, ACG and ACB. ACG is translationally upregulated in the posterior region of the early slug (Alvarez-Curto et al. 2007). The differentiation of prespore cells requires both extracellular cAMP acting on cAR1 (Wang et al. 1988b) and intracellular cAMP acting on PKA (Hopper et al. 1993). ACG produces cAMP for both purposes. ACB is specifically localized in prestalk cells; its activity increases at early fruiting body formation, where it is required for both spore and stalk cell maturation (Meima and Schaap 1999; Soderbom et al. 1999; Alvarez-Curto et al. 2007). The prespore cells produce polyketide based compounds that cause redifferentiation into anterior-like cells. One of these compounds is the differentiation-inducing-factor, DIF-1. DIF-1 triggers differentiation of a subpopulation of anterior-like cells that will sort downward during fruiting body formation to form the basal disk and lower cup (Saito et al. 2008).

Other polyketide-based compounds were purified that promote expression of other ALC, prestalk and prespore markers in cell suspension (Serafimidis and Kay 2005; Saito et al. 2006). Additionally, a glycoprotein, Psi, was identified that promotes prespore differentiation (Oohata et al. 1997; Kawata et al. 2004).

In addition to these diffusible molecules, cell differentiation in mounds also requires direct cell–cell interactions that are mediated by the cell adhesion protein LagC (Dynes et al. 1994). These observations suggest that prespore and prestalk differentiation are regulated by multiple levels of control. 3. Signalling during Fruiting Body Formation Incident light causes migrating slugs to point their tips upwards and initiate fruiting body formation (Fig. 4.3). Ammonia antagonizes this response and causes prolonged migration (Schindler and Sussman 1977). This is at least partly to inhibition of terminal stalk and spore differentiation by ammonia (Wang and Schaap 1989; Hopper et al. 1993). Ammonia and several other signals that regulate terminal differentiation are detected by sensor-linked histidine kinases, common sensors for external stimuli in bacteria, fungi and plants, but not in animals (Shaulsky et al. 1998; Thomason et al. 1998, 1999). The histidine kinase moiety can also act as a histidine phosphatase. Ligand binding to the sensor domain can trigger either the kinase or phosphatase activity. Active kinase phosphorylates itself on a histidine residue which starts a series of his-asp-his transfers. In Dictyostelium, the phosphoryl group is deposited via the phosphorelay intermediate RdeA on the response regulator of the intracellular cAMP phosphodiesterase RegA, thereby activating cAMP hydrolysis (Fig. 4.4). When ligand activates the histidine phosphatase, phosphorelay runs in reverse and inactivates RegA. Ammonia activates the histidine kinase DhkC and consequently RegA (Singleton et al. 1998), resulting in cAMP degradation and preventing activation of PKA. PKA activation is absolutely essential for spore and stalk cell maturation, and ammonia activation of RegA therefore effectively blocks both processes. (Harwood et al. 1992; Hopper et al. 1993; Mann et al. 1994). Slugs migrate in a thin layer of surface water that can accumulate fairly high levels of ammonia. However, when incident light causes the slug to

Evolution of Signalling and Morphogenesis in the Dictyostelids

63

Fig. 4.3. Induction of terminal differentiation. Protein degradation in starving cells yields large amounts of ammonia (NH3), which accumulates in soil moisture. Once the slugs reach the soil surface, light from above redirects the tip upwards across the air–water interface. Ammonia is now

lost by gaseous diffusion and ammonia depletion allows stalk cells to mature. Maturing prespore cells secrete a protein acbA that is processed by prestalk cells to produce the spore differentiation factor SDF2, which together with ammonia depletion triggers spore encapsulation

Fig. 4.4. Regulation of terminal differentiation of prestalk and prespore cells. Terminal spore and stalk maturation require high levels of PKA activation by cAMP. In prestalk cells cAMP is produced by the adenylate cyclase ACB and in prespore cells by the adenylate cyclase ACG. The cAMP phosphodiesterase RegA requires phosphorylation to be active. The phosphoryl group is provided by a sensor histidine kinase that can either act as a kinase or a phosphatase when a ligand binds to the sensor domain (DhkA, DhkC). The kinase autophosphorylates on a histidine

residue and, via a series of histidine to aspartate to histidine transfers, the phosphoryl group is carried via the phosphorelay intermediate RdeA over to RegA. Ammonia activates the histidine kinase activity of DhkC, thereby causing activation of RegA and preventing cAMP accumulation. The protease TagC that is expressed on prestalk cells cleaves acbA to yield SDF2, which activates the phosphatase activity of the histidine kinase DhkA. This results in dephosphorylation of RegA and consequent cAMP accumulation and PKA activation in prespore cells

lift its tip in the air, gaseous ammonia is lost from the rising structure, allowing stalk cells and spores to mature (Fig. 4.3). Prespore cells also require the presence of prestalk cells to proceed with encapsulation. Prespore cells secrete a protein acbA, which is processed by the protease TagC on prestalk cells to yield the peptide SDF2 (Fig. 4.4). SFD2 in turn binds to the sensor histidine kinase DhkA on prespore cells. This activates the phos-

phatase activity of DhkA, resulting in reverse phosphorelay and inactivation of RegA. cAMP accumulates and activates PKA which triggers spore encapsulation (Anjard et al. 1998; Wang et al. 1999; Anjard and Loomis 2005). By thus regulating the levels of intracellular cAMP, both loss of ammonia and production of SDF2 make sure that spores and stalk cells mature in the fruiting body at the right place and time.

64

Pauline Schaap

4. Signalling in the Spore Head Dictyostelid survival critically depends on spore dispersal to novel feeding grounds. Spore germination is therefore tightly regulated (Fig. 4.5). High intracellular cAMP levels and PKA activation maintain the dormant state (Van Es et al. 1996; Virdy et al. 1999). The high osmolality of the spore mass, which is caused by the presence of large amounts of ammonium phosphate blocks spore germination (Cotter 1977; Cotter et al. 1999). High osmolality has two targets: it acts on the intrinsic osmosensor of ACG to activate cAMP synthesis (Van Es et al. 1996; Saran and Schaap 2004) and it inhibits cAMP degradation by RegA by triggering reverse phosphorelay through the histidine kinase DokA (Schuster et al. 1996; Oehme and Schuster 2001). In conclusion, the transduction mechanisms of known signals that control the final stages of development all converge at regulating intracellular cAMP levels and the activation of PKA. cAMP is the most deeply conserved signalling intermediate in both pro- and eukaryotes, with a broad repertoire of functions in environmental sensing, hormone action, neurotransmission and developmental control. While intracellular use of cAMP is common to most organisms, extracellular cAMP signalling is thus far unique to social amoebas. This suggests that the extracellular use of cAMP is derived from its intracellular use. PKA is not essential for growth in social amoebas, but is thereafter required for all stages of starvation-

Fig. 4.5. Osmoregulation of spore germination. High ambient osmolarity prevents spores from germinating in the spore head. High osmolarity directly stimulates cAMP synthesis by ACG and indirectly inhibits cAMP hydrolysis by RegA by activating the phosphatase activity of the osmoregulated histidine kinase DokA. High cAMP levels in spores act on PKA to inhibit germination

induced development. It is therefore not unlikely that cAMP signalling in social amoebas has evolved from a stress response in their unicellular ancestors.

III. Evolution of Morphogenesis and Developmental Signalling A. Evolution of Morphology The evolution of novel forms in multicellular organisms results from alterations in the developmental programmes that generate these forms. For most groups of multicellular organisms, more is known of developmental signalling in the modern complex members of the group than in the often extinct unicellular ancestors. The evolutionary origins of these processes can therefore only be retraced by a top-down approach. To assess what came first and what was added later it is essential to know the phylogenetic relationships of the group of organisms under study. A family tree of most known social amoebas species, based on conserved small subunit ribosomal RNA (SSU rRNA) and b-tubulin protein sequences was recently constructed (Schaap et al. 2006a). This tree shows the subdivision of all known social amoeba species into four major groups (Fig. 4.6). The four groups do not coincide with the earlier subdivision into genera that was based on fruiting body morphology (Raper 1984). The genus Polysphondylium appears at two different locations: in group 2, and between groups 3 and 4. Group 2 also contains all members of the genus Acytostelium, as well as some Dictyostelium species.

By plotting morphological characters of individual species to the family tree, the order in which these characters evolved can be assessed. Social amoebas species differ in the shape and size of the fruiting bodies, which can be solitary or clustered, unbranched or carrying branches with specific topologies. Cellular structures such as basal disks and triangular supporters may support the stalk, while upper and lower cups may support the spore head. A migrating slug stage can be present and often a stalk is laid down during migration. Alternative life cycles, such as the microcyst or macrocyst can be present. Spores vary in size and shape and may carry granules at

Group 3

D. deminutivum MexM19A

]

]

Dermamoeba algensis

D. multi-stipes UK26b

1

1.00

Thecamoeba similis

D. laterosorum AE4 P. violaceum P6

with most species in group 2 and one species P. violaceum, clustered with D. laterosorum at the base of group 4. Group 2 also contains the genus Acytostelium and two members of the genus Dictyostelium (reproduced with permission of Science)

other amoebas

D. polycarpum VE1b D. polycarpum OhioWILDS P. filamentosum SU-1 P. luridum LR-2 P. pallidum TNS-C-98 P. equisetoides B75B P. nandutensis YA1 P. colligatum OH538 P. tikaliensis OH595 P. anisocaule NZ47B P. pseudocandidum TNS-C-91 P. tenuissimum TNS-C-97 Group 2 D. gloeosporum TCK52 P. pallidum PN500 P. asymmetricum OH567 D. oculare DB4B A. ellipticum AE2 A. anastomosans PP1 A. longisorophorum DB10A A. leptosomum FG12 A. digitatum OH517 A. serpentarium SAB3A A. subglobosum Lb1 D. antarcticum NZ43B D. fasciculatum SmokOW9A D. delicatum TNS-C-226 D. fasciculatum SH3 D. aureo-stipes var. helveticum GE1 D. aureo-stipes YA6 D. aureo-stipes JKS150 Group D. granulophorum CHII-4 D. medusoides OH592 D. mexicanum MexTF4B1 D. bifurcatum UK5 D. stellatum SAB7B D. microsporum TNS-C-38 D. parvisporum OS126 D. exiguum TNS-C-199

D. menorah M1 D. caveatum WS695 D. polycephalum MY1-1

D. lacteum

]

Group 4

D. gracile TNS-C-183 D. lavandulum B15 D. vinaceo-fuscum CC4 D. rhizopodium AusKY-4 D. coeruleo-stipes CRLC53B

D. tenue PJ6 D. tenue PR4

D. laterosorum AE4 P. violaceum P6 D. australe NZ80B D. monochasioides HAG653 D. tenue Pan52 D. potamoides FP1A D. minutum 71-2

Fig. 4.6. Family tree of the social amoebas. The phylogenetic tree of the social amoebas shows subdivision of all species into four major groups (Schaap et al. 2006a). The tree is based on small subunit ribosomal RNA sequences and is rooted on sequences of solitary amoebazoa. The genus Polysphondylium is polyphyletic

0.1

0.97-1.0 0.90-0.96 0.75-0.89 0.50-0.74 <0.50

Bayesian Inference posterior probabilities

D. brefeldianum TNS-C-115 D. mucoroides S28b D. capitatum 91HO-50 D. pseudobrefeldianum 91HO-8 D. aureocephalum TNS-C-180 D. aureum SL1 D. septentrionalis IY49 D. septentrionalis AK2 D. implicatum 93HO-1 D. medium TNS-C-205 D. mucoroides var. stoloniferum FOII-1 D. crassicaule 93HO-33 D. mucoroides TNS-C-114 D. sphaerocephalum GR11 D. rosarium M45 D. clavatum TNS-C-220 D. clavatum TNS-C-189 D. longosporum TNS-C-109 D. purpureum C143 D. purpureum WS321 D. macrocephalum B33 D. discoideum 91HO-9 D. discoideum AX4 D. discoideum AX2 D. discoideum X00134 D. discoideum NC4 D. discoideum AC4 D. citrinum OH494 D. discoideum V34 D. dimigraformum AR5b D. intermedium PJ11 D. firmibasis TNS-C-14 0.01 D. brunneum WS700 D. giganteum WS589 D. robustum TNS-C-219

Evolution of Signalling and Morphogenesis in the Dictyostelids 65

66

Pauline Schaap

their poles. The chemo-attractant used for aggregation may be cAMP, folic acid, glorin or pterin but is unknown for most species. Twenty consistently described characters were plotted to the family tree (Schaap et al. 2006a). The characters that showed clear evolutionary trends are summarized in Fig. 4.7. Sexual macrocysts are found in all groups, but microcysts appear to be lost from the most derived group 4. Spores are smallest in group 1 and largest in group 4, where they have also lost the polar granules. Clusters of fruiting bodies usually arise from a single aggregate in groups 1–3, and these fruiting bodies are also often branched. However, aggregates usually yield solitary unbranched fruiting bodies in group 4 species. As a consequence group 4 fruiting bodies are significantly larger than those of other groups, and they are buttressed by basal disks or triangular supporters. Outwith group 4 there is a single

species with a basal disc in group 1 and a cluster of species in group 3 with a crampon-shaped support. Remarkably, for all tested species in group 4, the chemoattractant is cAMP, while the tested species in the other groups all use other chemoattractants. There is apparently a correlation between the use of cAMP during aggregation, large fruiting body size with additional celltype specialization, large spore size and loss of microcysts. This suggests that elaboration of cAMP signalling in group 4 species may have contributed to these phenotypic changes.

Fig. 4.7. Major trends in the evolution of morphology. A total of 20 described morphological characters were mapped to the phylogenetic tree. Many of those are more or less randomly scattered over the four taxon groups. An example of such a trait is sexual macrocyst formation, which occurs in multiple species in each group. However encystation of solitary amoebas to form microcysts is lost in the most derived group 4. Species in groups 1–3 generally form many tips on aggregates and sorogens, which give rise to clusters of small branched fruiting bodies.

Group 4 species usually form only one or a few tips per aggregate and fruiting bodies are consequently solitary, unbranched and large. Virtually all group 4 species form cellular supporters or basal disks to hold up the stalk. A cluster of four species in group 3 forms unique cramponlike supports. Spore size is smallest in group 1 and largest in group 4, where spores are also devoid of the polar granules that characterize spores in groups 1–3. None of the species in groups 1–3 use cAMP as chemoattractant, but all investigated species in group 4 do

B. Evolution of Developmental Signalling The model system D. discoideum has been almost exclusively used to study developmental processes and information on developmental signals in other social amoebas is therefore limited. However, the

Evolution of Signalling and Morphogenesis in the Dictyostelids

genomes of two basal species D. fasciculatum (group 1), Polyshondylium pallidum (group 2) are currently being sequenced. This should immediately make evident whether protein-derived signals such as CMF, Psi, SDF2 and LagC are conserved, as well as the proteins that are involved in the synthesis, detection, processing and degradation of these and other signals. Equally interesting is information on conservation of specific members of the polyketide synthase gene family, since this may lead us to the signal for prestalk differentiation that has become increasingly elusive (Thompson and Kay 2000b; Saito et al. 2008). The basal disc inducer DIF-1 is present in the group 4 species, D. mucoroides, which also has the DIF-degrading enzyme, DIF-dechlorinase. D. minutum, in group 3, and Polyshondylium violaceum, between groups 3 and 4, synthesize other chlorinated factors that induce stalk cell differentiation in D. discoideum. However, they have neither DIF-dechlorinase nor basal disc cells, suggesting that DIF-1 induction of basal disc cells is a recent innovation (Kay et al. 1992; Van Es et al. 1994). Ammonia blocks aggregation and causes cells to form microcysts in the group 2 species P. pallidum (Lonski 1976; Choi and O’Day 1982). Effects on later development have not been investigated, but are very likely considering the fact that all species produce this catabolite. cAMP signalling has been studied more extensively in other species. Similar to D. discoideum, P. pallidum also requires PKA activation to proceed through development (Funamoto et al. 2003). The extracellular roles of cAMP are partly conserved. The cAR1 gene is present in representative species of all four groups (Alvarez-Curto et al. 2005), but the gene duplications that gave rise to cAR2, cAR3 and cAR4 only occurred in group 4 (Kawabe et al. 2009). The basal cAR1s are functionally identical to D. discoideum cAR1, but are only expressed after aggregation. D. discoideum and other group 4 species express a second cAR1 mRNA before the onset of aggregation (AlvarezCurto et al. 2005). Loss of cAR1 function in group 4 species blocks aggregation and further development. However, in groups 1–3 aggregation is normal, but the subsequent formation of slugs and fruiting bodies is disrupted. This implies that coordination of fruiting body formation is the ancestral role of extracellular cAMP signalling, while its role in aggregation is evolutionary derived.

67

In addition to using cAMP for aggregation, group 4 species stand out by having large solitary unbranched fruiting structures, as opposed to the clustered and branched small fruiting bodies that are found in groups 1–3. Are cAMP signalling and size related? The segmentation of aggregates into clusters of fruiting bodies and the formation of side branches all represent the formation of multiple body axes that are initiated by newly emerging tips (Raper 1940). D. discoideum tips are pacemakers for cAMP waves that suppress the emergence of competing pacemakers. This occurs in two ways: 1. More excitable cells that oscillate at high frequency will naturally entrain cells that oscillate at lower frequency (Lee et al. 2002). 2. A long-range inhibitor is produced by tips that reduces the excitability of surrounding cells (Durston 1976; Kopachik 1982). The cAMP hydrolysis product adenosine was proposed to fulfil this role (Newell and Ross 1982; Schaap and Wang 1986). Tip dominance is therefore intrinsic to oscillatory cAMP signalling and group 4 species have larger fruiting bodies because their cAMP oscillators are better at suppressing competitors. Two aspects of cAMP signalling are specific to group 4: (i) oscillatory cAMP signalling occurs much earlier in development than in groups 1–3 and (ii) the cAMP receptor gene was duplicated three times, and both expression and affinity of the daughter cARs were altered. It is conceivable that either of these novelties may have “improved” cAMP signalling to allow it to control larger numbers of cells and make it generally more robust. More recent work shows that the adenylate cyclase ACG and its role in osmoregulation of spore germination are conserved in all four taxon groups. Moreover, in species that form microcysts, ACG and PKA critically regulate encystation. Microcysts usually form when cells are starved, but high osmolality triggers encystation independently. For soil amoebas high osmolality may signal approaching drought, when the concentration of soil minerals increases. High osmolality triggers cAMP synthesis by ACG, which activates PKA and initiates the encystation process. Since encystation represents an ancestral mode of stress survival, this implies that the roles of ACG and PKA in prespore and spore differenti-

68

Pauline Schaap

ation during multicellular development were evolutionary derived from homologous roles in encystation (Ritchie et al. 2008).

IV. Conclusion The social amoebas with their unicellular/multicellular combination life style are an outstanding model to understand the evolution of multicellular development. Their recently completed molecular phylogeny provides a framework for evolutionary studies, but the ongoing comparative genome sequencing projects will provide information on the core components of developmental pathways and the patterns of gene duplication and loss in each of the major groups. This will show how ancestral signalling mechanisms were elaborated and how developmental robustness increased by bringing crucial steps in the life cycle under multiple levels of control. Whole genome comparison will show how the ratio of coding to non-coding DNA changed during evolution. This is important, since novel developmental roles for existing genes often arise through elaboration of promoter structure. Comparison of coding sequences will reveal changes in functional domain organization and conserved amino acids in proteins that allow formulation of hypotheses about changes in protein function during evolution. One of the strengths of the Dictyostelium system is that such hypotheses can be tested by gene replacement. In addition to the model D. discoideum, genetic tractability has also been developed for the group 2 species P. pallidum and this can hopefully be achieved for at least one species out of each major taxon group. Ecology, or rather adaptation to specific habitats is the ultimate cause for the transition to a multicellular life style and the evolution of multicellular complexity. We can only hope to elucidate how multicellularity evolved if we know what has driven it the first place. A combined effort of field biologists and developmental biologists is currently underway to map the factors that characterize the habitats of species to the phylogeny, such as latitude, altitude, (micro) climate, vegetation and soil type. In addition to the existing map of gross morphological traits, a map of deeper traits, such as developmental robustness, pattern formation, proportion regulation,

signalling dynamics is also being prepared. A combination of the three maps should allow us to discover relationships between ecological features and the evolution of phenotype. Only by incorporating this information with the gene and genome level changes that caused phenotypic change can we fully understand why and how multicellular organisms evolved.

References Alvarez-Curto E, Rozen DE, Ritchie AV, Fouquet C, Baldauf SL, Schaap P (2005) Evolutionary origin of cAMP-based chemoattraction in the social amoebae. Proc Natl Acad Sci USA 102:6385–6390 Alvarez-Curto E, Saran S, Meima M, Zobel J, Scott C, Schaap P (2007) cAMP production by adenylyl cyclase G induces prespore differentiation in Dictyostelium slugs. Development 134:959–966 Amagai A (1984) Induction by ethylene of macrocyst formation in the cellular slime mould Dictyostelium mucuroides. J Gen Microbiol 130:2961–2965 Anderson JS, Fennell DI, Raper KB (1968) Dictyostelium deminutivum, a new cellular slime mold. Mycologia 60:49–64 Anjard C, Loomis WF (2005) Peptide signaling during terminal differentiation of Dictyostelium. Proc Natl Acad Sci USA 102:7607–7611 Anjard C, Zeng CJ, Loomis WF, Nellen W (1998) Signal transduction pathways leading to spore differentiation in Dictyostelium discoideum. Dev Biol 193:146–155 Baldauf SL, Roger AJ, Wenk-Siefert I, Doolittle WF (2000) A kingdom-level phylogeny of eukaryotes based on combined protein data. Science 290:972–977 Bonner JT, Clarke Jr WW, Neely Jr CL, Slifkin MK (1950) The orientation to light and the extremely sensitive orientation to temperature gradients in the slime mold Dictyostelium discoideum. J Cell Comp Physiol 36:149–158 Bonner JT, Lamont DS (2005) Behavior of cellular slime molds in the soil. Mycologia 97:178–184 Castillo DI, Switz GT, Foster KR, Queller DC, Strassmann JE (2005) A cost to chimerism in Dictyostelium discoideum on natural substrates. Evol Ecol Res 7:263–271 Chang MT, Raper KB, Poff KL (1983) The effect of light on morphogenesis of Dictyostelium mucuroides. Exp Cell Res 143:335–341 Choi AH, O’Day DH (1982) Ammonia and the induction of microcyst differentiation in wild type and mutant strains of the cellular slime mold Polysphondylium pallidum. Dev Biol 92:356–364 Clarke M, Gomer RH (1995) PSF and CMF, autocrine factors that regulate gene expression during growth and early development of Dictyostelium. Experientia 51:1124–1134 Cotter DA (1977) The effects of osmotic pressure changes on the germination of Dictyostelium discoideum spores. Can J Microbiol 23:1170–1177

Evolution of Signalling and Morphogenesis in the Dictyostelids Cotter DA, Dunbar AJ, Buconjic SD, Wheldrake JF (1999) Ammonium phosphate in sori of Dictyostelium discoideum promotes spore dormancy through stimulation of the osmosensor ACG. Microbiology 145:1891–1901 Deery WJ, Gao T, Ammann R, Gomer RH (2002) A single cell density-sensing factor stimulates distinct signal transduction pathways through two different receptors. J Biol Chem 277:31972–31979 Devreotes PN (1994) G protein-linked signaling pathways control the developmental program of Dictyostelium. Neuron 12:235–241 Dormann D, Vasiev B, Weijer CJ (1998) Propagating waves control Dictyostelium discoideum morphogenesis. Biophys Chem 72:21–35 Dormann D, Weijer CJ (2001) Propagating chemoattractant waves coordinate periodic cell movement in Dictyostelium slugs. Development 128 4535–4543 Durston AJ (1976) Tip formation is regulated by an inhibitory gradient in the Dictyostelium discoideum slug. Nature 263:126–129 Dynes JL, Clark AM, Shaulsky G, Kuspa A, Loomis WF, Firtel RA (1994) LagC is required for cell–cell interactions that are essential for cell-type differentiation in Dictyostelium. Genes Dev 8:948–958 Eichinger D (2001) Encystation in parasitic protozoa. Curr Opin Microbiol 4:421–426 Feit IN, Bonner JT, Suthers HB (1990) Regulation of the anterior-like cell state by ammonia in Dictyostelium discoideum. Dev Gen 11:442–446 Feit IN, Medynski EJ, Rothrock MJ (2001) Ammonia differentially suppresses the cAMP chemotaxis of anterior-like cells and prestalk cells in Dictyostelium discoideum. J Biosci 26:157–166. Fosnaugh KL, Loomis WF (1991) Coordinate regulation of the spore coat genes in Dictyostelium discoideum. Dev Gen 12:123–132 Funamoto S, Anjard C, Nellen W, Ochiai H (2003) cAMPdependent protein kinase regulates Polysphondylium pallidum development. Differentiation 71:51–61. Gerisch G, Fromm H, Huesgen A, Wick U (1975) Control of cell-contact sites by cyclic AMP pulses in differentiating Dictyostelium cells. Nature 255:547–549 Ginsburg GT, Gollop R, Yu Y, Louis JM, Saxe CL, Kimmel AR (1995) The regulation of Dictyostelium development by transmembrane signalling. J Euk Microbiol 42:200–205 Gutierrez JC, Martin-Gonzalez A, Matsusaka T (1990) Towards a generalized model of encystment (cryptobiosis) in ciliates: a review and a hypothesis. Biosystems 24:17–24 Hartley AD, Ward MP, Garrett S (1994) The Yak1 protein kinase of Saccharomyces cerevisiae moderates thermotolerance and inhibits growth by an Sch9 protein kinase-independent mechanism. Genetics 136: 465–474 Harwood AJ, Hopper NA, Simon M-N, Driscoll DM, Veron M, Williams JG (1992) Culmination in Dictyostelium is regulated by the cAMP-dependent protein kinase. Cell 69:615–624 Hopper NA, Harwood AJ, Bouzid S, Ve´ron M, Williams JG (1993) Activation of the prespore and spore cell path-

69

way of Dictyostelium differentiation by cAMP-dependent protein kinase and evidence for its upstream regulation by ammonia. EMBO J. 12:2459–2466 Inouye K (1984) Measurement of the motive force of the migrating slug of Dictyostelium discoideum by a centrifuge method. Protoplasma 121:171–177 Kawabe Y, Morio T, James JL, Prescott AR, Tanaka Y, Schaap P (2009) Activated cAMP receptors turn encystation into sporulation. Proc Natl Acad Sci USA 106: 7089–7094 Kawata T, Nakagawa M, Shimada N, Fujii S, Oohata AA (2004) A gene encoding prespore-cell-inducing factor in Dictyostelium discoideum. Dev Growth Differ 46: 383–392 Kay RR, Taylor GW, Jermyn KA, Traynor D (1992) Chlorine-containing compounds produced during Dictyostelium development. Detection by labelling with 36Cl. Biochem J 281:155–161 King N (2004) The unicellular ancestry of animal development. Dev Cell 7:313–325 Kopachik WJ (1982) Size regulation in Dictyostelium. J. Embryol.Exp.Morph. 68:23–35 Kriebel PW, Parent CA (2004) Adenylyl cyclase expression and regulation during the differentiation of Dictyostelium discoideum. IUBMB Life 56:541–546 Lauwaet T, Davids BJ, Reiner DS, Gillin FD (2007) Encystation of Giardia lamblia: a model for other parasites. Curr Opin Microbiol 10:554–559 Leach CK, Ashworth JM, Garrod DR (1973) Cell sorting out during the differentiation of mixtures of metabolically distinct populations of Dictyostelium discoideum. J Embryol Exp Morphol 29:647–661 Lee KJ, Goldstein RE, Cox EC (2002) cAMP waves in Dictyostelium territories. Nonlinearity 15:C1–C5 Lonski J (1976) The effect of ammonia on fruiting body size and microcyst formation in the cellular slime molds. Dev Biol 51:158–165 Mann SKO, Richardson DL, Lee S, Kimmel AR, Firtel RA (1994) Expression of cAMP-dependent protein kinase in prespore cells is sufficient to induce spore cell differentiation in Dictyostelium. Proc Natl Acad Sci USA 91:10561–10565 Matsukuma S, Durston AJ (1979) Chemotactic cell sorting in Dictyostelium discoideum. J Embryol Exp Morph 50:243–251 McRobbie SJ, Jermyn KA, Duffy K, Blight K, Williams JG (1988) Two DIF-inducible, prestalk-specific mRNAs of Dictyostelium encode extracellular matrix protein of the slug. Development 104:275–284 Meima ME, Schaap P (1999) Fingerprinting of adenylyl cyclase activities during Dictyostelium development indicates a dominant role for adenylyl cyclase B in terminal differentiation. Dev Biol 212:182–190 Mercer SE, Friedman E (2006) Mirk/Dyrk1B: a multifunctional dual-specificity kinase involved in growth arrest, differentiation, and cell survival. Cell Biochem Biophys 45:303–315 Mutzel R, Lacombe M-L, Simon M-N, De Gunzburg J, Veron M (1987) Cloning and cDNA sequence of the regulatory subunit of cAMP-dependent protein kinase from Dictyostelium discoideum. Proc Natl Acad Sci USA 84:6–10

70

Pauline Schaap

Newell PC, Ross FM (1982) Inhibition by adenosine of aggregation centre initiation and cyclic AMP binding in Dictyostelium. J Gen Microbiol 128:2715–2724 Oehme F, Schuster SC (2001) Osmotic stress-dependent serine phosphorylation of the histidine kinase homologue DokA. BMC Biochem. 2:2 Oohata AA, Nakagawa M, Tasaka M, Fujii S (1997) A novel prespore-cell-inducing factor in Dictyostelium discoideum induces cell division of prespore cells. Development 124:2781–2787 Otto GP, Wu MY, Kazgan N, Anderson OR, Kessin RH (2003) Macroautophagy is required for multicellular development of the social amoeba Dictyostelium discoideum. J Biol Chem 278:17636–17645 Palsson E, Cox EC (1996) Origin and evolution of circular waves and spirals in Dictyostelium discoideum territories. Proc Natl Acad Sci USA 93:1151–1155 Palsson E, Lee KJ, Goldstein RE, Franke J, Kessin RH, Cox EC (1997) Selection for spiral waves in the social amoebae Dictyostelium. Proc Nat Acad Sci USA 94: 13719–13723 Peracino B, Borleis J, Jin T, Westphal M, Schwartz J-M, Wu L, Bracco E, Gerisch G, Devreotes P, Bozzaro S (1998) G Protein beta subunit-null mutants are impaired in phagocytosis and chemotaxis due to inappropriate regulation of the actin cytoskeleton J Cell Biol 141: 1529–1537 Raper KB (1940) Pseudoplasmodium formation and organization in Dictyostelium discoideum. J. Elisha Mitchell Sci Soc 56:241–282 Raper KB (1984) The Dictyostelids. Princeton University Press, Princeton, New Jersey Raper KB, Fennell DI (1952) Stalk formation in Dictyostelium. Bull Torrey Club 79:25–51 Rathi A, Kayman SC, Clarke M (1991) Induction of gene expression in Dictyostelium by prestarvation factor, a factor secreted by growing cells. Dev Gen 12:82–87 Ritchie, AV, van Es S, Fouquet C, Schaap P (2008) From drought sensing to developmental control: evolution of cyclic AMP signaling in social amoebas. Mol Biol Evol 25: 2109–2118 Saito T, Kato A, Kay R (2008) DIF-1 induces the basal disc of the Dictyostelium fruiting body. Dev Biol (in press) Saito T, Taylor GW, Yang JC, Neuhaus D, Stetsenko D, Kato A, Kay RR (2006) Identification of new differentiation inducing factors from Dictyostelium discoideum. Biochim Biophys Acta 1760:754–761 Saran S, Meima ME, Alvarez-Curto E, Weening KE, Rozen DE, Schaap P (2002) cAMP signaling in Dictyostelium – complexity of cAMP synthesis, degradation and detection. J Mus Res Cell Mot 23:793–802 Saran S, Schaap P (2004) Adenylyl cyclase G is activated by an intramolecular osmosensor. Mol. Biol. Cell 15: 1479–1486 Schaap P (1983) Quantitative analysis of the spatial distribution of ultrastructural differentiation markers during development of Dictyostelium discoideum. Roux’s Arch Dev Biol 192:86–94 Schaap P, Spek W (1984) Cyclic-AMP binding to the cell surface during development of Dictyostelium discoideum. Differentiation 27:83–87

Schaap P, Van Lookeren Campagne MM, Van Driel R, Spek W, Van Haastert PJ, Pinas J (1986) Postaggregative differentiation induction by cyclic AMP in Dictyostelium: intracellular transduction pathway and requirement for additional stimuli. Dev Biol 118:52–63 Schaap P, Wang M (1986) Interactions between adenosine and oscillatory cAMP signaling regulate size and pattern in Dictyostelium. Cell 45:137–144 Schaap P, Winckler T, Nelson M, Alvarez-Curto E, Elgie B, Hagiwara H, Cavender J, Milano-Curto A, Rozen DE, Dingermann T, Mutzel R, Baldauf SL (2006) Molecular phylogeny and evolution of morphology in the social amoebas. Science 314:661–663 Schindler J, Sussman M (1977) Ammonia determines the choice of morphogenetic pathways in Dictyostelium discoideum. J Mol Biol 116:161–169 Schulkes C, Schaap P (1995) cAMP-dependent protein kinase activity is essential for preaggegative gene expression in Dictyostelium. FEBS Lett 368:381–384 Schuster SC, Noegel AA, Oehme F, Gerisch G, Simon MI (1996) The hybrid histidine kinase DokA is part of the osmotic response system of Dictyostelium. EMBO J 15:3880–3889 Serafimidis I, Kay RR (2005) New prestalk and prespore inducing signals in Dictyostelium. Dev Biol 282: 432–441 Shaulsky G, Fuller D, Loomis WF (1998) A cAMPphosphodiesterase controls PKA-dependent differentiation. Development 125:691–699 Singleton CK, Zinda MJ, Mykytka B, Yang P (1998) The histidine kinase dhkC regulates the choice between migrating slugs and terminal differentiation in Dictyostelium discoideum. Dev Biol 203:345–357 Smith E, Williams KL (1981) The age-dependent loss of cells from the rear of a Dictyostelium discoideum slug is not tip controlled. J Embryol Exp Morph 61:61–67 Soderbom F, Anjard C, Iranfar N, Fuller D, Loomis WF (1999) An adenylyl cyclase that functions during late development of Dictyostelium. Development 126: 5463–5471 Souza GM, daSilva AM, Kuspa A (1999) Starvation promotes Dictyostelium development by relieving PufA inhibition of PKA translation through the YakA kinase pathway. Development 126:3263–3274 Souza GM, Lu SJ, Kuspa A (1998) YakA, a protein kinase required for the transition from growth to development in Dictyostelium. Development 125:2291–2302 Steinert M, Heuner K (2005) Dictyostelium as host model for pathogenesis. Cell Microbiol 7:307–314 Sternfeld J, David CN (1981) Cell sorting during pattern formation in Dictyostelium. Differentiation 20:10–21 Sternfeld J, David CN (1982) Fate and regulation of anterior-like cells in Dictyostelium slugs. Dev Biol 93:111–118 Strassmann JE, Zhu Y, Queller DC (2000) Altruism and social cheating in the social amoeba Dictyostelium discoideum. Nature 408:965–967 Swanson AR, Vadell EM, Cavender JC (1999) Global distribution of forest soil dictyostelids. J Biogeography 26:133–148 Thomason PA, Traynor D, Cavet G, Chang W-T, Harwood AJ, Kay RR (1998) An intersection of the cAMP/PKA

Evolution of Signalling and Morphogenesis in the Dictyostelids and two-component signal transduction systems in Dictyostelium. EMBO J. 17:2838–2845 Thomason PA, Traynor D, Stock JB, Kay RR (1999) The RdeA-RegA system, a eukaryotic phospho-relay controlling cAMP breakdown. J Biol Chem 274:27379–27384 Thompson CR, Kay RR (2000a) Cell-fate choice in Dictyostelium: intrinsic biases modulate sensitivity to DIF signaling. Dev Biol 227:56–64 Thompson CR, Kay RR (2000b) The role of DIF-1 signaling in Dictyostelium development. Mol Cell 6:1509–1514 Toama MA, Raper KB (1967) Microcysts of the cellular slime mold Polysphondylium pallidum. I. Factors influencing microcyst formation. J Bacteriol 94:1143–1149 Vadell EM, Cavender JC (2007) Dictyostelids living in the soils of the Atlantic Forest, Iguazu region, Misiones, Argentina: description of new species. Mycologia 99:112–124 Van Es S, Hodgkinson S, Schaap P, Kay RR (1994) Metabolic pathways for differentiation-inducing factor-1 and their regulation are conserved between closely related Dictyostelium species, but not between distant members of the family. Differentiation 58:95–100 Van Es S, Virdy KJ, Pitt GS, Meima M, Sands TW, Devreotes PN, Cotter DA, Schaap P (1996) Adenylyl cyclase G, an osmosensor controlling germination of Dictyostelium spores. J Biol Chem 271:23623–23625 Verkerke-van Wijk I, Fukuzawa M, Devreotes PN, Schaap P (2001) Adenylyl cyclase A expression is tip-specific in Dictyostelium slugs and directs StatA nuclear translocation and CudA gene expression. Dev Biol 234: 151–160 Virdy KJ, Sands TW, Kopko SH, van Es S, Meima M, Schaap P, Cotter DA (1999) High cAMP in spores

71

of Dictyostelium discoideum: association with spore dormancy and inhibition of germination. Microbiology 145:1883–1890 Wang M, Aerts RJ, Spek W, Schaap P (1988a) Cell cycle phase in Dictyostelium discoideum is correlated with the expression of cyclic AMP production, detection, and degradation. Dev Biol 125:410–416 Wang M, Schaap P (1989) Ammonia depletion and DIF trigger stalk cell differentiation in intact Dictyostelium discoideum slugs. Development 105:569–574 Wang M, Van Driel R, Schaap P (1988b) Cyclic AMPphosphodiesterase induces dedifferentiation of prespore cells in Dictyostelium discoideum slugs: evidence that cyclic AMP is the morphogenetic signal for prespore differentiation. Development 103:611–618 Wang N, Soderbom F, Anjard C, Shaulsky G, Loomis WF (1999) SDF-2 induction of terminal differentiation in Dictyostelium discoideum is mediated by the membrane-spanning sensor kinase DhkA. Mol Cell Biol 19:4750–4756 Weijer CJ, Duschl G, David CN (1984) Dependence of celltype proportioning and sorting on cell cycle phase in Dictyostelium discoideum. J.Cell Sci. 70:133–145 West CM (2003) Comparative analysis of spore coat formation, structure, and function in Dictyostelium. Int Rev Cytol 222:237–293 Wilkins MR, Williams KL (1995) The extracellular matrix of the Dictyostelium discoideum slug. Experientia 51: 1189–1196 Yamamoto A, Takeuchi I (1983) Vital staining of autophagic vacuoles in differentiating cells of Dictyostelium discoideum. Differentiation 24:83–87

.

5

Function and Evolution of Pheromones and Pheromone Receptors in Filamentous Ascomycetes

STEFANIE PO¨GGELER1

CONTENTS I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Pheromones and Pheromone Receptors in S. cerevisiae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Pheromones of S. cerevisiae . . . . . . . . . . . . . . . . . . . B. Pheromone Receptors of S. cerevisiae . . . . . . . . III. Structure and Evolution of Pheromones and Pheromone Receptors in Filamentous Ascomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Pheromones of Filamentous Ascomycetes. . . . 1. Characteristics of MFa-Like Peptide Pheromones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Characteristics of MFa-Like Lipopeptide Pheromones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. H-Type Pheromone Precursors – a Novel Class of Peptide Pheromone Precursors in Hypocreales. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Pheromone Receptors of Filamentous Ascomycetes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Function of Pheromone/Receptor Systems in Filamentous Ascomycetes. . . . . . . . . . . . . . . . . . . . . . A. Functional Analysis of Pheromones and Pheromone Receptors in Heterothallic Filamentous Ascomycetes . . . . . . . . . . . . . . . . . . . . . B. Functional Analysis of Pheromones and Pheromone Receptors in Homothallic Filamentous Ascomycetes . . . . . . . . . . . . . . . . . . . . . V. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

73 75 75 76 77 77 79 82 84 85 86 89 90 92 92

I. Introduction Sexual reproduction is intrinsically connected to cell-type specification and the organization of different sexes. Genetic sex determination requires the establishment of genes that are encoded only in a given cell type. These may be localized on sex chromosomes, such as the X and Y chromosomes in mammals (Jime´nez and Burgos 1998; Koopman 1999). 1

Genetics of Eukaryotic Microorganisms, Institute of Microbiology and Genetics, August University Go¨ttingen, Grisebachstrasse 8, 37077 Go¨ttingen, Germany; e-mail: [email protected]

In fungi, sexual dimorphism is nearly nonexistent. More than 100 years ago, Blakeslee discovered that, in the common bread mould Rhizopus stolonifer, sexual reproduction occurs between morphologically indistinguishable partners; and he introduced the term mating type (MAT) to differentiate sexually compatible single spore strains of the same species (Blakelee 1904). Blakeslee also introduced the terms heterothallism to describe the breeding mechanism of selfsterile fungi in which the sexual cycle can be completed only by the union of two physiological different haploid nuclei and homothallism for the self-fertile fungi in which a complete sexual cycle can begin with a single haploid nucleus (Blakelee 1904; Korf 1952). The mucoraceous fungus R. stoloinfer has not, however, been amenable to genetic analysis. Therefore, the first molecular characterization of mating-type loci was not surprisingly accomplished using the model fungus Saccharomyces cerevisiae. The baker´s yeast S. cerevisiae is classified into the phylum Ascomycota, or sac fungi, a monophyletic group that accounts for approximately 64% of all described fungi (Stajich et al. 2009). Members of this group are characterized by the formation of ascospores in a sac-like structure called the ascus and are phylogenetically grouped into three subphyla: (i) the Saccaromycotina, which harbors S. cerevisiae as well as the parasitic yeast Candida albicans, (ii) the subphylum Taphrinomycotina, with species such as the fission yeast Schizosaccharomyces pombe and the pathogen Pneumocystis carinii and (iii) the Pezizomycotina, containing species that protect their asci with multicellular fruiting bodies such as the model fungus Neurospora crassa (Stajich et al. 2009; see Chapter 8 in this volume). Moreover, two morphologically distinct groups can be distinguished among the Ascomycota, the unicellular yeasts and the filamentous fungi. Pezizomycotina are generally filamentous. Both filamentous and yeastlike growth are found in Taphrinomycotina, and hyphal growth was lost in the Saccaromycotina (Stajich et al. 2009). Evolution of Fungi and Fungal-Like Organisms, The Mycota XIV S. Po¨ggeler and J. Wo¨stemeyer (Eds.) © Springer-Verlag Berlin Heidelberg 2011

74

Stefanie Po¨ggeler

The budding yeast S. cerevisiae exists as one of two haploid cell types, MATa or MATa. The MAT alleles encode gene regulatory proteins, which in combination with other regulatory proteins are responsible for a distinct pattern of expression in the three yeast cell types: haploid MATa and MATa cells, and diploid MATa/a cells (Herskowitz 1989). Each mating type locus carries two genes: the MATa locus has a1 and a2, and the MATa locus has a1 and a2 (Fig. 5.1). Since the DNA sequences at the mating-type locus are completely different in MATa and MATa cells, they have been termed idiomorphs instead of alleles to denote that the sequences at the same locus in different strains exhibit no sequence similarity (Glass et al. 1988; Metzenberg and Glass 1990). Sexual reproduction is driven by the secretion of mating-type specific pheromones activating a signal transduction pathway that regulates the cell fusion between partners of the opposite mating type (Herskowitz 1989). Pheromones are diffusible substances released by an individual member of a species that trigger a biological response by another individual of the same species (Karlson and Luscher 1959).

The pheromones are sensed by G protein-coupled cell surface receptors that initiate the mating response (Bardwell 2005; Dohlman and Slessareva 2006; Kurjan 1993; Wang and Dohlman 2004). The MATa gene a1 encodes a specific class of fungal transcription factors, namely an a-domain transcription factor that is a positive regulator of MATa-specific gene of transcription. The regulated genes are required for cell–cell recognition and fusion, and include two genes MFa1 and MFa2 for the MATa-specific pheromone called

mating factor a (MFa), and the gene for the mating factor a (MFa) receptor (STE3). The gene product of a2 is a homeodomain protein that acts as a negative regulator of transcription of the MATa-specific genes. MFa1 and MFa2 that code for the MATa specific MFa, and the gene for the MFa receptor (STE2; Fig. 5.1). The MATa locus of MATa cells encodes only one functional protein, a1, a transcription factor with a homeodomain. However, unlike the a2 protein of MATa cells, a1 does determine the MATa cell type. In the absence of the a2 protein, a-specific genes are expressed constitutively and a-specific genes are not expressed because of the absence of the positive regulator a1. The second gene a2 of the MATa idiomorph encodes a nonfunctional version of a2 (Dolan and Fields 1991). In diploid MATa/a cells, the homeodomain proteins a2 and a1 interact to form a novel negative regulatory protein that is functional during mating and meiosis (Herskowitz 1989; Fig. 5.1). In filamentous ascomycetes, mating types were originally described as + or –, but were also called either a and a or A and a. According to the nomenclature of Turgeon and Yoder (Turgeon and Yoder 2000), The MATa-like idiomorph is termed MAT1-1 and the MATa-like idiomorph is termed MAT1-2 (Debuchy and Turgeon 2006; Debuchy et al. 2010). In most cases, homothallic filamentous ascomycetes contain either linked or unlinked sequences similar to both the MAT1-1 and the MAT1-2 idiomorph (Lin and Heitman 2007; Po¨ggeler et al. 1997). This review will start from the well studied pheromone/receptor system of S. cerevisiae and will give an overview of the current molecular knowledge on the structure, evolution and function of pheromone/receptor systems in filamentous ascomycetes. a-factor receptor

a-factor receptor STE3

STE2

a–cell

Fig. 5.1. Signaling between MATa and MATa cells of Saccharomyces cerevisiae. Mating-type loci encode transcription factors (a1, a1, a2) which determine the cell type and regulate the transcription of mating-type specific pheromones and pheromone receptors

a–cell a-factor

MATa-Locus

a2 a2

a1 a1

a-factor

MATa-Locus (a2)

a1 a1

Function and Evolution of Pheromones and Pheromone Receptors in Filamentous Ascomycetes

II. Pheromones and Pheromone Receptors in S. cerevisiae Conjugation between two haploid yeast cells is generally controlled by the reciprocal action of diffusible mating pheromones, with cells releasing pheromones that induce mating-specific changes in cells of the opposite type. The underlying pheromone/receptor system is best understood in S. cerevisiae. A. Pheromones of S. cerevisiae As early as 1956, Levi observed that the mating reaction of S. cerevisiae MATa cells does not require initial cell contact and showed that a morphological response, the formation of one or more projections on the cell surface (commonly termed shmoo morphology) is caused by a diffusible substance (Levi 1956). Duntze et al. (1970) confirmed this observation, partially purified the diffusible factor from filtrates of liquid cultures of MATa cells and showed that it inhibits cellular division in MATa cells in the Gl phase of the cell cycle (Bu¨cking-Throm et al. 1973). Later, the diffusible factor was termed MFa. Duntze and co-workers determined that MFa is the 13-amino-acid (aa) peptide: WHWLQLKPGQPMY (Sto¨tzler and Duntze 1976; Stotzler et al. 1977). When the tridecapeptide sequence was chemically synthesized, it was found to exhibit all the properties of the natural pheromone (Ciejek et al. 1977). Two genes (MFa1 and MFa2, specifically expressed in MATa cells) encode precursors that contain four and two tandem repeats of the tridecapeptide, respectively (Brake et al. 1983; Kurjan and Herskowitz 1982; Singh et al. 1983). MFa is secreted by the classical yeast secretory pathway (Julius et al. 1983; Singh et al. 1983). The first step of MFa precursor processing is the removal of the signal sequence from the pre-pro-MFa factor (Waters et al. 1988). Further processing requires the KEX2 endoprotease, which cleaves the pro-protein at characteristic KR dipeptides (Julius et al. 1984), the dipeptidyl aminopeptidase STE13, which recognizes the dipeptide XA or XP (Julius et al. 1983) and the carboxypeptidase KEX1, which removes C-terminal extensions (Cooper and Bussey 1989; Fig. 5.2). Similar to MFa, partially purified fractions exhibiting MFa activity have been obtained from culture filtrates of a MATa cells by ultrafiltration,

75

Signalpeptidase SS

13 13

13

13

Endopeptidase Kex2p

13 13

13

13

Carboxypeptidase Kex1p

13 13 13

13

Aminopeptidase Ste13p

13 13 13

13

13 13

13

13 = WHWLQLKPGQPMY

13

Tridecapeptide (mature pheromone)

SS = Signal sequence

= Dipeptide KR = Dipeptide XA or XP

Fig. 5.2. Processing of the S. cerevisiae pre-pro-pheromone MFa1

ion-exchange chromatography and gel filtration. MFa preparations specifically cause both G1 arrest and morphological alterations in MATa cells (Betz et al. 1977). Tandem mass spectrometry revealed that MFa is a mixture of carboxylated and farnesylated dodecapeptides (YIIKGV/LFWDPAC[Farnesyl]-OCH3). The posttranslational modification with farnesyl and carboxymethyl groups is required for full biological activity of MFa (Marcus et al. 1991). Similar to MFa, MFa is encoded by two genes, MFa1 and MFa2, encoding precursor proteins of 36 and 38 aa, respectively (Brake et al. 1985). Neither precursor contains a signal sequence, but both have a C-terminal CaaX farnesylation motif (where C is cysteine, a is an aliphatic residue and X is any amino acid). Genetic analysis of sterile (ste) mutants that disturb the mating process (Hartwell 1980) and mutants deficient in ras and MFa modification (ram; Powers et al. 1986) led to the elucidation of the MFa biosynthesis reviewed by Cladwell et al. (1995; see Fig. 5.3). Farnesyltransferase, composed of an a subunit encoded by ram2 and a b subunit, encoded by ram1, leads to the farnesylation of the MFa precursor. The C-terminal CaaX protease activitiy corresponds to the endoplasmic reticulum membrane-localized RCE1 and STE24 proteases (Schmidt et al. 1998). The carboxymethylation of the C-terminal cysteine residue is mediated by STE14 (Hrycyna et al. 1991). N-terminal processing involves two sequential

Stefanie Po¨ggeler

76

MCPSATAAPKETSSEKKDNYIIKGVFWDPACVIA SH RAM1

Isoprenylation

RAM2

+

Cytoplasm

MCPSATAAPKETSSEKKDNYIIKGVFWDPACVIA

Cleavage of aaX

STE24

VIA

or

S

MCPSATAAPKETSSEKKDNYIIKGVFWDPAC

Carboxy-Methylation

CH3

S

STE14

MCPSATAAPKETSSEKKDNYIIKGVFWDPAC CH3 STE24

N-terminal Processing MCPSATAAPKETSSEKKDN

S

and AXL1

Plasmamembrane

RCE1

YIIKGVFWDPAC CH3 S STE6

Secretion

YIIKGVFWDPAC CH3 S

mature lipopeptide

Extracellular

Fig. 5.3. Processing of the S. cerevisiae pheromone precursor MFa1

proteolytic cleavages by STE24 and AXL1 (Huyer et al. 2006). Finally, the mature lipopeptide pheromone is exported via the ATP-binding cassette transporter STE6 (Michaelis 1993; Fig. 5.3). Experience with purification of the S. cerevisiae pheromones facilitated purification of pheromones and the identification of the pheromone precursor genes in the fission yeast Schizosaccharomyces pombe M (minus) cells produce a farnesylated, carboxylated 9-aa peptide (YTPKVPYMC[Farnesyl]-OCH3), the M-factor; while P (plus) cells produce an unmodified P-factor peptide of 23 residues (TYADFLRAYQSWNTFVNPDRPNL; Davey 1992; Imai and Yamamoto 1994; Wang et al. 1994). Two genes, MFm1 and MFm2, encode the M-factor precursor which is posttranslationally cleaved and modified to yield the active pheromone (Davey 1992). The P-factor precursor is the product of the map2 gene. It carries a hydrophobic signal sequence and four tandem repeats of the mature 23-aa peptide. The P-factor precursor is predicted to undergo processing events similar to the S. cerevisiae a-factor precursor (Imai and Yamamoto 1994).

B. Pheromone Receptors of S. cerevisiae In S. cerevisiae, the pheromone signal is sensed by the MATa cell-specific MFa receptor STE2 and the MATa cell-specific MFa receptor STE3 (Fig. 5.1; Burkholder and Hartwell 1985; Hagen et al. 1986). Both pheromone receptors are G protein-coupled receptors (GPCRs), which are the largest and most diverse superfamily of membrane receptors.

GPCRs are characterized by seven helical transmembrane (TM) domains and the ability to activate heterotrimeric G proteins (Gabg). The seven TM domains are connected by three extracellular and three intracellular loops. The N-terminus is oriented towards the extracellular space while the C-terminus is intracellular (Pierce et al. 2002). Because of their low level of expression in native tissue and poor thermal stability the crystal structure of only four GPCRs have been solved so far. These include the human b2 adrenergic receptor, the avian b1 adrenergic receptor, the human A2A adenosine receptor and the bovine rhodopsin, reviewed by Rosenbaum et al. (2009).

Like all other GPCRs, activation of STE2 by MFa and STE3 by MFa leads to well characterized G protein-mediated mitogen-activated protein (MAP) kinase signal transduction events. Upon pheromone binding, STE2 and STE3 switch from an inactive to an active conformation. Both receptors thereby activate the GPA1 Ga subunit, triggering the exchange of GDP for GTP on GPA1. This leads to the dissociation of the G protein from the receptor and of the Ga subunit from the Gbg complex. The released Ga-GTP and the Gbg complex then activate downstream effectors (Hamm 1998). Mutagenesis studies indicate that STE2 is similar to other GPCRs in that the third intracellular loop plays an impor´ elic´ et al. 2003; Choi and tant role in G protein activation (C Konopka 2006; Stefan and Blumer 1994).

Function and Evolution of Pheromones and Pheromone Receptors in Filamentous Ascomycetes

In S. cerevisiae, pheromone receptor activation results in G1 cell cycle arrest and polarized growth of the mating projection. Subsequently cell contact, fusion and karyogamy lead to the formation of the diploid zygote (Bardwell 2005; Dohlman and Slessareva 2006; Elion et al. 2005). In addition to its function in the initial pheromone response leading to G1 arrest that involves the well analyzed MAP kinase signal transduction pathway, the STE2 receptor was recently implicated at the level of cell wall degradation as a possible modulator of mating efficiency (Shi et al. 2007). Shi et al. (2007) demonstrated that truncation of the 20 N-terminal amino acid residues reduced mating efficiency, while maintaining a wild-type-like G1 arrest signaling. MATa cells expressing the truncated STE2 receptor arrest mating at the prezygotic stage after cell contact. The STE2 receptor was implicated in modulation of the interaction between activated STE2 and STE3 receptors in a hypothesis that N-terminal, extracellular residues of STE2 mediate novel receptor interactions with ligands or proteins, leading to alternate signaling effector pathways (Shi et al. 2009). According to Xue et al. (2008), fungal GPCRs can be classified into six classes based on sequence homology and ligand sensing. The two S. cerevisiae receptors, STE2 and STE3, exhibit no sequence similarity, aside from a general enrichment of basic residues in the third intracellular loop, even though they are known to activate the same cytoplasmic Ga protein (GPA1) in response to pheromone binding. STE2 and STE3 belong to GPCR classes of STE2-like pheromone receptors and STE3-like pheromone receptors, respectively (Xue et al. 2008).

III. Structure and Evolution of Pheromones and Pheromone Receptors in Filamentous Ascomycetes Recent genome sequencing projects and sequencing of expressed sequence tag (EST) databases, allowed the identification of pheromone precursor genes and pheromone receptor genes in several species of filamentous ascomycetes. These include not only heterothallic and homothallic filamentous ascomycetes, but also species that were

77

originally assumed to be asexual (Galagan et al. 2005; Hoff et al. 2008; Machida et al. 2005; Pel et al. 2007; Po¨ggeler 2000, 2002; Po¨ggeler and Ku¨ck 2001; Po¨ggeler et al. 2008; Woo et al. 2006). Similar to S. cerevisiae, filamentous ascomycetes encode two types of pheromone receptors and pheromones, which are related to STE2 and STE3 receptors and MFa and MFa precursors, and all four genes are present in strains of both mating types. Only MFa-like pheromones and STE3-like receptors have been identified in basidiomycetes. In contrast to ascomycetes, the genes encoding pheromones and receptors are linked to a matingtype locus (Bo¨lker et al. 1992; O’Shea et al. 1998; Vaillancourt et al. 1997; Wendland et al. 1995) (see Chapter 6 in this volume). A. Pheromones of Filamentous Ascomycetes The first pheromone genes of filamentous ascomycetes were cloned in the plant pathogen Cryphonectria parasitica, which causes chestnut blight. A natural biological control of the plant disease can be achieved with a mycovirus that infects C. parasitica. The virus causes a perturbation of fungal development and results in low virulence (hypovirulence). The virus was demonstrated to affect the transcription of several fungal genes, among them two genes, initially termed vir1 and vir2, which are produced abundantly in non-infected strains but are not detectable in virus-infected strains (Powell and Van Alfen 1987; Zhang et al. 1993). Interestingly, vir1 and vir2 were later recognized encoding pheromone precursors related to the MFa precursors of S. cerevisiae. The genes have therefore been renamed Mf2/1 and Mf2/2. Sequence comparison of both genes revealed that they contain an identical 83bp open reading frame (ORF), encoding a small 23-aa lipopeptide pheromone precursor with a Cterminal CaaX motif (Zhang et al. 1998). The Mf1/ 1 gene was cloned by PCR-mediated subtractive hybridization using cDNA libraries of MAT1-1 and MAT1-2 strains. A MAT1-1 specific clone encoding a 530-aa protein with seven repeats of the decapeptide WCLFHGEGCW was identified (Table 5.1). Similar to the S. cerevisiae MFa precursor, the MF1/1 protein contains an N-terminal hydrophobic signal sequence and all seven repeats of the decapeptide are preceded by repeats of the dipeptide XA (or XP) and have the dipeptide KR at the C-terminus, with KEKR or KVKR in some

Stefanie Po¨ggeler

78

Table 5.1. Mfa-like pheromones of filamentous ascomycetes

Order

Species

Mfa-like pheromone precursor (name/length, aa/accession number)

Eurotiales

Aspergillus nidulans A. fumigatus A. terreus A. flavus A. niger A. clavatus Neosartorya fischeri Penicillium chrysogenum Verticillium albo-atrum

PPGA/128/Q7SI74 PPGA/102/Q4WNH4 PPGA/136/Q0CFX7 PPGA/103/B8NF30 PPGA/108/A2RBH8 PPGA/105/A1CE26 PPGA/102/A1DM33 PPG1/111/B1GVB6 PPG1/174/C9SJ18

Cryphonectria parasitica Sclerotinia sclerotiorum Botryotinia fuckeliana Pyrenophora triticirepentis

MF1-1/530/AAC39328.1 PPG1/326/A7EFR4 PPG1/392/A6RID0 PPG1/162/B2VZL9

Pezizales

Phaeosphaeria nodorum Tuber melanosporum

PPG1/182/Q0UXN0 PPG1/135/D5G4G7

Hypocreales

Nectria haematococca

PPG1/294/C7Z625

Gibberella zeae

PPG1/421/FGSG_05061.3a

Fusarium oxysporum

PPG1/524/FOXG_08636.2a

F. verticilloides

PPG1/594/FVEG_06038.3b

Trichoderma atroviride

PPG1/188/Triat2|297240| estExt_fgenesh1_kg. C_contig_140145 ** PRE2/296/Trire2|64018| e_gw1.13.399.1b PPG1/227/A4QYP1 Mf1/1/530/AAC39328.1 PPG1/277/Q9P3Y6 PPG1/299/Q01301 PPG1/290/Neute_mat_a1| 143059| estExt_fgenesh2_kg. C_10700** PPG1/304/Neudi1|92063| estExt_Genewise1Plus. C_21207** MFM/221/Q5MGQ6 PPG1/154/Q2HF71

Phyllochorales Diaporthales Helotiales Pleosporales

T. reesei Magnaporthales Magnaporthe grisea Diaporthales Cryphonectria parasitica Sordariales Sordaria macrospora Neurospora crassa N. tetrasperma

N. discreta Podospora anserina Chaetomium globosum

Predicted sequence of the peptide pheromone (length/repeats) WCRFAGRIC (9/1x)WCRFRGQVC (9/1x) WCHLPGQGC (9/2x) WCWLPGQGC (9/2x) WCSLPAQGC (9/2x) WCVLPGQPC (9/2x) WCELPGQGC (9/2x) WCHLPGQGC (9/2x) WCGHIGQGC (9/2x) GKPCPRPGQGCW (12/2x) QKPCPRPGQGCW (12/2x) WCLFHGEGCW (10/7x) WCGRPGQPC (9/11x) WCGRPGQPC (9/15x) WTQKRPYGMPVG (12/1x) WTQHRPYGMPVG (12/2x) WTQKRPYGMPVG (12/1x) WRQKRPYGMPVG (12/1x) GWRYRPYGLPVG (12/5x) WTPRPGRGAY (10/2x)WHPRAGPPAY (10/1x)WHPRAGDPMA (10/1x) WEVNGNRGVY (10/1x) WCFYPGQPCW (10/6x)WGFFPGQSQW (10/1x) WCTWKGQPCW (10/4x)WCWWKGQPCW (10/4x) WCTWRGQPCW (10/6x)WCLWRGQPCW (10/2x)WCMWRGQPCW (10/2x) WCTWRGQPCW (10/7x)WCMWRGQPCW (10/5x) WCWRVGESCW (10/4x) WCYRIGEPCW (10/4x)WCWILGGKCW (10/1x) WCLFHGEGCW (10/7x) QWCRIHGQSCW (11/5x) QWCRIHGQSCW (11/5x) QWCRIHGQSCW (11/5x)

QWCRIHGQSCW (11/5x) QWCLRFVGQSCW (12/2x) WCKQFLGMPCW (11/1x)WCTQFLGQPCW (11/1x)FCTRFTGSSCW (11/1x)

a

Broad Institute DOE Joint Genome Institute

b

cases. The posttranslation processing of both pheromone precursors was concluded to be similar to that of S. cerevisiae. Northern blot analyses revealed MAT1-2 specific expression of Mf2/1 as

well as of Mf2/2, and MAT1-1 specific expression of Mf1/1. Furthermore, Southern blot analysis of C. parasitica genomic DNA demonstrated that MAT1-1 strains carry non-expressed copies of

Function and Evolution of Pheromones and Pheromone Receptors in Filamentous Ascomycetes

Mf2/1 and Mf2/2 while a non-expressed copy of Mf1/1 exist in MAT1-2 strains (Zhang et al. 1998). These results showed that, unlike basidiomycetes, the pheromone genes of filamentous ascomycetes are not encoded by the mating-type locus. Two mating type-specific expressed pheromone precursor genes were also isolated from the plant pathogen Magnaporthe grisea by a subtractive cloning strategy (Shen et al. 1999). Initially, the M. grisea lipopeptide pheromone precursor gene was termed MF1-1 and the gene encoding a precursor protein processed by the Kex2-like protease was termed MF2-1 (Shen et al. 1999). However for consistency, the MF1-1 gene is termed here MF2-1, and MF1-1 is termed MF2-1.

The M. grisea MF2-1 gene encodes a 26-aa polypeptide that is processed to produce a lipopeptide. MF1-1 encodes a precursor with four repeats of an undecamer sequence that is predicted to yield a peptide pheromone with striking similarity to the predicted Mf1/1 pheromone precursor sequence of C. parasitica (Table 5.1). Conserved sequences of the known pheromone genes from C. parasitic and M. grisea were later used to extract putative pheromone precursor gene sequences by a tblastn search from the Neurospora crassa EST libraries. The DNA sequences of the identified sequences were used to design primers and amplify pheromone precursor genes from the homothallic ascomycete Sordaria macrospora, which is a close relative of N. crassa (Po¨ggeler 2000). The putative pheromone genes of N. crassa and S. macrospora were named ppg1 and ppg2. In N. crassa, ppg1 is also termed ccg-4 because the gene was first identified as a gene that is expressed under control of the circadian biological clock (Bell-Pedersen et al. 1996). The ppg2 gene of N. crassa is also termed mfa-1 (mating factor expressed in mat a strains; Bobrowicz et al. 2002).

The ppg1 gene is predicted to encode an MFalike precursor that is processed by a KEX2-like protease to yield the mature peptide pheromone, while ppg2 encodes a short 24-aa polypeptide with a putative farnesylation motif (Po¨ggeler 2000). Within the N. crassa and S. macrospora PPG1 precursor, the same five repeats of a decapeptide motif can be identified: QWCRIHGQSCW (Table 5.1). Moreover, the coding sequence of the N. crassa ppg2 gene is 100% identical to the

79

S. macrospora ppg2 coding region. Thus, both fungi seem to secrete an identical set of pheromones. The expression of the pheromone precursor genes is mating-type-specific and under the control of the MAT locus in the heterothallic N. crassa, whereas transcripts of ppg1 and ppg2 are co-expressed during all stages of the homothallic S. macrospora life cycle (Bobrowicz et al. 2002; Po¨ggeler 2000). In the heterothallic Podospora anserina, genetic data indicate that both pheromone genes are transcribed at low levels in vegetative hyphae of both mating types. However, their transcription is induced by the mating types in male spermatia (Coppin et al. 2005). Furthermore, Coppin et al. (2005) showed that constitutively expressed pheromones gene constructs are repressed at the posttranscriptional level by the non-cognate mating type.

1. Characteristics of MFa-Like Peptide Pheromones Genes encoding MFa-like pheromone genes have been identified in different orders of filamentous ascomycetes (Table 5.1). A common feature of all MFa-like pheromone precursors is 2–15 repeats of a presumed mature pheromone. While members of the order Eurotiales encode MFa-like precursors with only two repeats, the MFa-like precursor of Botrytis cinerea carries 15 copies of a nonamer sequence (Table 5.1). The length of the repeated sequence and therefore the predicted mature peptide pheromone varies between 9 and 12 aa. Within a given precursor the length of the repeated sequence is constant; however, some positions may vary within the repeated copies. The highest degree of sequence variation can be found in the Chaetomium globosum PPG1 precursor. The three copies of the C. globosum undecapeptide motif vary at six positions ([WF]C[KT] [QR]F[LT]G[MQS][PS]CW), while in the B. cinerea 15 copies of the nonamer motif are identical (Table 5.1). Structure–activity studies of S. cerevisiae MFa by the systematic substitution of each residues with alanine revealed that the N- and C-termini of the tridecamer play important roles in receptor activation and receptor binding, respectively, while the central residues of MFa orient the signaling and binding domains of the pheromone (Naider and Becker 2004; Fig. 5.4A).

Stefanie Po¨ggeler

80

Table 5.2. STE2- and STE 3-like pheromone receptors in ascomycetes Order

Species

Saccharomycetales

STE2-like receptor (name/length, STE2-like receptor (name/ aa/accession number) length, aa/accession number)

Saccharomyces cerevisiae Kluyveromyces lactis Ashbya gossypii Candida albicans C. glabrata Pichia stipitis Vanderwaltozyma polyspora Schizosaccharomycetales Schizosaccharomyces pombe S. japonicus Pneumocystidales Pneumocystis carinii

STE2/431/P06842 STE2/436/Q6CIP0 STE2/458/Q752Q1 STE2/470/Q59Q04 STE2/4 17/Q6FLY8 STE2/321/A3LXU7 STE2/473/A7TQX4

STE3/470/NP_012743.1 STE3/435/XP_451291.1 STE3/455/NP_984730.1 STE3/417/XP_718594.1 STE3/452/XP_449701.1 STE3/300/XP_001387433.1 STE3/477/XP_001646391.1

NAM2/348/Q00619

MAP3/365/NP_593942.1

NAM2/346/B6JZE2 PRE2/354/A2TJ26

Eurotiales

Aspergillus nidulans A. fumigatus A. oryzae A. terreus A. flavus A. niger A. clavatus Neosartorya fischeri Penicillium chrysogenum P. marneffei Talaromyces stipitatus Verticillium albo-atrum Ajellomyces capsulata A. dermatitidis Paracoccidioides brasiliensis Arthroderma benhamiae Nannizzia otae Trichophyton verrucosum Uncinocarpus reesii Coccidioides posadasii Sclerotinia sclerotiorum Botryotinia fuckeliana Pyrenophora triticirepentis Phaeosphaeria nodorum Tuber melanosporum

PREB/377/Q7SI72 PREB/369/Q4WYU8 PREB/369/Q2U819 PREB/373/Q0CS34 PREB/374/B8NF30 PREB/378/A2QU32 PREB/370/A1CLD3 PREB/369/A1D5Z2 PRE2/380/B1GVB8 PREB/357/Q0MR03 PREB/359/B8M557 PRE2/377/C9SGY3 PREB/357/A6QUU6 PREB/355/C5JV36 PREB/356/C1H517

MAP3/360/XP_002172594.1 STE3a/439/AAG38536.1; STE3b/336/AF309805.1a PREA/349/DAA01795.1 PREA/460/XP_753848.1 PREA/465/XP_001823505.1 PREA/442/XP_001216959.1 PREA/465/XP_002378906.1 PREA/523/XP_001390270.1 PREA/442/XP_001274260.1 PREA/407/XP_001259896.1 PRE1/469/XP_002565481.1 PREA/448/XP_002145523.1 PREA/354/XP_002487009.1 PRE1/316/XP_003000881.1 PREA/432/EEH10416.1 PREA/431/XP_002621095.1 PREA/452/EEH23272.1

PREB/369/D4AND1 PREB/369/C5FBT2 PREB/369/D4DBK6 PREB/358/C4JL18 PREB/362/C5PF60 PRE2/395/A7EY95 PRE2/380/A6SLH2 PREB/345/B2WIP5

PREA/455/XP_003015770.1 PREA/432/XP_002849417.1 PREA/406/XP_003025539.1 PREA/456/XP_002544713.1 PREA/371/XP_003069522.1 PRE1/607/XP_001592017.1 PRE1/571/XP_001554250.1 PRE1/453/XP_001931407.1

PREB/370/Q0UCT8 PRE2/367/D5GJK5

Nectria haematococca Gibberella zeae Fusarium oxysporum

PRE2/386/C7ZA34 PRE2/386/XP_382831.1 PRE2/380/supercontig 2: 2453834-2455018b PRE2/354/supercontig 6: 672208673392b PRE2/294/Triat2|17079|Triat1. gw1.3.1314.1d PRE2/296/Trire2|64018| e_gw1.13.399.1d PRE2/415/A4QV61 PRE2/554/Q8WZM9 PRE2/568/D2N2E6 PRE2/571/D2N2E9 PRE2/576/D2N2F3 PRE2/568/D2N2F4

PRE1/485/XP_001799363.1 PRE1/374/ GSTUMT00012510001c PRE1/375/XP_003052288.1 PRE1/374/XP_387446.1 PRE1/494/FOXG_02147b

Phyllochorales Onygenales

Helotiales Pleosporales Pezizales Hypocreales

F. verticilloides Trichoderma atroviride T. reesei Magnaporthales Sordariales

Magnaporthe grisea Sordaria macrospora Neurospora crassa N. intermedia N. sitophila N. tetrasperma

PRE1/238/FVEG_05310.3b PRE1/371/Triat2|147894|d PRE1/356/Trire2|57526| e_gw1.4.314.1d PRE1/456/XP_369937.1 PRE1/717/CAC87067.2 PRE1/724/CAO78593.1 PRE1/724/CAO78581.1 PRE1/724/CAO78582.1 PRE1/724/CAO78586.1 continued

Function and Evolution of Pheromones and Pheromone Receptors in Filamentous Ascomycetes

81

Table 5.2. continued Order

Species N. discreta Podospora anserina Chaetomium globosum

STE2-like receptor (name/length, STE2-like receptor (name/ aa/accession number) length, aa/accession number) PRE2/577/Neudi1|140892| fgenesh3_pm. C_scaffold_9000121d PRE2/465/B2ADL1 PRE2/346/Q2GU85

PRE1/722/CAO78599.1 PRE1/624/XP_001908273.1 PRE1/666/XP_001221914.1

a

Alternative splice product of P. carinii STE3a Broad Institute c http://mycor.nancy.inra.fr/IMGC/TuberGenome/index.html d DOE Joint Genome Institute b

For S. macrospora the predicted signal sequence of the PPG1 precursor was demonstrated to be sufficient to enable fused green fluorescent protein (GFP) to pass through the secretory pathway (Mayrhofer and Po¨ggeler 2005).

The repeated sequences of MFa-like pheromone precursors are surrounded by maturation signals similar to those of the S. cerevisiae MFa precursor (Fig. 5.2). Usually, the repeated motifs are preceded by two to several copies of the dipeptide XA or XP, a putative signal for cleavage by a STE13-like dipeptidyl aminopeptidase. These dipeptide repeats are mostly preceded by the diba-

A Loop domain P8

G9

K7

Q10

L6

P11

Q5

M12

L4

Y13

W3

Binding domain

H2 W1 Signaling domain

B 4 3 bits

Motif searches by the MEME program (http:// meme.sdsc.edu) using the 143 identified repeated copies of MFa-like pheromone precursors listed in Table 5.1 resulted in an 8-aa consensus motif (CR[RW]PGQPC; Fig. 5.4B). Similar to S. cerevisiae MFa, peptide pheromones of filamentous ascomycetes contain an invariant glycine residue in the center of the peptide pheromone. Moreover, the adjacent amino acid residues, Gln and Pro, seem to be conserved (Fig. 5.4). Analysis of S. cerevisiae MFa indicated that the central Pro-Gly sequence is involved in the pheromone biological activity and that this region created the proper overall conformation of the molecule (Naider and Becker 2004). A similar function may attributed to the conserved central residues of MFa-like peptide pheromones of filamentous ascomycetes. An N-terminal signal sequence is predicted for nearly all of the predicted precursors by signal P (http://www.cbs.dtudk). Missing N-terminal secretion signals may be explained by genome annotation errors. Thus, in filamentous ascomycetes, MFa-like pheromones seem to be secreted via the classical secretion pathway.

2 1 0

1

2

3

4

5

6

7

8

Fig. 5.4. MFa-like pheromones of filamentous ascomycetes. (A) Cartoon representing functional segmentation of the S. cerevisiae MFa. According to Naider and Becker (2004) the MFa can be dived into a signaling, loop (conformational hinge) and binding domains. (B) Consensus sequence by NEME (http://meme.sdsc.edu; E-value 2.1e-470) as determined from the 143 repeats of MFa-like pheromones of filamentous ascomycetes listed in Table 5.1

sic sequence KR or RR, a cleavage signal of the KEX2-like endopeptidase (Bader et al. 2008; Brenner and Fuller 1992; Julius et al. 1984). In some cases, however, the repeat is directly preceded by a KEX2 signal sequence. Only pheromone

82

Stefanie Po¨ggeler

precursors of Eurotiales and Pleosporales deviate from this principle. Here, repeats of MFa precursors are not preceded by the dipeptide KR or by copies of the dipeptide XA or XP. Often the pheromone precursors contain additional KEX2 cleavage sites downstream of the repeats of the predicted mature pheromone sequence. The C-terminus of the S. cerevisiae MFa precursor is processed by the carboxypeptidase KEX1 (Fig. 5.2), which removes basic residues left by KEX2 cleavage. In the MFalike precursors of filamentous ascomycetes, additional residues often lie between the KEX2 site and the predicted pheromone sequence. These residues were assumed to also be removed during C-terminal processing by a KEX1-like enzyme (Shen et al. 1999). Alternatively, the repeats may produce pheromones with additional C-terminal residues. Using a heterologous yeast assay it has been shown that the S. macrospora PPG1 precursor is processed into an active pheromone by S. cerevisiae MATa cells (Mayrhofer and Po¨ggeler 2005).

Genes involved in pheromone processing of the a-factor precursor in S. cerevisiae such as KEX1, KEX2 and STE13 have homologues in the genomes of filamentous ascomycetes (Hoff et al. 2008; Mayrhofer and Po¨ggeler 2005; Po¨ggeler 2002). Thus, processing of the MFa-like pheromone precursors is likely to have been conserved in filamentous ascomycetes. In contrast to S. cerevisiae, only one MFa-like pheromone precursor gene has been identified in the genomes of all filamentous ascomycetes analyzed to date. 2. Characteristics of MFa-Like Lipopeptide Pheromones The identification of genes encoding MFalike pheromone precursors is difficult because: (i) MFa-like pheromones are poorly conserved, (ii) small peptide-encoding ORFs<100 aa are often not correctly annotated in fungal genomes and (iii) the genomes of filamentous ascomycetes contain many putative small ORFs with the potential to be prenylated at the C-terminus. As mentioned above, the first MFa-like pheromone precursor genes of filamentous were isolated by chance or cloned by PCR-mediated subtractive hybridization using cDNA libraries of MAT1-1 and MAT1-2 strains from C. parasitica and M. grisea (Shen et al. 1999; Zhang et al. 1998).

To date, MFa-like precursors have been identified in only 11 different filamentous ascomycetes (Fig. 5.5). Genes encoding MFa-like pheromones are identified by their relatively close phylogenetic relationship to C. parasitica and M. grisea using tblastn searches of fungal genomes sequence and EST libraries or using degenerated primers for PCR (Coppin et al. 2005; Po¨ggeler 2000). In some filamentous ascomycetes genomes, the lipopeptide-encoding gene was found in the vicinity of a cyanate lyase gene (Elleuche and Po¨ggeler 2008). Therefore, an alternative strategy to isolate MFa-like pheromone precursor genes has been to analyze this genomic region in detail (Kim et al. 2008; Lee et al. 2008; Schmoll et al. 2010). In the yeast Candida albicans, comparisons of genome sequences of closely related species was successfully used to identify a gene encoding an MFa-like mating factor. Searching the C. albicans genome for short ORFs with a cysteine four residues from the C-terminus identified more than 1000 ORFs. However, performing the same analysis with the genomic sequence of the closely related C. dubliniensis identified only one candidate protein with an identical sequence, and the encoding gene was syntenic in both species. The gene was named CaMFA1 and homologues were also identified in C. guilliermondii, C. tropicalis, C. lusitaniae and C. paralopsiloisis. Disruption of CaMFA1 blocks mating in MTLa cells but not in MTLa cells of C. albicans (Dignard et al. 2007).

Within closely related species, the lipopeptide pheromone genes are highly conserved. The amino acid sequence of the lipopeptide precursor from S. macrospora, N. crassa and N. tetrasperma are 100% identical. However, lipopeptide precursors are poorly conserved among more distantly related filamentous fungi (Fig. 5.5). As in ascomycetous yeasts and basidiomycetes, common features of MFa-like pheromone precursors from filamentous ascomycetes are the C-terminal CaaX motif that acts as a recognition site for isoprenylation, and the short precursor length. MFa-like pheromone precursors identified in filamentous ascomycetes are only 21–32 aa (Fig. 5.5). Analysis of the MFa-like pheromones shown in Fig. 5.5 by MEME (http://meme.sdsc. edu) revealed a 9-aa conserved motif comprising the CaaX motif and five preceding residues (Fig. 5.5B). In addition to the invariant C four amino acids from the end, the last residue is an invariant methionine and the penultimate residue an aliphatic amino acid.

Function and Evolution of Pheromones and Pheromone Receptors in Filamentous Ascomycetes

The last X residue of the CaaX motif is proposed to determine specificity for farnesyltransferases (methionine, serine, glutamine, alanine) or geranylgeranyltransferase type I (leucine, phenylalanine; Moores et al. 1991). Thus, the MFa-like A

83

pheromones of filamentous ascomycetes are predicted to be farnesylated. Farnesyltransferases seem to recognize the sequence downstream of the conserved cysteine as a set of two or three co-operative, interconnected

Sordariales Neurospora crassa PPG2 (AAN03594.1)

MPSTAASTKVPQTTMNFNGYCVVM Sordaria macrospora PPG2 (CAB96171.1)

MPSTAASTKVPQTTMNFNGYCVVM Neurospora tetrasperma scaffold_2: 3376878:3376949*

MPSTAASTKVPQTTMNFNGYCVVM Podospora anserina MFP (Q5MGQ5)

MPSTTAQTKVPQTSTNF NSYCVVM Chaetomium globosum PPG2 Supercontig 2: 1423081-1423152 - **

MPSTTAQTKVPQTSTNF NSYCVVM Magnaporthales Magnaporthe grisea MF2/1 (A4R2S0)

MSPSTKNIPAPVAGARAGPIHYCVIM Diaporthales Cryphonectria parasitica MF2/1 (AAC39329.1)

MPSNTQTSNSSMGVNGYSYCVVM Pleosporales Phaeosphaeria nodorum PPG2 Supercontig 10: 445402-445458 +**

MSETGGFSLAVEGYRRCLVM Hypocreales Gibberella zeae PPG2 (Schmoll et al. 2010)

MPSTKPTSSQKPGYPLSCTVM Epichloe festuca_PPG2 (Schmoll et al. 2010)

MPSTTNNQATQVTTASSEENAHTASYVGCTIM *DOE Joint genome institute; JGI **Broad Institute

B 4 3 bits

Fig. 5.5. MFa-like pheromones of filamentous ascomycetes. (A) Primary structure of MFa-like pheromones of species of the orders Sordariales, Magnaporthales, Diaporthales, Pleosporales and Hypocreales. The accession numbers are given in brackets or are given according to Schmoll et al. (2010). The conserved NF dipeptide of MFa-like precursors in species of the order Sordariales is indicated in bold italics. (B) Consensus sequence by NEME (http://meme.sdsc.edu; E-value 6.3e-12) as determined from MFa-like pheromone precursors shown in A

2 1 0

1

2

3

4

5

6

7

8

9

84

Stefanie Po¨ggeler

recognition elements as opposed to three independent amino acids (Hougland et al. 2009). Therefore, the internal a-residues of the CaaX motif do not necessarily need to be aliphatic. In species in the order Hypocreales, the third last residue is a polar, uncharged threonine (Fig. 5.5). Bioinformatic analysis and biochemical studies of known substrates of farnesyltransferases indicate that sequences immediately upstream of the conserved cysteine residue may also play a role in farnesytransferases substrate selectivity (Hicks et al. 2005; Maurer-Stroh and Eisenhaber 2005). While the requirement for specific amino acids at single positions appears minimal outside of the CaaX box, physicochemical constraints that extend up to 11 residues N-terminal from the modified cysteine can be found (Maurer-Stroh and Eisenhaber 2005). MFa-like pheromone precursors of filamentous predominantly contain a tyrosine residue at the –1 position (Fig. 5.5). In S. cerevisiae and basidiomycetes, the Nterminus of the lipopeptide pheromone precursor is removed by the STE24 and AXL1 proteases (Fig. 5.3). The cleavage site between the Nterminal extension and the mature lipopeptide pheromone has a conserved motif for both the S. cerevisiae MFa and the S. pombe M-factor with cleavage occurring between asparagine and tyrosine. Only MFa-like precursors of species of the order Sordariales contain a conserved Asn-Phe motif that might serve as a cleavage site. However, this motif is not conserved in MFa-like pheromone precursors from other filamentous ascomycetes (Fig. 5.5). Extensive tblastn analysis failed to identify a MFa-like pheromone precursor in several filamentous ascomycetes. This might be because some species either lack MFa-like pheromone encoding genes, or MFa-like genes were not identified because they are very short and only poorly conserved. However, homologues of genes encoding MFa-like pheromone-processing enzymes and pheromone transporters in S. cerevisiae, such as ram1, ram2, ste24, rce1 ste14, axl1 and ste6, are present in the genomes of filamentous ascomycetes, including members of the order Eurotiales (Po¨ggeler 2002; Po¨ggeler et al. 2008; Woo et al. 2006). These data suggest that in addition to the species listed in Fig. 5.5, many other filamentous ascomycetes may also have preserved the ability to secrete a prenylated MFa-like pheromone.

3. H-Type Pheromone Precursors – a Novel Class of Peptide Pheromone Precursors in Hypocreales Recently, Schmoll et al. (2010) reported the presence of a novel class of pheromone precursors in species in the order Hypocreales. The protein resembles an MFa-like pheromone precursor, which also shows characteristics of MFa-like pheromone precursors. Based on these characteristics, the new class of peptide pheromone precursors was named hybrid (h)-type pheromone precursors. The first gene encoding a h-type pheromone was identified in Trichoderma reesei (anamorph of Hypocrea jecorina) and was originally isolated as a gene reported to be associated with cellulose signal transduction in T. reesei (Schmoll et al. 2004). The gene termed h-type pheromone precursor gene 1 (hpp1), encodes the presumed MFa-like pheromone precursor of T. reesei. Htype pheromone precursors contain three copies of the motif [LI]GC[TS]VM, a CaaX motif at the C-terminus and two putative KEX2 processing sites. Similar to MFa-like pheromone precursors, h-type pheromone precursors are predicted to contain no signal sequence and are therefore suggested to be secreted via a non-classical STE6mediated secretion pathway. An extensive screen of available fungal genomes for homologues of the T. reesei h-type pheromone revealed that, thus far, h-type pheromones are encoded by only members of the order Hypocreales. They have been identified in Trichoderma virens, T. atroviride, Nectria haematococca, Fusarium verticilloides, and F. oxysporum, but not in Gibberella zeae or Epichloe festucae (Schmoll et al. 2010). The latter two encode typical MFa-like pheromone precursors in addition (Fig. 5.5). Thus far, all known members of the order Hypocreales encode typical MFa-like pheromone precursors (Table 5.1). In N. haematococca, in addition to the h-type pheromone gene, a small ORF coding for a polypeptide with a CaaX motif was identified near the cyanase locus (see Section III.A.2). However, this protein does not fit the MFa-pheromone consensus of filamentous ascomycetes shown in Fig. 5.5. In most cases, the h-type pheromone precursors seem to have replaced the MFa-like pheromone. Investigation of the maturation of this novel class of pheromone precursors will reveal whether their processing resembles MFa- or MFa-like pheromone precursors.

Function and Evolution of Pheromones and Pheromone Receptors in Filamentous Ascomycetes

85

B. Pheromone Receptors of Filamentous Ascomycetes

pre1 gene of N. crassa and related Neurospora species (Karlsson et al. 2008).

The finding of pheromone genes in filamentous ascomycetes implies that they also encode cognate pheromone receptors. The availability of genome sequences allowed the identification of pheromone receptor genes from many filamentous ascomycetes. A tblastn search of the N. crassa genome with the S. cerevisiae a-factor receptor STE2 and STE3 receptor sequences led to the first identification of pheromone receptors in filamentous ascomycetes. The genes encoding the putative receptors were named pre1 and pre2. The deduced pre1 gene product is a putative seven-TM protein with a high-level amino acid identity to the a-factor receptor STE3 of S. cerevisiae, and high similarity to lipopeptide pheromone receptors of basidiomycetes. The deduced pre2 product displays significant sequence similarity with the S. cerevisiae STE2 gene product, the MFa receptor. PRE1 and PRE2 are also predicted to have seven TM-spanning domains, an extracellular N-terminal tail, three outer and three cytoplasmic loops, and an inner C-terminal tail (Po¨ggeler and Ku¨ck 2001). A PCR-based assay with primers modeled on pre1 and pre2 sequences of N. crassa isolated the corresponding genes from the closely related homothallic S. macrospora. A heterologous yeast system showed that the S. macrospora receptor PRE2 facilitated all aspects of the S. cerevisiae pheromone response in S. cerevisiae MATa cells lacking the STE2 receptor, when activated by the S. macrospora peptide pheromone. Therefore, the conclusion is that the receptor encoded by the pre2 gene also functions as GPCR in S. macrospora (Mayrhofer and Po¨ggeler 2005).

Similar to the pheromone genes, pheromone receptor genes of the heterothallic N. crassa are predominantly expressed in a mating-type dependent manner, pre1 in MAT1-1 strains and pre2 in MAT1-2 strains (Karlsson et al. 2008; Kim and Borkovich 2004; Kim and Borkovich 2006). However, at reduced levels the transcripts of pre1 and pre2 could be identified in MAT1-2 and MAT1-1 strains, respectively (Kim and Borkovich 2004; Po¨ggeler and Ku¨ck 2001). Not surprisingly, both pheromone receptor genes are expressed constitutively during all stages of development in the homothallic S. macrospora (Po¨ggeler and Ku¨ck 2001). Interestingly, STE2- and STE3-like receptor genes and the MFa-like pheromone gene ppgA are expressed independently of mating-type in the heterothallic filamentous ascomycetes Neosartorya fumigata (anamorph: Aspergillus fumigatus) and the asexual filamentous fungi Penicillium chrysogenum (Hoff et al. 2008; Paoletti et al. 2005). In contrast to amino acid sequences of pheromones that are completely conserved between N. crassa and S. macrospora (Bobrowicz et al. 2002; Po¨ggeler and Ku¨ck 2000), pheromone receptor genes display extremely high sequence deviation when compared between the heterothallic N. crassa and the homothallic S. macrospora. Consequently, pheromone precursor genes are suggested to have evolved at a faster rate than genes encoding for metabolic enzymes (Po¨ggeler and Ku¨ck 2001). A detailed analysis of the molecular evolution of pheromone receptor genes, pre1 and pre2, in heterothallic members of the genus Neurospora confirmed this suggestion. The C-terminal region encoding the C-terminal intracellular loop domain of both PRE1 and PRE2 was demonstrated to constitute the rapidly evolving domain. In the C-terminal domain of PRE2 three highly variable regions encoding numerous repeats of glycine, threonine or aspartic acid make a decisive contribution to this variability. The Neurospora pre1 and pre2 genes evolve under low selective constrains, and pre1 contains additional sites under positive selection (Karlsson et al. 2008). Recently, Strandberg et al. (2010) found conflicting topologies between the genealogies of

In the first study, the pre1 ORF of S. macrospora was determined to have a size of 1232 bp, interrupted by one intron of 56 bp (Po¨ggeler and Ku¨ck 2001). RT-PCR analysis preformed in a later study revealed the presence of a second intron in the pre1 ORF. When the second intron is spliced from the pre1 transcript, 326 aa are added to the Nterminal 376 aa of the previously defined 391 aa PRE1 protein (Mayrhofer and Po¨ggeler 2005). The pre1 cDNA encoding the 391 aa polypeptide was amplified by RT-PCR using a primer within the second intron (Po¨ggeler and Ku¨ck 2001), indicating that the second intron is not removed in a portion of the pre1 transcripts. A second intron was also identified at a conserved position in the

86

Stefanie Po¨ggeler

pheromone receptor genes with the species phylogeny derived from non-coding sequences among heterothallic and pseudohomothallic taxa of Neurospora. The authors concluded that reproductive genes like pheromone receptor genes are more often introgressed between Neurospora species, than other parts of the genome. To date, both STE2- and STE3-like receptors have been identified in the genome of filamentous ascomycetes of many different orders and include heterothallic, homothallic and presumed asexual fungi. Many of these receptors have been demonstrated experimentally to function in mating and pheromone sensing (Table 5.2; see Section IV). Similar to their putative homologues in the ascomycetous yeast, the pheromone receptors PRE1 and PRE2 of filamentous ascomycetes have no obvious sequence similarity and belong to different classes of GPCRs (Xue et al. 2008). Therefore the pheromone receptor genes pre1 and pre2 probably did not arisen from a gene duplication event and seem to be orthologous genes. The length of pheromone receptors in filamentous ascomycetes differs from 294 to 577 aa in PRE2 homologues and from 300 to 724 aa in PRE2 homologues (Table 5.2). Some receptors in Table 5.2 were taken from genome sequencing projects and might contain annotation errors. The highest degree of sequence similarity among STE2- and STE3-like receptors of filamentous ascomycetes can be observed within the region that contains the TM helices, while the C-termini of the encoded proteins display a very high level of amino acid deviation. Mutational analysis of STE2 and STE3 from S. cerevisiae indicated that functional domains are organized in a manner similar to other GPCRs. TM segments are thought to form a pocket that acts as a ligand-binding domain, and the third intracellular loop is important for G protein activation. The cytoplasmic C-terminus is a target for a receptor kinase that negatively regulates receptor signaling and functions in ligand-induced endocytosis (Dohlman and Thorner 2001). Most likely, the latter function differs among different filamentous ascomycetes. The topology of phylogenetic trees obtained from maximum parsimony analyses of STE2- and STE3-like pheromone receptors of ascomycetous yeasts and filamentous ascomycetes appears to be consistent with the view derived from other systematic studies, and placed the taxa according to their predicted affiliation (Fig. 5.6).

IV. Function of Pheromone/Receptor Systems in Filamentous Ascomycetes Chemotropic interactions and diffusible attractants have long been known to be involved in the mating process of heterothallic filamentous ascomycetes, such as N. crassa (Bistis 1981, 1983), but recent research has demonstrated that pheromone signaling plays an important role in sexual reproduction (Bobrowicz et al. 2002; Coppin et al. 2005; Kim and Borkovich 2004, 2006; Kim et al. 2002). An important step in sexual reproduction of filamentous ascomycetes is bringing two compatible nuclei into the same cell and establishing a dikaryotic stage. Usually heterothallic ascomycetes send out a specialized hypha, termed a trychogyne, from a female prefruiting body that grows towards a fertilizing male cell of the opposite mating type. A functional male cell may be a uninucleate spermatiun, a microconidium or a multinucleate macroconidium or vegetative hyphae. In self-fertile, homothallic ascomycetes, spermatia and trychogynes are absent, conidia may be missing, and mating recognition does not seem to be necessary for sexual reproduction. This suggests that a pheromone/ receptor system is not needed for sensing a mating partner or initializing fertilization events. Nevertheless, pheromone genes and receptor genes are transcriptionally expressed in homothallic ascomycetes (Kim et al. 2008; Lee et al. 2008; Paoletti et al. 2007; Po¨ggeler 2000; Po¨ggeler and Ku¨ck 2001). Two functions of pheromones and pheromone receptors are important for filamentous ascomycetes. One is the regulation of the initial recognition between sexual structures of opposite mating types, which is essential for sexual reproduction in heterothallic filamentous ascomycetes. The other is possible function of the pheromone/receptor system in postfertilization events. This is equally important for homothallic and heterothallic fungi. Studies in the fission yeast Schizosaccharomyces pombe and basidiomycetes showed that pheromones can also be involved in meiosis, maintenance of the dikaryotic state, nuclear migration and clamp cell fusion (Casselton 2002; Spellig et al. 1994; Willer et al. 1995; see Chapter 6 in this volume; also see The Mycota vols I, XV).

Function and Evolution of Pheromones and Pheromone Receptors in Filamentous Ascomycetes

100 93

97 96 92 100

87 100

89

100 100

94

96 98 99

100

Fig. 5.6. Phylogenetic analysis of pheromone receptors from ascomycetes. (A) STE2-like receptors. (B) STE3-like receptors. Because of gaps and errors in the draft genome data, only the TM domains were used in this analysis. The TM domains were found to be common to all fungal STE2like and STE3-like pheromone receptors. The amino acid sequences of the TM domains 1–7 of STE2-like receptors and TM domains 2–7 of STE3-like receptors of a total of 51 receptors (Table 5.2) were aligned using the ClustalX program (Thompson et al. 2002). Phylogenetic analysis was

es s

Eu ro tia l ale es ial ar

rd s ale

Sa Sc cc m hiz Pne P ha yc os um ez Pl Helo ro et ac o iz P H eo m ale ch cy al sp tiale hyl ypo y ce s ar sti es or s loc cre ta o- da a ho al les les les ra es les

Ma

gn

ap

92

th

73

or

87 86 94 100

T. stipitatus N . sitophila N . crassa N . tetrasperma N . intermedia N . discreta S . macrospora P . anserina C . globosum M . grisea F . oxysporum F . verticilloides G . ze a e N . haematococca T . reesei T . atroviride V . albo-atrum B . fuckeliana S . sclerotiorum P . nodorum P . tritici-repentis A . gossypii K. lactis S . cerevisiae V . polyspora C . glabrata C . albicans P . stipitis T. melanosporum P. carinii S . pombe S . japonicus

So

100

Eu ro t

ial

100

en

100 100

A . terreus A . niger N . fischeri A . fumigatus A . c lavatus P . chrysogenum A. nidulans T . verrucosum A . benhamiae N . otae C . posadasii U . reesii A . capsulata A . dermatitidis P . brasiliensis P . marneffei

yg

100 100

A . oryzae A. flavus

On

100

es

A

87

made with programs contained in the program package PHYLIP ver.3.6 (Felsenstein 2004). Program PROTPARS was used for constructing phylogenetic trees. Statistical significance was evaluated by bootstrap analysis, with 1000 iterations of bootstrap samplings and reconstructing of trees by PROTPARS. A majority rule consensus tree was subsequently generated using the program CONSENSE. S. pombe and S. japonicus were used as outgroups. Bootstrap values greater than 70% are given (in percentages)

100

98

s

Eu ro tia l m s ale

tid

m

93

100 100 100

97 99 92

95

99

89

100 100

ial

Sc H M Ph m hizo elo ag yllo Hy yc t n po et sac iale apo cho cr ale ch s rth ral ea ar s e les a s oles

So

rd

89 100

ar

82

es

Pn

eu

97

oc

Sa

100

ys

cc

ha

100

ro

72

yc

Pl

100

eo

99

Eu ro t

72

sp Pez or iza et ale le ale s s s

ial

100 99

ale

82 100

en

99

yg

100

A . fumigatus N . fischeri A . c lavatus P . chrysogenum A. flavus A . oryzae A . terreus A . niger A. nidulans A . dermatitidis A . capsulata P . brasiliensis U . reesii C . posadasii A . benhamiae T . verrucosum N . otae P . marneffei T. stipitatus T. melanosporum P . nodorum P . tritici-repentis S . cerevisiae V . polyspora C . glabrata K. lactis A . gossypii P . stipitis C . albicans P . carinii N . tetrasperma N . sitophila N . intermedia N . crassa N . discreta S . macrospora P . anserina C . globosum F . verticilloides F . oxysporum G . ze a e N . haematococca T . atroviride T . reesei V . albo-atrum M . grisea S . sclerotiorum B . fuckeliana S . pombe S . japonicus

On

100 92

es

B

es

Stefanie Po¨ggeler

88

Fig. 5.6 (continued) An essential postfertilization event is the development of dikaryotic hyphae, which is a prerequisite for karyogamy and meiosis in filamentous ascomycetes. Recognition between nuclei has been suggested to be mediated by nuclear limited expression of mating type-specific pheromones and receptors. In addition, pheromones and receptors are thought to be limited to the plasma membrane region close to the individual nucleus. The spatial restric-

tion of signaling components is proposed to facilitate recognition between two nuclei (Debuchy 1999; Schuurs et al. 1998; Shiu and Glass 2000).

To examine functions of pheromone and pheromone receptors and analyze their impact during fertilization, fruiting body and ascospore

Function and Evolution of Pheromones and Pheromone Receptors in Filamentous Ascomycetes

development, the relevant genes have been deleted and the phenotypes of the mutants have been studied in detail in various heterothallic and homothallic filamentous ascomycetes. A. Functional Analysis of Pheromones and Pheromone Receptors in Heterothallic Filamentous Ascomycetes In heterothallic filamentous ascomycetes, male and female fertility were found to dependent on specific interaction of pheromones with their cognate receptors. When pheromone genes were deleted, male spermatia or macroconidia were no longer able to fertilize female partners. Pheromone deletion mutants are male sterile but do not generally display defects in vegetative development and female fertility. Thus, the main function of pheromones in heterothallic filamentous ascomycetes is to ensure the fertility of the male partner (Coppin et al. 2005; Kim and Borkovich 2006; Kim et al. 2002; Turina et al. 2003). In N. crassa, heterologous expression of a pheromone gene in male cells can direct chemotropic polarized growth of trychogynes from an otherwise incompatible mating-type background (Kim and Borkovich 2006). In contrast to the inability of conidia to fuse with the trychogyne of the opposite mating type, N. crassa pheromone-deleted strains can fuse with neighboring conidia and vegetative hyphae, indicating that pheromones are not required for vegetative cell fusion (Kim and Borkovich 2006).

Additional functions of pheromones beyond their role in fertilization have been reported for MFa-like pheromones in C. parasitica and N. crassa (Kim et al. 2002; Zhang et al. 1993). Deletion of the Mf2/2 gene, one of the two copies of the MFa-like pheromone genes, in C. parasitica results in a pleiotropic phenotype. When Mf2/ 2 deletion strains are used as the female partner, the perithecia that formed are barren; in rare cases they contain shrunken and deformed asci but these do not contain ascospores. Moreover, the production of asexual spores is reduced, whereas the Mf2/2 deletion strain is fully competent in sexual crosses when used as male partner (Turina et al. 2003; Zhang et al. 1993). This may be explained by the second MFa-like pheromone gene, Mf2/1, in the Mf2/2 deletion strain. Simi-

89

larly, deletion of three of the seven putative pheromone peptide repeats within the MFa-factor-like Mf1/1 gene has no effect on mating. Thus, the gene dosage effect does not appear to be essential for male fertility under laboratory conditions. The role of the C. parasitica Mf2/2 gene in both asexual and sexual sporulation is currently not understood, but the Mf2/2 gene encodes four small ORFs, only one of which corresponds to the MFa-like pheromone precursor. The role of other peptides translated from the Mf2/2 transcript is not known but these may regulate asexual and sexual development (Turina et al. 2003; Zhang et al. 1993). As in C. parasitica, mutations generated by repeat-induced point (RIP) mutation in the N. crassa MFa-like pheromone precursor gene ppg2 result in pleiotropic phenotypes. Mutants carrying point mutations in both the ppg2 ORF and the 3’ untranslated region (UTR), display male sterility in MAT1-2 strains because they are unable to attract MAT1-1 trychogynes and possess additional defects in female sexual development, ascospore production and vegetative growth (Kim et al. 2002). Interestingly, deletion mutants of N. crassa lacking the complete ppg2 ORF do not display obvious defects in vegetative development and female fertility, suggesting that the pleitropic defects of the ppg2 RIP mutants result from mutations outside the ppg2 ORF (Kim and Borkovich 2006). In P. anserina, disruption of the 3’UTR mfp (a homologue of the N. crassa ppg2 and the C. parasitica Mf2/2) results in reduced fertility. Thus, the 3’UTR of MFa-like pheromone transcripts may mediate functions other than initial fertilization events. Due to the block in trychogyne attraction and cell fusion exhibited by the pheromone gene deletion mutants functions in postfertilization events could not be analyzed in N. crassa (Kim and Borkovich 2006). However, Coppin et al. (2005) managed to cross pheromone double-deletion strains of opposite mating types in P. anserina by complementation and transient expression of the pheromone gene. They showed that these crosses were fertile and demonstrated that, once fertilization occurs, pheromones are unnecessary for the complementation of the sexual cycle. Deletions of pheromone receptor genes have been analyzed only in the heterothallic N. crassa. The STE3-like pheromone receptor gene pre1 was shown to be essential for mating type-specific directional growth and fusion of trychogynes

90

Stefanie Po¨ggeler

and thus for female fertility. Analysis of pre1 deletion strains revealed that pre1 does not affect vegetative growth, asexual sporulation and male fertility of either mating type. Furthermore, heterokaryon formation is not affected. However, prefruiting bodies from MAT1-1 strains do not undergo fertilization. This defect results from the inability of the MAT1-1 pre1 deletion strains to recognize and fuse with MAT1-2 male cells. When examined microscopically, trychogynes of MAT1-1 pre1 deletion strains do not display directional growth toward, and do not coil around the male cells of opposite mating type (Kim and Borkovich 2004). Interestingly, loss of the PRE1 receptor leads to significantly reduced expression of both pheromone genes in the appropriate mating type. Kim and Borkovich (2004) therefore suggested that the low level of pre1 expression in MAT1-2 cells may be involved in regulating pheromone expression. They further suggested that the regulation could result from a pheromoneindependent signaling activity of PRE1 leading to activation of a common transcription factor that affects the expression of both pheromone genes. Recently, the pheromone precursor gene hpp1 was found to encode a novel class of peptide pheromone precursor (see Section III.A.3) essential for male fertility in T. reessei. Interestingly, the expression of hhp1 and the MFa-like pheromone gene ppg1 is not dependent on the mating-type in the supposedly heterothallic T. reesei (Schmoll et al. 2010).

Sequencing of the complete genome of the supposedly asexual human pathogen A. fumigatus led to the identification of genes, as well as pheromone and receptor genes and a number of other genes involved in the sexual cycle of perfect fungi (Paoletti et al. 2005; Po¨ggeler 2002). Mating-type genes and pheromone, receptor and other sexrelated genes were found to be expressed in A. fumigatus (Paoletti et al. 2005). While pheromone-receptor genes preA and preB are expressed with no clear differences between MAT1-1 and MAT1-2 strains, the expression level of the MFa gene ppgA is higher in a MAT1-1 than in a MAT12 isolate. Recently, pairwise crossing of MAT1-1 and MAT1-2 strains, showed that sexual reproduction of A. fumigatus can be induced in vitro after six months of incubation on parafilm-sealed oatmeal agar plates. Thus, A. fumigatus is a heterothallic ascomycete and was named Neosartorya fumigata (O’Gorman et al. 2009). No differ-

ence in expression of pheromone and receptor genes was detected in sealed versus unsealed plates of A. fumigatus (Paoletti et al. 2005) and the effect of pheromone and pheromone receptors in the sexual reproduction of A. fumigatus has not yet been analyzed.

B. Functional Analysis of Pheromones and Pheromone Receptors in Homothallic Filamentous Ascomycetes Unlike heterothallic filamentous ascomycetes, the role of pheromones and pheromone receptors has been questioned in homothallic filamentous ascomycetes (see above). However, the presence of transcripts for pheromone and pheromone receptor genes in homothallic filamentous ascomycetes such as S. macrospora, A. nidulans and G. zeae suggest that pheromone receptor systems function beyond their role in fertilization (Kim et al. 2008; Lee et al. 2008; Paoletti et al. 2007; Po¨ggeler 2000; Po¨ggeler and Ku¨ck 2001; Seo et al. 2004). Pheromone receptor genes identified in homothallic ascomycetes are not expressed in a matingtype-dependent manner (Kim et al. 2008; Klix et al. 2010; Paoletti et al. 2007; Po¨ggeler et al. 2006). They are expressed under vegetative growth conditions and during the entire sexual development in S. macrospora, and are upregulated under conditions favoring sexual development in A. nidulans (Paoletti et al. 2007; Po¨ggeler 2000; Po¨ggeler and Ku¨ck 2001). Similar to A. nidulans, the G. zeae MFa receptor gene pre2 is expressed in all cells under conditions favoring sexual development, but the pre1 gene is expressed weakly and exclusively in mature ascospores (Lee et al. 2008). In contrast to the pheromone receptor genes, the pheromone precursor genes of homothallic filamentous ascomycetes are regulated by the mating-type locus (Kim et al. 2008; Klix et al. 2010; Paoletti et al. 2007; Po¨ggeler et al. 2006). The S. macrospora DSmta-1 mutant that lacks the MAT1-2-specific HMG transcription factor SMTa-1 displays a 500-fold reduction of lipopeptide ppg2 gene expression, while ppg1, the other pheromone precursor, is not affected. This indicates that MAT1-2 has a direct or indirect impact on the activation of ppg2 gene expression (Po¨ggeler et al. 2006). Both pheromone genes of S. macrospora are significantly downregulated in the DSmtA-1 mutant which lacks the a-domain gene

Function and Evolution of Pheromones and Pheromone Receptors in Filamentous Ascomycetes

MAT1-1-1. Only ppg2 is significantly upregulated in the DSmtA-2 mutant lacking the Sordarialesspecific MAT1-1-2 gene (Klix et al. 2010). The GzPPG1 transcript in the homothallic G. zeae cannot be detected in strains deleted for the MAT1-1-specific or the MAT1-2-specific genes. However, the GzPPG2 transcript level increases in a G. zeae DMAT1-1 strain (Kim et al. 2008). In the homothallic A. nidulans, deletion of MAT1-11 gene has no effect on the expression of the pheromone gene ppgA while its expression increases in a DMAT1-2 mutant (Paoletti et al. 2007). To investigate the function of pheromones and receptor in homothallic filamentous ascomycetes, single, double- and even quadruple-mutant strains have been generated (Kim et al. 2008; Lee et al. 2008; Mayrhofer and Po¨ggeler 2005; Mayrhofer et al. 2006; Seo et al. 2004). In S. macrospora, deletion of any single pheromone or receptor gene has no effect on vegetative growth or sexual development, but double-deletion strains lacking any compatible pheromone receptor pair (Dpre1/ Dppg1 and Dpre2/Dppg2) and the double pheromone mutant (Dppg1/Dppg2) display a drastically reduced number of fruiting bodies and sexual spores. Deletion of both receptor genes (Dpre1/D pre2) completely eliminates fruiting body and ascospore formation (Mayrhofer and Po¨ggeler 2005; Mayrhofer et al. 2006). In cases in which perithecia are produced in double-deletion strains, ascospore production is extremely impaired and only very few croziers are formed (Mayrhofer et al. 2006). This suggests that, in S. macrospora, at least one pheromone/receptor system is necessary for proper formation of dikaryotic ascogenous hyphae. In filamentous ascomycetes, the nuclear sorting that occurs in the dikaryotic ascogenous hyphae is reminiscent of the synchronous division of two nuclei and clamp cell formation in the dikaryotic hyphae of basidiomycetes (Casselton 2002). In basidiomycetes the pheromone/receptor system is required to promote nuclear migration and clamp cell fusion after the dikaryon is established (O’Shea et al. 1998; Vaillancourt et al. 1997; Wendland et al. 1995). Pheromones secreted by the clamp cells are believed to activate receptors on the surface of the subterminal cell and vice versa (Brown and Casselton 2001; see Chapter 6 in this volume). Based on the basidiomycete system, the pheromone/receptor system of filamentous ascomycetes was proposed to be possibly involved in nuclear migration and crozier formation in ascogenous hyphae (Mayrhofer et al. 2006).

91

Similar to S. macrospora, a double deletion of two receptor genes in A. nidulans (DpreA/DpreB) completely abolishes fruiting body and ascospore formation (Seo et al. 2004). Single pheromone receptor mutants produce smaller cleistothecia with a reduced number of ascospores. This result also suggests a postfertilization role for pheromone receptors in A. nidulans. In contrast, outcrossing of A. nidulans receptor mutants (DpreA/DpreB
DpreA/DpreB) results in fruiting body and ascospore formation at wild-type level, indicating that in A. nidulans pheromone receptors are required only for self-fertilization and not for sexual development per se (Seo et al. 2004). In the homothallic G. zeae, pheromone and receptor mutants from single- to quadrupledeletion strains are similar to the self-fertile progenitor in both male and female fertility and in vegetative growth. However, deletion combinations that include either the ppg1 pheromone gene or the pre2 receptor gene cause increased numbers of immature perithecia (Kim et al. 2008). The Dppg1 and the Dpre2 mutant displays approximately 50% reduced fertility in self-fertilization tests. Moreover, deletion of ppg1 reduced male fertility and deletion of pre2 reduced female fertility in outcrossing tests (Lee et al. 2008). Similar to S. macrospora and A. nidulans, G. zeae produces no obvious trychogyne, but its macroconidia can be forced to act as male partners (Lee et al. 2003). Thus, it is possible to test effects of pheromone and receptor gene deletions on the male and female fertility in this homothallic fungus.

Evidence shows that one of the putative pheromone/receptor pairs of G. zeae (ppg1/pre2) enhances, but is not essential for selfing and outcrossing, whereas no functional role was identified for the other pair (Lee et al. 2008). However, even the Dpgg1/Dpre2/Dppg2/Dpre1quadruple mutant generated by Kim et al. (2008) produces fruiting bodies and ascospores, so the putative pheromone/receptor system seems to play a nonessential role in the sexual development of G. zeae. As an adaption mechanism to the homothallic lifestyle many variations have occurred not only in the regulation of the expression but also in the functions of pheromone and pheromone receptor genes. Pheromone/receptor systems of homothallic filamentous seem to be more important at later stages of the sexual development than for mate recognition as commonly seen in their heterothallic relatives.

92

Stefanie Po¨ggeler

V. Conclusions Genomic mining of fungal genomes revealed pheromone and pheromone receptor genes in heterothallic, homothallic and supposedly asexual filamentous ascomycetes. The encoded proteins resemble the MFa and MFa pepetide precursor and the STE2 and STE3 pheromone receptors of the ascomycetous yeast S. cerevisiae. Recently a novel type of pheromone precursor genes was identified in the Hypocreales that is assumed to act as a MFa-like pheromone in some species of this order. Similar to S. cerevisiae, the main function of the pheromone/receptor system in heterothallic filamentous ascomycetes is initialization of the fertilization event, while pheromone/receptor systems of homothallic ascomycetes seem to promote postfertilization events. Future studies on the structure and functions of pheromone/receptor systems in heterothallic as well as homothallic filamentous ascomycetes will reveal new insights into the regulation of sexual development of filamentous ascomycetes and the evolution of fungal mating strategies.

References Bader O, Krauke Y, Hube B (2008) Processing of predicted substrates of fungal Kex2 proteinases from Candida albicans, C. glabrata, Saccharomyces cerevisiae and Pichia pastoris. BMC Microbiol 8:116 Bardwell L (2005) A walk-through of the yeast mating pheromone response pathway. Peptides 26:339–350 Bell-Pedersen D, Shinohara ML, Loros JJ, Dunlap JC (1996) Circadian clock-controlled genes from Neurospora crassa are late night- to early morning-specific. Proc Natl Acad Sci USA 93:13096–13101 Betz R, MacKay VL, Duntze W (1977) a-Factor from Saccharomyces cerevisiae: partial characterization of a mating hormone produced by cells of mating type a. J Bacteriol 132:462–472 Bistis GN (1981) Chemotropic interactions between trychogynes and conidia of the opposite mating type in Neurospora crassa. Mycologia 73:959–975 Bistis GN (1983) Evidence for diffusible, mating-type specifictrichogyne attractants in Neurospora crassa. Exp Mycol 7:292–295 Blakelee AF (1904) Sexual reproduction in the Mucorinae. Proc Am Acad Arts Sci 40:205–319 Bobrowicz P, Pawlak R, Correa A, Bell-Pedersen D, Ebbole DJ (2002) The Neurospora crassa pheromone precursor genes are regulated by the mating type locus and the circadian clock. Mol Microbiol 45:795–804

Bo¨lker M, Urban M, Kahmann R (1992) The a mating type locus of U. maydis specifies cell signaling components. Cell 68:441–450 Brake AJ, Brenner C, Najarian R, Laybourne P (1985) Structure of genes encoding precursors of the yeast peptide mating pheromone a-factor. In: Gething MJ (ed) Protein transport and secretion. Cold Spring Habor Laboratory Press, Cold Spring Habor, NY, pp 103–108 Brake AJ, Julius DJ, Thorner J (1983) A functional preproalpha-factor gene in Saccharomyces yeasts can contain three, four, or five repeats of the mature pheromone sequence. Mol Cell Biol 3:1440–1450 Brenner C, Fuller RS (1992) Structural and enzymatic characterization of a purified prohormone-processing enzyme: secreted, soluble Kex2 protease. Proc Natl Acad Sci USA 89:922–926 Brown AJ, Casselton L (2001) Mating in mushrooms: increasing the chances but prolonging the affair. Trends Genet 17:393–400 Bu¨cking-Throm E, Duntze W, Hartwell LH, Manney TR (1973) Reversible arrest of haploid yeast cells in the initiation of DNA synthesis by a diffusible sex factor. Exp Cell Res 76:99–110 Burkholder AC, Hartwell LH (1985) The yeast alpha-factor receptor: structural properties deduced from the sequence of the STE2 gene. Nucleic Acids Res 13: 8463–8475 Caldwell GA, Naider F, Becker JM (1995) Fungal lipopeptide mating pheromones: a model system for the study of protein prenylation. Microbiol Rev 59: 406–422 Casselton LA (2002) Mate recognition in fungi. Heredity 88:142–147 ´ elic´ A, Martin NP, Son CD, Becker JM, Naider F, Dumont C ME (2003) Sequences in the intracellular loops of the yeast pheromone receptor ste2p required for G protein activation. Biochemistry 42:3004–3017 Choi Y, Konopka JB (2006) Accessibility of cysteine residues substituted into the cytoplasmic regions of the a-factor receptor identifies the intracellular residues that are available for G protein interaction. Biochemistry 45:15310–15317 Ciejek E, Thorner J, Geier M (1977) Solid phase peptide synthesis of a-factor, a yeast mating pheromone. Biochem Biophys Res Commun 78:952–961 Cooper A, Bussey H (1989) Characterization of the yeast KEX1 gene product: a carboxypeptidase involved in processing secreted precursor proteins. Mol Cell Biol 9:2706–2714 Coppin E, de Renty C, Debuchy R (2005) The function of the coding sequences for the putative pheromone precursors in Podospora anserina is restricted to fertilization. Eukaryotic Cell 4:407–420 Davey J (1992) Mating pheromones of the fission yeast Schizosaccharomyces pombe: purification and structural characterization of M-factor and isolation and analysis of two genes encoding the pheromone. EMBO J 11:951–960 Debuchy R (1999) Internuclear recognition: a possible connection between euascomycetes and homobasidiomycetes. Fungal Genet Biol 27:218–223

Function and Evolution of Pheromones and Pheromone Receptors in Filamentous Ascomycetes Debuchy R, Berteaux-Leceleir V, Silar P (2010) Mating systems and sexual morphogenesis in ascomycetes. In: Borkovich KA, Ebbole DJ (eds) Cellular and molecular biology of filamentous fungi. ASM Press, Washington, DC, pp 501–535 Debuchy R, Turgeon BG (2006) Mating-type structure, evolution, and function in euascomycetes. In: Ku¨es U, Fischer R (eds) Growth, differentiation and sexuality (The Mycota). Springer, Berlin Heidelberg New York, pp 293–323 Dignard D, El-Naggar AL, Logue ME, Butler G, Whiteway M (2007) Identification and characterization of MFA1, the gene encoding Candida albicans a-factor pheromone. Eukaryot Cell 6:487–494 Dohlman HG, Slessareva JE (2006) Pheromone signaling pathways in yeast. Sci STKE 2006:cm6 Dohlman HG, Thorner JW (2001) Regulation of G proteininitiated signal transduction in yeast: paradigms and principles. Annu Rev Biochem 70:703–754 Dolan JW, Fields S (1991) Cell-type-specific transcription in yeast. Biochim Biophys Acta 1088:155–169 Duntze W, MacKay V, Manney TR (1970) Saccharomyces cerevisiae: a diffusible sex factor. Science 168: 1472–1473 Elion EA, Qi M, Chen W (2005) Signal transduction: signaling specificity in yeast. Science 307:687–688 Elleuche S, Po¨ggeler S (2008) A cyanase is transcriptionally regulated by arginine and involved in cyanate decomposition in Sordaria macrospora. Fungal Genet Biol 45: 1458–1469 Felsenstein J (2004) PHYLIP. Phylogeny inference package version 3.6. Distributed by the author. University of Washington, Seattle Galagan JE, Calvo SE, Cuomo C, Ma LJ, Wortman JR, Batzoglou S, Lee SI, Bastu¨rkmen M, Spevak CC, Clutterbuck J, Kapitonov V, Jurka J, Scazzocchio C, Farman M, Butler J, Purcell S, Harris S, Braus GH, Draht O, Busch S, D’Enfert C, Bouchier C, Goldman GH, Bell-Pedersen D, Griffiths-Jones S, Doonan JH, Yu J, Vienken K, Pain A, Freitag M, Selker EU, Archer DB, Pen˜alva MA, Oakley BR, Momany M, Tanaka T, Kumagai T, Asai K, Machida M, Nierman WC, Denning DW, Caddick M, Hynes M, Paoletti M, Fischer R, Miller B, Dyer P, Sachs MS, Osmani SA, Birren BW (2005) Sequencing of Aspergillus nidulans and comparative analysis with A. fumigatus and A. oryzae. Nature 438:1105–1115 Glass NL, Vollmer SJ, Staben C, Grotelueschen J, Metzenberg RL, Yanofsky C (1988) DNAs of the two mating-type alleles of Neurospora crassa are highly dissimilar. Science 241:570–573 Hagen DC, McCaffrey G, Sprague GF (1986) Evidence the yeast STE3 gene encodes a receptor for the peptide pheromone a factor: gene sequence and implications for the structure of the presumed receptor. Proc Natl Acad Sci USA 83:1418–1422 Hamm HE (1998) The many faces of G protein signaling. J Biol Chem 273:669–672 Hartwell L (1980) Mutants of Saccharomyces cerevisiae unresponsive to cell division control by polypeptide mating hormone. J Cell Biol 85:811–822

93

Herskowitz I (1989) A regulatory hierarchy for cell specialization in yeast. Nature 342:749–757 Hicks KA, Hartman HL, Fierke CA (2005) Upstream polybasic region in peptides enhances dual specificity for prenylation by both farnesyltransferase and geranylgeranyltransferase type I. Biochemistry 44: 15325–15333 Hoff B, Po¨ggeler S, Ku¨ck U (2008) Eighty years after its discovery, Fleming’s Penicillium strain discloses the secret of its sex. Eukaryot Cell 7:465–470 Hougland JL, Lamphear CL, Scott SA, Gibbs RA, Fierke CA (2009) Context-dependent substrate recognition by protein farnesyltransferase. Biochemistry 48: 1691–1701 Hrycyna CA, Sapperstein SK, Clarke S, Michaelis S (1991) The Saccharomyces cerevisiae STE14 gene encodes a methyltransferase that mediates C-terminal methylation of a-factor and RAS proteins. EMBO J 10: 1699–16709 Huyer G, Kistler A, Nouvet FJ, George CM, Boyle ML, Michaelis S (2006) Saccharomyces cerevisiae a-factor mutants reveal residues critical for processing, activity, and export. Eukaryot Cell 5:1560–1570 Imai Y, Yamamoto M (1994) The fission yeast mating pheromone P-factor: its molecular structure, gene structure, and ability to induce gene expression and G1 arrest in the mating partner. Genes Dev 8:328–338 Jime´nez R, Burgos M (1998) Mammalian sex determination: joining pieces of the genetic puzzle. Bioassays 20:696–699 Julius D, Blair L, Brake A, Sprague G, Thorner J (1983) Yeast alpha factor is processed from a larger precursor polypeptide: the essential role of a membranebound dipeptidyl aminopeptidase. Cell 32:839–852 Julius D, Brake A, Blair L, Kunisawa R, Thorner J (1984) Isolation of the putative structural gene for the lysinearginine-cleaving endopeptidase required the processing of yeast prepro-a-factor. Cell 36:309–318 Karlson P, Luscher M (1959) Pheromones: a new term for a class of biologically active substances. Nature 183: 55–56 Karlsson M, Nygren K, Johannesson H (2008) The evolution of the pheromonal signal system and its potential role for reproductive isolation in heterothallic neurospora. Mol Biol Evol 25:168–178 Kim H-K, Lee T, Yun S-H (2008) A putative pheromone signaling pathway is dispensable for self-fertility in the homothallic ascomycete Gibberella zeae. Fungal Genet Biol 45:1188–1196 Kim H, Borkovich KA (2004) A pheromone receptor gene, pre1, is essential for mating type-specific directional growth and fusion of trichgynes and female fertility in Neurospora crassa. Mol Microbiol 52:1781–1798 Kim H, Borkovich KA (2006) Pheromones are essential for male fertility and sufficient to direct chemotropic polarized growth of trichogynes during mating in Neurospora crassa. Eukaryotic Cell 5:544–554 Kim H, Metzenberg RL, Nelson MA (2002) Multiple functions of mfa-1, a putative pheromone precursor gene of Neurospora crassa. Eukaryotic Cell 1:987–999 Klix V, Nowrousian M, Ringelberg C, Loros JJ, Dunlap JC, Poggeler S (2010) Functional characterization of

94

Stefanie Po¨ggeler

MAT1-1-specific mating-type genes in the homothallic ascomycete Sordaria macrospora provides new insights into essential and nonessential sexual regulators. Eukaryot Cell 9:894–905 Koopman P (1999) Sry and Sox9: mammalian testisdetermining genes. Cell Mol Life Sci 55:839–885 Korf RP (1952) The terms homothallism and heterothallism. Nature 170:534–535 Kurjan J (1993) The pheromone response pathway in Saccharomyces cerevisiae. Annu Rev Genet 27:147–179 Kurjan J, Herskowitz I (1982) Structure of a yeast pheromone gene (MFa): a putative a-factor precursor contains four tandem copies of mature a-factor. Cell 30:933–943 Lee J, Lee T, Lee YW, Yun SH, Turgeon BG (2003) Shifting fungal reproductive mode by manipulation of mating type genes: obligatory heterothallism of Gibberella zeae. Mol Microbiol 50:145–152 Lee J, Leslie JF, Bowden RL (2008) Expression and function of sex pheromones and receptors in the homothallic ascomycete Gibberella zeae. Eukaryotic Cell 7: 1211–1221 Levi JD (1956) Mating reaction in yeast. Nature 177: 753–754 Lin X, Heitman J (2007) Mechanisms of homothallism in Fungi and transitions between heterothallism and homothallism. In: Heitman J, Kronstad JW, Taylor JW, Casselton LA (eds) Sex in fungi. ASM Press, Washington, DC, pp 35–57 Machida M, Asai K, Sano M, Tanaka T, Kumagai T, Terai G, Kusumoto K-I, Arima T, Akita O, Kashiwagi Y, Abe K, Gomi K, Horiuchi H, Kitamoto K, Kobayashi T, Takeuchi M, Denning DW, Galagan JE, Nierman WC, Yu J, Archer DB, Bennett JW, Bhatnagar D, Cleveland TE, Fedorova ND, Gotoh O, Horikawa H, Hosoyama A, Ichinomiya M, Igarashi R, Iwashita K, Juvvadi PR, Kato M, Kato Y, Kin T, Kokubun A, Maeda H, Maeyama N, Maruyama J-i, Nagasaki H, Nakajima T, Oda K, Okada K, Paulsen I, Sakamoto K, Sawano T, Takahashi M, Takase K, Terabayashi Y, Wortman JR, Yamada O, Yamagata Y, Anazawa H, Hata Y, Koide Y, Komori T, Koyama Y, Minetoki T, Suharnan S, Tanaka A, Isono K, Kuhara S, Ogasawara N, Kikuchi H (2005) Genome sequencing and analysis of Aspergillus oryzae. Nature 438:1157–1161 Marcus S, Caldwell GA, Miller D, Xue CB, Naider F, Becker JM (1991) Significance of C-terminal cysteine modifications to the biological activity of the Saccharomyces cerevisiae a-factor mating pheromone. Mol Cell Biol 11:3603–3612 Maurer-Stroh S, Eisenhaber F (2005) Refinement and prediction of protein prenylation motifs. Genome Biol 6:R55 Mayrhofer S, Po¨ggeler S (2005) Functional characterization of an a-factor-like Sordaria macrospora peptide pheromone and analysis of its interaction with its cognate receptor in Saccharomyces cerevisiae. Eukaryot Cell 4:661–672 Mayrhofer S, Weber JM, Po¨ggeler S (2006) Pheromones and pheromone receptors are required for proper sexual development in the homothallic ascomycete Sordaria macrospora. Genetics 172:1521–1533

Metzenberg RL, Glass NL (1990) Mating type and mating strategies in Neurospora. Bioessays 12:53–59 Michaelis S (1993) STE6, the yeast a-factor transporter. Semin Cell Biol 4:17–27 Moores SL, Schaber MD, Mosser SD, Rands E, O’Hara MB, Garsky VM, Marshall MS, Pompliano DL, Gibbs JB (1991) Sequence dependence of protein isoprenylation. J Biol Chem 266:14603–14610 Naider F, Becker JM (2004) The [alpha]-factor mating pheromone of Saccharomyces cerevisiae: a model for studying the interaction of peptide hormones and G protein-coupled receptors. Peptides 25: 1441–1463 O’Gorman CM, Fuller HT, Dyer PS (2009) Discovery of a sexual cycle in the opportunistic fungal pathogen Aspergillus fumigatus. Nature 457:471–474 O’Shea SF, Chaure PT, Halsall JR, Olesnicky NS, Leibbrandt A, Connerton IF, Casselton LA (1998) A large pheromone and receptor gene complex determines multiple B mating type specificities in Coprinus cinereus. Genetics 148:1081–1090 Paoletti M, Rydholm C, Schwier EU, Anderson MJ, Szakacs G, Lutzoni F, Debeaupuis JP, Latge´ JP, Denning DW, Dyer PS (2005) Evidence for sexuality in the opportunistic fungal pathogen Aspergillus fumigatus. Curr Biol 15:1242–1248 Paoletti M, Seymour FA, Alcocer MJ, Kaur N, Calvo AM, Archer DB, Dyer PS (2007) Mating type and the genetic basis of self-fertility in the model fungus Aspergillus nidulans. Curr Biol 17:1384–1389 Pel HJ, de Winde JH, Archer DB, Dyer PS, Hofmann G, Schaap PJ, Turner G, de Vries RP, Albang R, Albermann K, Andersen MR, Bendtsen JD, Benen JA, van den Berg M, Breestraat S, Caddick MX, Contreras R, Cornell M, Coutinho PM, Danchin EG, Debets AJ, Dekker P, van Dijck PW, van Dijk A, Dijkhuizen L, Driessen AJ, d’Enfert C, Geysens S, Goosen C, Groot GS, de Groot PW, Guillemette T, Henrissat B, Herweijer M, van den Hombergh JP, van den Hondel CA, van der Heijden RT, van der Kaaij RM, Klis FM, Kools HJ, Kubicek CP, van Kuyk PA, Lauber J, Lu X, van der Maarel MJ, Meulenberg R, Menke H, Mortimer MA, Nielsen J, Oliver SG, Olsthoorn M, Pal K, van Peij NN, Ram. A.F, Rinas U, Roubos JA, Sagt CM, Schmoll M, Sun J, Ussery D, Varga J, Vervecken W, van de Vondervoort PJ, Wedler H, Wo¨sten HA, Zeng AP, van Ooyen AJ, Visser J, Stam H (2007) Genome sequencing and analysis of the versatile cell factory Aspergillus niger CBS 513.88. Nat Biotechnol 25:221–231 Pierce KL, Premont RT, Lefkowitz RJ (2002) Seven-transmembrane receptors. Nat Rev Mol Cell Biol 3:639–650 Po¨ggeler S (2000) Two pheromone precursor genes are transcriptionally expressed in the homothallic ascomycete Sordaria macrospora. Curr Genet 37:403–411 Po¨ggeler S (2002) Genomic evidence for mating abilities in the asexual pathogen Aspergillus fumigatus. Curr Genet 42:153–160 Po¨ggeler S, Hoff B, Ku¨ck U (2008) Asexual cephalosporin C producer Acremonium chrysogenum carries a functional mating type locus. Appl Environ Microbiol 74:6006–6016

Function and Evolution of Pheromones and Pheromone Receptors in Filamentous Ascomycetes Po¨ggeler S, Ku¨ck U (2000) Comparative analysis of the mating-type loci from Neurospora crassa and Sordaria macrospora: identification of novel transcribed ORFs. Mol Gen Genet 263:292–301 Po¨ggeler S, Ku¨ck U (2001) Identification of transcriptionally expressed pheromone receptor genes in filamentous ascomycetes. Gene 280:9–17 Po¨ggeler S, Nowrousian M, Ringelberg C, Loros JJ, Dunlap JC, Ku¨ck U (2006) Microarray and real-time PCR analyses reveal mating type-dependent gene expression in a homothallic fungus. Mol Gen Genomics 275:492–503 Po¨ggeler S, Risch S, Ku¨ck U, Osiewacz HD (1997) Matingtype genes from the homothallic fungus Sordaria macrospora are functionally expressed in a heterothallic ascomycete. Genetics 147:567–580 Powell WA, Van Alfen NK (1987) Differential accumulation of poly(A)þ RNA between virulent and double-stranded RNA-induced hypovirulent strains of Cryphonectria (Endothia) parasitica. Mol Cell Biol 7:3688–3693 Powers S, Michaelis S, Broek D, Sonia Santa A-A, Field J, Herskowitz I, Wigler M (1986) RAM, a gene of yeast required for a functional modification of RAS proteins and for production of mating pheromone afactor. Cell 47:413–422 Rosenbaum DM, Rasmussen SGF, Kobilka BK (2009) The structure and function of G-protein-coupled receptors. Nature 459:356–363 Schmidt WK, Tam A, Fujimura-Kamada K, Michaelis S (1998) Endoplasmic reticulum membrane localization of Rce1p and Ste24p, yeast proteases involved in carboxyl-terminal CAAX protein processing and amino-terminal a-factor cleavage. Proc Natl Acad Sci USA 95:11175–11180 Schmoll M, Seibel C, Tisch D, Dorrer M, Kubicek CP (2010) A novel class of peptide pheromone precursors in ascomycetous fungi. Mol Microbiol 77:1483– 1501 Schmoll M, Zeilinger S, Mach RL, Kubicek CP (2004) Cloning of genes expressed early during cellulase induction in Hypocrea jecorina by a rapid subtraction hybridization approach. Fungal Genet Biol 41:877–887 Schuurs TA, Dalstra HJ, Scheer JM, Wessels JG (1998) Positioning of nuclei in the secondary mycelium of Schizophyllum commune in relation to differential gene expression. Fungal Genet Biol 23:150–161 Seo JA, Han KH, Yu JH (2004) The gprA and gprB genes encode putative G protein-coupled receptors required for self-fertilization in Aspergillus nidulans. Mol Microbiol 53:1611–1623 Shen WC, Bobrowicz P, Ebbole DJ (1999) Isolation of pheromone precursor genes of Magnaporthe grisea. Fungal Genet Biol 27:253–263 Shi C, Kaminskyj S, Caldwell S, Loewen MC (2007) A role for a complex between activated G protein-coupled receptors in yeast cellular mating. Proc Natl Acad Sci USA 104:5395–5400 Shi C, Kendall SC, Grote E, Kaminskyj S, Loewen MC (2009) N-terminal residues of the yeast pheromone

95

receptor, Ste2p, mediate mating events independently of G1-arrest signaling. J Cell Biochem 107:630–638 Shiu PK, Glass NL (2000) Cell and nuclear recognition mechanisms mediated by mating type in filamentous ascomycetes. Curr Opin Microbiol 3:183–188 Singh A, Chen EY, Lugovoy JM, Chang CN, Hitzeman RA, Seeburg PH (1983) Saccharomyces cerevisiae contains two discrete genes coding for the a-factor pheromone. Nucleic Acids Res 11:4049–4063 Spellig T, Bo¨lker M, Lottspeich F, Frank RW, Kahmann R (1994) Pheromones trigger filamentous growth in Ustilago maydis. EMBO J 1994:1620–1627 Stajich JE, Berbee ML, Blackwell M, Hibbett DS, James TY, Spatafora JW, Taylor JW (2009) The fungi. Curr Biol 29:R840–R845 Stefan CJ, Blumer KJ (1994) The third cytoplasmic loop of a yeast G-protein-coupled receptor controls pathway activation, ligand discrimination, and receptor internalization. Mol Cell Biol 14:3339–3349 Sto¨tzler D, Betz R, Duntze W (1977) Stimulation of yeast mating hormone activity by synthetic oligopeptides. J Bacteriol 132:28–35 Sto¨tzler D, Duntze W (1976) Isolation and characterization of four related peptides exhibiting a factor activity from Saccharomyces cerevisiae. Eur J Biochem 65: 257–262 Strandberg R, Nygren K, Menkis A, James TY, Wik L, Stajich JE, Johannesson H (2010) Conflict between reproductive gene trees and species phylogeny among heterothallic and pseudohomothallic members of the filamentous ascomycete genus Neurospora. Fungal Genet Biol 47:869–878 Thompson JD, Gibson TJ, Higgins DG (2002) Multiple sequence alignment using ClustalW and ClustalX. Curr Protocols Bioinform 2:Unit 2.3 Turgeon BG, Yoder OC (2000) Proposed nomenclature for mating type genes of filamentous ascomycetes. Fungal Genet Biol 31:1–5 Turina M, Prodi A, Alfen NKV (2003) Role of the Mf1-1 pheromone precursor gene of the filamentous ascomycete Cryphonectria parasitica. Fungal Genet Biol 40:242–251 Vaillancourt LJ, Raudaskoski M, Specht CA, Raper CA (1997) Multiple genes encoding pheromones and a pheromone receptor define the b1 mating-type specificity in Schizophyllum commune. Genetics 146: 541–551 Wang SH, Xue CB, Nielsen O, Davey J, Naider F (1994) Chemical synthesis of the M-factor mating pheromone from Schizosaccharomyces pombe. Yeast 10: 595–601 Wang Y, Dohlman HG (2004) Pheromone signaling mechanisms in yeast: a prototypical sex machine. Science 306:1508–1509 Waters MG, Evans EA, Blobel G (1988) Prepro-a-factor has a cleacable signal sequence. J Biol Chem 263: 6209–6214 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

96

Stefanie Po¨ggeler

gene and putative pheromone genes. EMBO J 14:5271–5278 Willer M, Hoffmann L, Styrkarsdottir U, Egel R, Davey J, Nielsen O (1995) Two-step activation of meiosis by the mat1 locus in Schizosaccharomyces pombe. Mol Cell Biol 15:4964–4970 Woo PC, Chong KT, Tse H, Cai JJ, Lau CC, Zhou AC, Lau SK, Yuen KY (2006) Genomic and experimental evidence for a potential sexual cycle in the pathogenic thermal dimorphic fungus Penicillium marneffei. FEBS Lett 580:3409–3416

Xue C, Hsueh YP, Heitman J (2008) Magnificent seven: roles of G protein-coupled receptors in extracellular sensing in fungi. FEMS Microbiol Rev 32:1010–1032 Zhang L, Baasiri RA, Van Alfen NK (1998) Viral repression of fungal pheromone precursor gene expression. Mol Cell Biol 18:953–959 Zhang L, Churchill AC, Kazmierczak P, Kim DH, Van Alfen NK (1993) Hypovirulence-associated traits induced by a mycovirus of Cryphonectria parasitica are mimicked by targeted inactivation of a host gene. Mol Cell Biol 13:7782–7792

6

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

URSULA KU¨ES1, TIMOTHY Y. JAMES2, JOSEPH HEITMAN3

CONTENTS I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 97 II. A Short Treatise on the Molecular Functions of Mating Type Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 A. The Mating Type Locus Encoding Two Classes of Homeodomain Transcription Factors (HD Locus) . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 B. The Mating Type Locus Encoding Pheromone Precursors and Pheromone Receptors (P/R Locus) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 106 C. On the Origin of Tetrapolar Mating Type Determination . . . . . . . . . . . . . . . . . . . . . . . . . . . 108 III. Mating Type Genes in Pucciniomycotina . . . . . . . . 109 A. Microbotryomycetes, Sporidiobolales . . . . . . . . 109 B. Microbotryomycetes, Microbotryales . . . . . . . . . 114 C. Pucciniomycetes, Pucciniales. . . . . . . . . . . . . . . . . . 114 D. Evolutionary Considerations . . . . . . . . . . . . . . . . . . 115 IV. Mating Type Genes in Ustilaginomycotina . . . . . . 116 A. Ustilaginomycetes, Ustilaginales . . . . . . . . . . . . . . 116 B. Ustilaginomycotina Incertae Sedis, Malasseziales . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120 C. Evolutionary Considerations . . . . . . . . . . . . . . . . . . 121 V. Mating Type Genes in Agaricomycotina . . . . . . . . . 126 A. Tremellomycetes, Filobasidiales. . . . . . . . . . . . . . . 126 B. Tremellomycetes, Tremellales . . . . . . . . . . . . . . . . . 133 1. Mitosporic Tremellales/Sensu Lato Kwoniella Clade . . . . . . . . . . . . . . . . . . . . . . . . . . . . 133 2. Tremellacecae. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 134 C. Agaricomycetes, Auriculariales. . . . . . . . . . . . . . . . 134 D. Agaricomycetes, Cantharellales . . . . . . . . . . . . . . . 135 E. Agaricomycetes, Agaricales, Polyporales, and Russulales. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135 1. Tetrapolar Species . . . . . . . . . . . . . . . . . . . . . . . . . . 135 2. Bipolar Species . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 139 3. Homothallic Species. . . . . . . . . . . . . . . . . . . . . . . . 141 F. Evolutionary Considerations . . . . . . . . . . . . . . . . . . 142

1

Division of Molecular Wood Biotechnology and Technical Mycology, Bu¨sgen-Institute, University of Go¨ttingen, Bu¨sgenweg 2, 37077 Go¨ttingen, Germany; e-mail: [email protected] 2 Department of Ecology and Evolutionary Biology, University of Michigan, 830 N. University, Ann Arbor, MI 48109, USA; e-mail: [email protected] 3 Department of Molecular Genetics and Microbiology, Duke University Medical Center, 322 CARL Building, Research Drive, Durham, NC 27710, USA; e-mail: [email protected]

VI. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 144 References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 146

I. Introduction The Basidiomycota is a diverse phylum of economically and environmentally important fungi that produce meiotic spores on the surface of flask-shaped cells termed basidia. The Basidiomycota shares a monophyletic origin with the Ascomycota as the two phyla of the Dikarya. The new taxonomic system introduced by Hibbett et al. (2007) defines three subphyla within the Basidiomycota: Pucciniomycotina (including rust fungi, basidiomycetous yeasts, smut-like fungi, and jelly-like fungi), Ustilaginomycotina (true smut fungi), and the Agaricomycotina (including some yeasts, jelly fungi, and the mushroomforming species). Representatives from all three subphyla have been analysed for their mating behaviour (Whitehouse 1949, 1952), and the number of species for which data is available on the molecular control of mating and mating type is rapidly increasing (for a compilation, see Table 6.1). According to the data collected on ~230 species by Whitehouse (1949), ~90% of all basidiomycetes are heterothallic and require cell–cell fusion between two independent strains of different mating type for sexual reproduction (heterothallism), whereas ~10% of species are self-fertile, homothallic, and undergo sexual reproduction without a partner of another mating type (primary homothallism). Not included in this analysis are pseudo-homothallic species that have reduced outcrossing by incorporating haploid nuclei of distinct mating types within one basidiospore (secondary homothallism), which upon germination gives rise to a sexually competent mycelium. In this mating type configuration, the existence of different mating types is masked (Whitehouse 1949). Evolution of Fungi and Fungal-Like Organisms, The Mycota XIV S. Po¨ggeler and J. Wo¨stemeyer (Eds.) © Springer-Verlag Berlin Heidelberg 2011

Ustilaginomycotina Ustilaginomycetes

Pucciniomycetes

Ustilaginales

Pucciniales

Ustilaginaceae

Cronartiaceae

Melampsoraceae

Pucciniaceae

Microbotryales Microbotryaceae

Microbotryomycetes Sporidiobolales Sporidiobolaceae

Family

Pucciniomycotina

Order

Class

Sub-phylum

Table 6.1. Mating systems in selected basidiomycetes

Bipolar/pseudobipolar

Tetrapolar

U. maydis


50 (2 a, 25 b)

2 (MAT-1 and MAT-2)

? ?

Tetrapolar Tetrapolar Bipolar

?

2 (+ and –)

? 2 (+ and –) 2 (+ and –)

2 (A1 and A2)

0?

2 (A1 and A2)

?

Bipolar

Tetrapolar Bipolar Bipolar

Bipolar

Gymnosporangium asiaticum Melampsora laricipopulina M. lini Cronartium quercuum Ustilago hordei

Microbotryum violaceum Puccinia coronata P. graminis P. triticina

Sporobolomyces Anamorphic? spec. IAM 13481

Sporobolomyces metaroseus (anamorph S. roseus) Sporidiobolus salmonicolor

Sporidiobolus johnsonii

Bipolar/pseudo- 2 (A1 and A2) bipolar Bipolar/pseudo- 2 (A1 and A2) bipolar, some strains are homothallic, others anamorphic Bipolar, some 2 (A1 and A2) strains are anamorphic

R. babjevae

2 (A1/A and A2/a)

Total mating typesa

Bipolar

Mating system

Rhodosporidium toruloides

Species

Lawrence (1980) Yamazaki and Katsuya (1988) Groth (1975), Martinez-Espinoza et al. (1993) Puhalla (1970), Silva (1972)

Narisawa et al. (1994) Craigie (1927, 1931) Allen (1932), Boulton et al. (2008) Tetsuka and Katsuya (1984) Duplessis et al. (2011)

Statzell-Tallman and Fell (1998), Vale´rio et al. (2008a), Coehlo et al. (2010) Coelho et al. (2008), Valerio et al. (2008b) Day (1979)

Valerio et al. (2008b)

Banno (1967), Abe et al. (1975), Fell and StatzellTallman (1998) Golubev (1993), Coehlo et al. (2010) Fell and Statzell Tallman (1981), Valerio et al. (2008b)

References

98 Ursula Ku¨es et al.

Agaricomycotina

Agaricomycetes

Tremellomycetes

Ustilaginomycotina Exobasidiomycetes incertae sedis

Mitosporic Tremellales/ sensu lato Kwoniella clade

Tremellales

Ceratobasidiaceae

Cantharellales

Hydnaceae

Auriculariaceae

Auriculariales

Tremellaceae

Filobasidiaceae

Tilletiaceae

Tilletiales

Filobasidiales

Malasseziaceae

Malasseziales Bipolar Bipolar

T. tritici

Tetrapolar

Bipolar

Thanatephorus cucumeris (Rhizoctonia solani) species complex

Homothallic or bipolar

?

Multiple (20 A, 14 B) ?

2 A (a and A), 68 B (146 B estimated) 2 A (a and A), ? B Multiple

continued

Wong and Wells (1987) Cubeta and Vilgalys (1997), Julian et al. (1997), Toda and Hyakumachi (2006), Qu et al. (2008a, b)

Ishibashi et al. (1984) Whitehouse (1949)

Statzell-Tallman et al. (2008) Bandoni (1965), Wong and Wells (1985)

2 (A and a)

?

?

Rodriguez-Carres et al. (2010) Hsueh et al. (2011) Metin et al. (2010)

0

Homothallic

Hsueh et al. (2011)

Hoffmann and Kendrick (1965) Holton and Kendrick (1957), Kollmorgen and Trione (1980), Trail and Mills (1990) Fraser et al. (2003) Kwon-Chung (1976), Kwon-Chung and Bennett (1978)

Albert and Schenk (1996) Schirawski et al. (2005) Xu et al. (2007)

Multiple (2 A,
6 B)

?

Heterothallic

Tetrapolar

2 (a and a) 2 (a and a)

2

Mating never observed Multiple (
5)


15 (3 a,
5 b)

2 (+ and –)

Bipolar Bipolar

T. brasiliensis Tetrapolar Auricularia auricula Bipolar judae A. polytricha Tetrapolar

Kwoniella mangroviensis Tremella mesenterica

Cryptococcus gattii C. neoformans (Filobasidiella neoformans) var. neoformans and var. grubii C. amylolentus (F. amylolenta) Filobasidiella depauperata Tsuchiyaea wingfieldii C. (Kwoniella) heveanensis

Tetrapolar

Sporisorium reilianum Malassezia globosa and M. restricta Tilletia controversa ?

Bipolar

U. scitamineum

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality 99

Class

Russulales

Polyporales

C. scobicola Coprinellus disseminatus Schizophyllum commune

Psathyrellaceae

Phanerochaetaceae Phanerochaete chrysosporium Amylostereaceae Amylostereum areolatum

Polyporaceae

Fomitopsidaceae

Strophariaceae

Pholiota microspora (¼P. nameko) Fomitopsis palustris (¼ Polyporus palustris) Postia placenta Lentinus squarrosulus

Tetrapolar Bipolar

Coprinopsis cinerea

Mitosporic Marasmiaceae Pleurotaceae

Schizophyllaceae

Tetrapolar

Moniliophthora perniciosa Pleurotus djamor

Marasmiaceae

Boidin and Lanquentin (1984), van der Nest et al. (2008, 2009)

? ? A;
3 B Tetrapolar

Bipolar

Nobles (1943) Dhitaphichit and Nawawongwiwat (2006) James et al. (2011)

? ?

Bipolar Tetrapolar

Raper (1966)

20

Bipolar

Bipolar

Tetrapolar

Tetrapolar

Homothallic

Tetrapolar

Lentinula edodes

Hydnangiaceae

Agaricales

References

Lemke (1969), Ullrich and Raper (1975) Raffle et al. (1995), Niculita-Hirzel et al. (2008) Multiple (40–65 A, Takemaru (1961), 63–100 B) Tokimoto et al. (1973), Fox et al. (1994) 0 Griffith and Hedger (1994) Multiple James et al. (2004b) (estimated: 58 A, 231 B) Multiple Raper (1966) (estimated 160 A, 81 B)
49 (7 A, 7 B) Kemp (1974) Multiple (123 James et al. (2006) estimated) Multiple Raper (1966) (estimated: 288 A, 81 B) Multiple (
6) Aimi et al. (2005)

Homothallic and bipolar Tetrapolar Multiple (45 A, 24 B)

Sistotrema brinkmannii Laccaria bicolor

Total mating typesa

Mating system

Species

Family

Order

a Note that by historical reason the naming of mating type loci a in the Ustilaginomycotina, A in the Tremellomycetes, and B in the Agaricomycetes correspond to P/R mating type genes; and b in the Ustilaginomycotina, B in the Tremellomycetes, and A in the Agaricomycetes correspond to HD mating type genes.

Sub-phylum

Table 6.1. continued 100 Ursula Ku¨es et al.

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

Mating in heterothallic basidiomycetes involves the fusion of two self-sterile strains (haploid yeast cells or filamentous monokaryons) each with a distinct type of haploid nucleus. Following cell– cell fusion, nuclear fusion is delayed, and the two nuclei congress and pair but do not fuse, forming a dikaryon with two distinct types of haploid nuclei in its cells, one from each parent (Meinhardt and Esser 1990; Hiscock and Ku¨es 1999). The dikaryon grows as hyphae, in some cases with fused clamp connections, and sexual reproduction occurs, with nuclear fusion (karyogamy), meiosis, and the formation of basidia and basidiospores. The culmination of the sexual cycle is restricted to a single specialized cell, the basidium, which forms at the termini of dikaryotic hyphae (Ku¨es 2000; Hull and Heitman 2002). Reproduction in homothallic fungi may include either uniparental reproduction (apomixis) without a partner or biparental reproduction (amphimixis). Mating in homothallic fungi can involve either: (1) the fusion between two selffertile strains (outcrossing), or (2) cell(s) or nuclei of the same strain undergoing sexual reproduction (inbreeding by selfing) (Lin and Heitman 2007). There are several different molecular bases for homothallic sexual reproduction in fungi. First, in the Ascomycota, mating type switching enables mother cells to switch mating type via recombination of silent mating type cassettes into the expressed mating type locus (MAT locus), allowing cell–cell fusion of mother with daughter cells (Haber 2007; Lin and Heitman 2007; Nielsen and Egel 2007). This pattern of homothallism is well established and arose independently in Saccharomyces cerevisiae (and closely related species) and Schizosaccharomyces pombe (Arcangioli and Thon 2004; Butler et al. 2004). Recent studies reveal that, while the three mating type cassettes share a common ancestral origin in S. cerevisiae and Kluveromyces lactis, the mechanisms and enzymes involved in high efficiency mating type switching evolved independently (Barsoum et al. 2010; Rusche and Rine 2010). There is only one known example of mating type switching in the Basidiomycota, and it is unknown whether the mating type switching in Agrocybe aegerita also involves silent mating type cassettes (Labare`re and Noe¨l 1992). Second, in other homothallic species, fusions have occurred between two alternate forms of the mating type locus (idiomorphs;

101

Metzenberg and Glass 1990) to produce hybrid mating type loci that promote self-fertility (such as Cochliobolus heterostrophus and Sordaria macrospora; Lin and Heitman 2007; Po¨ggeler 2007; Turgeon and Debuchy 2007; see Chapters 5 and 12 this volume). Third, in certain homothallic species including Aspergillus nidulans and Neosartorya fischeri, both mating type idiomorphs are present in the genome, but are unlinked (Lin and Heitman 2007; Rydholm et al. 2007; see Chapters 5 and 12 in this volume). Finally, a novel form of homothallism has recently been discovered, known as same-sex mating or unisexual reproduction (Lin et al. 2005). In this case, isolates of a single mating type are capable of mating with either themselves or other members of the population of the same mating type. This unusual form of self-fertile mating was first discovered for the human pathogenic basidiomycete Cryptococcus neoformans and population genetics studies provide evidence that this form of mating occurs in nature for both this and the related species Cryptococcus gattii (Fraser et al. 2005; Lin et al. 2007, 2009, 2010; Bui et al. 2008; Hiremath et al. 2008; Saul et al. 2008; Byrnes et al. 2010; see Section V.A). Moreover, the most common human fungal pathogen, the ascomycete Candida albicans has also recently been discovered to undergo samesex mating (Alby et al. 2009; Heitman 2009). Mating cells (“gametes”) in the Basidiomycota typically do not differ in shape and overall size or cell fusion properties, and sexual reproduction is thus isogamous (Billiard et al. 2011). However, for C. neoformans the responses of mating partners to pheromones can differ, and in some heterothallic mating situations one mating partner produces long conjugation tubes whereas the other forms isotropically enlarged round cells. This may therefore present a form of fungal anisogamy that deserves further exploration in this and other species (Wang et al. 2000; Hull and Heitman 2002; McClelland et al. 2004; see Section V.A). For heterothallic mating to be successful, the haploid individuals must be of complementary mating type to undergo cell fusion. Strains of the same mating type are usually incompatible and incapable of forming a dikaryon (Whitehouse 1949; Raper 1966). As noted above, recent studies with Cryptococcus species indicate that in some cases species can also undergo same-sex mating, i.e., fusion of cells of the same mating type in which sexual reproduction involves ploidy changes

102

Ursula Ku¨es et al.

(possibly via karyogamy, meiosis, and basidiospore formation; Lin et al. 2005; see Section V.A). Mating types and mating type incompatibility in fungi are physiologically defined and the fact that most species exhibit isogamy may have facilitated the evolution of multiple mating types by reducing a potential morphological barrier to fertility of novel mating types (Billiard et al. 2011). A striking and defining feature of many species of the Basidiomycota is the existence of multiple mating types (more than two in a single species), and this phenomenon has been verified for the majority of the Basidiomycota (Bensaude 1918; Kniep 1928; Whitehouse 1949). Beyond the Basidiomycota of the fungal kingdom, there are relatively few other documented examples of sexual reproduction known with more than two mating types. Notable examples include the slime mold Dictyostelium discoideum, which has three mating types defined by a tri-allelic mating type locus (Bloomfield et al. 2010), and the ciliate Tetrahymena thermophila, which has seven mating types but for which the molecular basis is as yet unknown (Phadke and Zufall 2009). Similar to the Ascomycota (see Chapter 12 in this volume), in the simplest cases, two different mating types exist within a basidiomycetous species. Examples include the wheat stem rust Puccinia graminis (Craigie 1931), smuts of the Microbotryum species complex (Devier et al. 2010), the barley smut Ustilago hordei (Bakkeren and Kronstad 1993), and the human pathogen C. neoformans (KwonChung 1976; Kwon-Chung and Bennett 1978; Hull and Heitman 2002). However, more frequently, species exhibit multiple mating types and these include the corn smut Ustilago maydis that has about 50 mating types (Puhalla 1970; Silva 1972; Banuett and Herskowitz 1989) and many mushrooms such as Coprinopsis cinerea, Pleurotus djamor and Schizophyllum commune that may have up to 12000– 24000 different ones when both mating loci are considered simultaneously (Raper 1969; James et al. 2004b). Further examples of multiple mating types can be found in Table 6.1.

The mating types in heterothallic basidiomycetes are determined by either one or two chromosomal mating type loci, which each have at least two alleles (Table 6.1). Depending on whether one or two loci control mating type and sexual reproduction, such heterothallic species are called bipolar (one mating type locus) or tetrapolar (two mating type loci), respectively. In tetrapolar species, compatibility between individuals only occurs when they differ at both mating type loci. ~55–65% of Agaricomycotina species

are thought to be tetrapolar (Whitehouse 1949; Raper and Flexer 1971), whereas the majority of the Ustilaginomycotina (Whitehouse 1952) and possibly also of the Pucciniomycotina (Allen 1936; Buller 1950) appear to be bipolar with just two different mating type alleles. The origin of the terms bipolar and tetrapolar derives from the patterns of F1 progeny produced by sexual reproduction. In bipolar systems, progeny of the two parental mating types are produced, whereas in tetrapolar species, mating of parental a1b1 and a2b2 isolates yields progeny of four mating types (tetrapolar): a1b1, a1b2, a2b1, and a2b2 (where a1, a2 and b1, b2 refer to different phenotypes determined by different alleles of the two mating type loci). In the recently discovered unique example of homothallic same-sex mating of C. neoformans, a parent or parents of just one mating type produce offspring that are all of the same parental mating type, and we introduce here the novel term “unipolar” to codify this pattern of sexuality relative to the well established nomenclature for bipolar and tetrapolar species. It is worth stressing in this context that C. neoformans does have a well established bipolar mating type system involving two mating types, one of which is extremely uncommon in nature (see Section V.A). Thus, it is like the well established model species S. cerevisiae in which Hoþ homothallic (which express the HO endonuclease required for mating type cassette switching) and ho– heterothallic (which lack the HO endonuclease) isolates co-exist in a well defined species (Butler et al. 2004; Haber 2007). Analogously, C. neoformans maintains both a heterothallic bipolar and a homothallic unipolar sexual cycle (see Section V.A.), illustrating that fungi are not strictly heterothallic or homothallic, but that isolates can co-exist in a species population that exhibit either or both mating patterns and can even mate with each other based on the isolation of natural HO+/ho diploid S. cerevisiae isolates (McCusker 2006). The mating type loci in the genomes of basidiomycetes differ but appear to be limited to three classes of genes encoding: (1) homeodomain transcription factors, (2) pheromone precursors, and (3) cognate G protein-coupled 7-transmembrane pheromone receptors. The principal functions of the different mating type proteins have been clarified over the past two decades by elegant molecular studies. Much insight has been gained into how these proteins orchestrate complex intracellular

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

signalling and gene regulation processes occurring during mating and sexual development and reproduction (for more detailed information on special aspects and individual fungi, see reviews by Kothe 1996, 1999, 2001; Kronstad and Staben 1997; Bo¨lker 1998; Casselton and Olesnicky 1998; Hiscock and Ku¨es 1999; Lengeler et al. 2000a; Feldbru¨gge et al. 2004, 2006; McClelland et al. 2004; Casselton and Challen 2006; Banuett 2007; Casselton and Ku¨es 2007; Fowler and Vaillancourt 2007; Kahmann and Schirawski 2007; Stanton and Hull 2007; Bakkeren et al. 2008; Hsueh and Heitman 2008; Xue et al. 2008; Morrow and Fraser 2009; Kothe and Raudaskoski 2010; Lee et al. 2010b; for a short overview on the general mechanisms, see Sections II.A and II.B). Importantly, mating type-like genes encoding either homeodomain transcription factors or pheromone precursors or pheromone receptors are also found in self-fertile, homothallic species lacking a mating type phenotype (Casselton and Challen 2006; Coelho et al. 2010; Ku¨es and Navarro-Gonza´lez 2010; Rodriguez-Carres et al. 2010) and in self-sterile anamorphic species that have not been observed to undergo mating reactions (Xu et al. 2007; Coelho et al. 2008, 2010). Furthermore, in a number of bipolar and tetrapolar species mating type-like genes are present that are unlinked to the functional mating type loci (James et al. 2004b, 2006; Aimi et al. 2005; Niculita-Hirzel et al. 2008; Hsueh et al. 2009; Martinez et al. 2009; Ohm et al. 2010). In the next sections, we summarize current knowledge on the structure of mating type loci in different species of basidiomycetes, giving special consideration to bipolar and tetrapolar heterothallic species (including pseudo-homothallic species) as well as related genes of true homothallic species and of anamorphic species not currently recognized to have an extant sexual cycle (see Table 6.1).

II. A Short Treatise on the Molecular Functions of Mating Type Genes A. The Mating Type Locus Encoding Two Classes of Homeodomain Transcription Factors (HD Locus) Mating type genes encoding homeodomain transcription factors (HD locus) are divided into two

103

different classes based on structural homologies and distinct sequences of the DNA-binding motifs of the homeodomains in the deduced products (Fig. 6.1). Generally for all of the basidiomycetes, the two classes of genes have been termed HD1 and HD2, respectively (Ku¨es and Casselton 1992, 1993), although the individual genes in different species may have been given different names (see below). The products of the HD2 genes of the basidiomycetes have a canonical 60-amino-acid homeodomain forming a three-helical structure and a typical conserved DNA-binding motif WFXNXR within helix 3 (Duboule 1994; Fig. 6.1B, see below). In contrast, the products of HD1 genes have an atypical homeodomain (known as a TALE homeodomain; TALE ¼ three amino acid loop extension; Bu¨rglin 1997) with an extra three amino acids forming a loop between helix I and helix II of the three-helical homeodomain structure. Furthermore, in some cases extra amino acids are present between helix II and helix III of the folded homeodomain structure. Another consistent difference is that the amino acid sequence is more variable for the HD1 homeodomain proteins in helix III and within the DNA binding motif (Duboule 1994; Fig. 6.1A, see below). Congruent with this, HD1 homeodomain proteins are more permissive of DNA mutations in the DNA binding motifs compared to HD2 homeodomain proteins (Ku¨es et al. 1994b; Asante-Owusu et al. 1996; Pardo et al. 1996; Schlesinger et al. 1997). In contrast, the functions of both homeodomain proteins are apparently required in C. neoformans, because when amino acid substitution mutations were introduced into the DNA-binding motifs of either protein the heterodimeric complex was no longer functional (Stanton et al. 2009). In most basidiomycetes, the HD1 and HD2 genes are arranged within the mating type locus as a pair of divergently transcribed genes (Ku¨es and Casselton 1992, 1993; Fig. 6.2). For a compatible mating, the product of an HD1 gene has to interact with the product of an HD2 gene from an allelic pair of genes, i.e., from a gene pair of another mating type specificity (e.g., see Gillissen et al. 1992; Specht et al. 1992; Bakkeren and Kronstad 1993; Ku¨es et al. 1994a, c; Pardo et al. 1996; Hull et al. 2002, 2005; Schirawski et al. 2005; Yi et al. 2010). Sequences of HD1 and HD2 alleles are generally highly divergent from each other and, as a consequence, the allelic gene products differ in

Ursula Ku¨es et al.

104 A. HD1 transcription factors SjHD1 SsHD1 SspbE1 RbHD1 MvbE1 MpbE1 PgbE1 PtbE1

HHSSPMGDDSASFSPYRRWFLDNFSNPYPSASDKTTLLALVPS--------------HNKQQLDTWFTNTRRRSGWQD SPPAPMGDDSASFAPYRRWFLGNFSNPYPSAFEKTTLLALVPS--------------HNKQQLDTWFTNTRRRSGWQD QAPIGDAGDLDPFH PLRLWFLDHFVNPYPDSKDKEHFLLVYPN--------------HTKQQIDTWFTNIRRRSGWQD PSPEPPADDQAHCAPYREYFLAHFSFPYPT VMDACSRTEDELVQPYRLWFLDNFAWPYPESSERLRLLAQVPS--------------HGRAQLSNWFVNSRRRSGWSR SPPPCSNLRKDPQDLLRCWSSQHLSYLFPTKEELMELSGQTK---------------LSAKQVDLWFRNARTRSGWSK PVAQDNSCSSPSHTILKNWSQAHMTYLFPTQPQLQELASQTS---------------STETKVNSWFRNARSRSGWSK EATEATSDTSTSHTILKNWSQARLSYLFPSPLELQELVSQTS---------------MSETKLNSWFRNARSRSGWAK

UhbE1 UmbE1 SrbE1 MgbE1

PSQAPDLSETLPSYHMRRHFLATLHDPYPAAEDKKALVRITNESVLQPGTPLKKRTLLSVEQLTLWFINARRRSGWSQ DVGCRNLSEDLPAYHMRKHFLLTLDNPYPTQEEKETLVRLTNESTARVGQSSVNRPPLEVHHVTLWFINARRRSGWSH FDVKQDL SETLPSYHMRKHFLATLDNPYPSQADKDTILRTTNDSAAGTG-----RPPLDAHQLTLWFINARRRSGWTH IRESDRRDPTLNAKH MRDWFLRHLGHPFPSREDKEQILAETNACIRDRT------LRLKYTQIVLWFINTRRRSGWTA

CgSxi1 CngSxi1 CnnSxi1 ChSxi1 TmSxi1

HISPKPLESAPDHTIVRLWFLNNLAYPYPTAQQKDSLAKYAG---------------MQRGKVDSDLTNYRRRAGWTD GNSPKPSESATDHTVVRLWFLDNLAYPYPTAQQKDFLAKTAG---------------IQRSQVDSDLTNYRRRAGWTD DNSPKPSEPPIDHTVVRLWFLDNLAYPYPTAQQKDFLAKTAG---------------IQRSQVDSDLTNYRRRAGWTD KKSSKAHSQEPDHSLIRHWFLHNISYPYPTVSMKETLASKAG---------------ITRAKVDSDLTNFRRRAGWTE SIPPFTTLLSPPTDPIRTWFISHLASPYPTPLEKVELATSAN---------------ITTNKLDSDLVNWRRRSGWAN

Cca1-2 Ccb1-2 Ccd1-1 Csa1-1 Lba1-1 PdA1-7 ScAαZ4 ScAβQ6 ScAβS6 ScAβT6 ScAβU6 CdA1-1 CdB1-1 Pca1-1 PnA3-hox1 MpHD1

TEDEKSDASPAYIAPAVQWLYQNLHNPYPSPQVRESIATKSK---------------SPRKDVDNWFVDARRRIGWND SENTDASRTPSYIEPCARWLKNNCHNPYPSSQMRAEIAVETG---------------ASTKDIDAWFIDARKRIGWND DASPKEPAEPAYIEPSCRWLKDNWYNPYPSPQVRSSIAKQTG---------------ASRKDIDAWFIDARKRIGWNE SSEPQNNSGPSYILPAAKWLYENVHAPYPSVEVRSSLAKQSG---------------CPRKDVDNWFIDARRRIGWND RSPRSSNTCPQYIEHAYKWLLRNLHNPYPTREDKASFLRHTT---------------STKCDLDHWFVDARKRMGWNS ASEEESLGHPSYIAPAYKWLLNNLHAPYPSKKQKSLISRQTS---------------TSVHHIDTWFLNVRRRIGWTA GAELSATPLPPYIEPCYRWLVNHLDNPYPTKAIKEELLDQARQRTSPDVAQH-----LALGDIDNWFIAARARMGWGD ASRSPRTCPPKRIDACYKWLLNHLHNPYPTREEREDLARQSS---------------SDIKSITAWFVTARQRIGWSN TPHPRSLAALEKLEPCYSWLASHLHNPYPTREEREALARECD---------------RTMPAITAWFVTARQRIGWSQ SSSRHKPAMSKVNEPCYRWLLNHLYNPYPTREEWEALSKESK---------------TSVKEVASWFMSARKRIGWSL KASTRVPAAPSLNEPCYRWLLANLHNPYPTQQQRERLRLESK---------------ATAKAITTWFVNARKRIGWSH SEDEMLGSLPPYFKLASQWLLANLHNPYPSSKVKESMALQTG---------------SPRKDIDHWFSEARKRVGWGD ATRISQAASPWYIEPSVRWLRTNIHNPYPPRAVRACIAGASR---------------SAIKDIDAWFIDARKRIGWSR SNRRHNPSLSPFIPKAYNWLVDNLHNPYPPTSLKSAIASDAG---------------LSLRTVDDWFKSVRRHIGWVS PISKVLDSIPVYVKPCCQWLLNNLHNPYPAKSVRNSICRQTG---------------YSIKDIDAWFVDARKRIGWNQ EPEEPPISSSPYVRAAYEWLIQNLHNPYPTTAQRDAISFQSG---------------TSRKTIDNWFIDVRKRIGWNA _____________ ________ _-_--_WFXNXR-----

Helix

I

II

III

B. HD2 transcription factors SsHD2 SspbW1 MvbW1 MpbW1 PgbW1 PtbW1

DDSPRGIFTEEVLDILNAAFEAGD--TVTRAEVQQLLTAVTG---------------LRARQVRTWFANQRQRRHKRA AAEPSGEFTEEVVEILRRAFDKC---ETLTRGEVKGLQTVTK---------------LNAKQIRTWFANQRQRRGRKA LPVVFGEFNEEVNNVLSTATNQF---EHLTKGERLALAKRTG---------------LTQKQVATWFANQRQRRTRRV RRGRPFVFSKDQTAVLRALLVHD---DQFSNEEKELIAQTLN---------------LTRAQVNRWFCNARARGGRKK AGIRPPKFTHKQTVVLNALLARD---NQYSTEEKDLIAHELD---------------MTPDQVNRWFCNARARKKPYS PRGRAPKFTKEQTAALNALLARD---NQYSSEDKELIAHELN---------------LTREQVNRWFCNARARKKPYS

UhbW1 UmbW1 SrbW1 MgbW1

STKPCRGHDSDAVRILEQAFQHT---PNITQAEKYRLAEVTG---------------LKPKQVTIWFQNRRNRKGRKG PLKTGRGHDSEAVRILEQAFKHS---PNITPAEKFRLSEVTG---------------LKPKQVTIWFQNRRNRKGKKN DTKTARGHDSDAIRILEQAFEIT---PNITQAEKFRLAEVTG---------------LQPKQVTIWFQNRRNRKGRKG PHETHRPHSVRATAILERAFAHA---PNITQAEKYKLAQATG---------------LQPRQVTIWFQNRRNRRAHAR

CgSxi2 CngSxi2 CnnSxi2 ChSxi2 TmSxi2

DIKRPSTFAPGTLNILESAYARC---TILSAAETALIAEAAS---------------ITPQQVRTWFQNKRNRGKKTR GIKRPSPFAPGTLHVLESAYSRC---TILSAAETALIAEAAS---------------ITPQQVRTWFQNKRNRGKKTR GTKRPSPFAPETLHVLESAYSRC---TVLSAAESALIAEAAS---------------ITPQQVRTWFQNKRNRGKKTR SARKLSTFSPHTIIVLDAAYSRS---KSLSQAETIIIAQAAG---------------ISPHQVRTWFQNKRNRGKNKK CPTSVNGFPSGALQVLESAYSRT---HMLSVRECHLIAQAAQ---------------LDPAQVRTWFQNRRAREKRAA

Cca2-2 Ccb2-2 Csa2-1 Lba2-1 PdA2-7 ScAαY4 ScAßR6 ScAßV6 CdA2-1 CdB2-1 Pca2-1 PnA3-hox2 Ppa2-1

SLPKRTPFNAEYTPLLEKYFEYN---AFPPLRDREWLARKSM---------------MTPRQIEVWFQNHRRRARNEG PSAKRTPFNSEYTPLLEKYFEYH---AYPSARDREWLARKTM---------------MTTRQIEVWFQNHRRRARKEG QESKRTPFNAEYTPLLEKYFEYN---AYPSVRDRERLARKSM---------------MTPRQIEVWFQNHRSRAKKEG KVNERRTFNQEYTPLLEKYFEYN---AYPSAPDRLLLARKSM---------------MTPRQIEVWFQNHRTRARKDG KPAAKTSFNADFVPFLEKYFEFN---AYPSAADRSLMARKSM---------------MTPRQIEVWFQNHRNRARKEG GKRSRPKFHSEYTPVLELYFHFN---AYPTYADRRILAEKTG---------------MLTRQITVWFQNHRRRAKGPL EAQPVNLLKRARR PLLDRYFDLN---AYPSVTDKKALAAHEG---------------ATYRQIHVWFQNRRAKARAMN ISSGKDALDNRNHPVLEEYLKHN---AYPSPIDKQRLAEQEG---------------MSYRQVHVWFQNRRARAKEKH AHQAKRPFNHMCLPLLQTYFEHN---AYPSAQDRAMLARKST---------------LCPRQIEVWFQNHRRRAKKEG PSLKKAAFNVEYTPVLEQYFEYN---AYPSARDRELLARKSS---------------MTPRQIEVWFQNHRRRARREG RNIPDINLFNQALPLLKQAFSHD---QYPSRVEKERLANITG---------------LEYKQVSVWFQNHRSRTKKGG RENKRPAFNNEYTPMLENYFENN---AYPSRPDRVLLARKSC---------------MTERQIEVWFQNHRNRSKKEG PRHPHCPFNQNALPILVDFFERN---AFPSPSQKSRLASRTG---------------MNYRQIAVWFQNRRHRFRKDG _____________ ________ _-_--_WFXNXR-----

Helix

I

Fig. 6.1. Homeodomain sequences as found in HD1 (A) and HD2 (B) transcription factors from mating type loci of selected basidiomycetes. Underneath the aligned sequences, the three helical regions of the homeodomains are indicated by black blocks and the positions of the conserved amino acids of the DNA binding motif are indicated within the third helix by bold white letters (after Duboule 1994). Sequences in A and B are arranged in groups corresponding to the sub-phylum Pucciniomycotina, the sub-phylum Ustilaginomycotina, and then the

II

III

classes Tremellomycetes and Agaricomycetes of the subphylum Agaricomycotina, respectively. Available sequences for homeodomains were taken from databases as follows: Sporidiobolus johnsonii (Sj; GenBank GU474692), S. salmonicolor (Ss; GenBank GU474641; translated from BAUG37624, http://www.ncbi.nlm.nih.gov/Traces/trace.cgi?), Sporobolomyces spec. strain IAM 13481 (Ssp; protein IDs 34712 and 34718 on http://genome.jgi-psf.org/Sporo1/Sporo1. home.html), Rhodosporidium babjevae (Rb; incomplete sequence GenBank HM143857), Microbotryum violaceum

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality HD 1-1 HD 2-1

HD 1-2 HD 2-2

Fig. 6.2. Simplified model of two alleles of a basidiomycete mating type locus containing divergently transcribed genes for HD1 and HD2 transcription factors. The gene regions encoding the relatively conserved homeodomains HD1 and HD2 are marked by blue and light blue bars, respectively. Different shading of gene sequences indicates high levels of sequence divergence between allelic genes of the HD1 class and the HD2 class. Different numbers added with a dash to the end of gene class names refer to different alleles of these genes. Interactions occurring during mating between products of foreign HD1 and HD2 genes are marked by arrows. Note that the actions of HD1 and HD2 gene products occur only after cellular fusion and function to both establish and maintain the dikaryon (e.g., for the mushroom Coprinopsis cinerea, see Tymon et al. 1992; Ku¨es et al. 1998, 2002; Ku¨es 2000). Therefore, with a complete HD1 and HD2 gene pair in both the two allelic loci, there are two redundant interactions possible that exert the same regulatory function. As a consequence, while the most efficient mating type locus structure has a pair of allelic loci, an HD1 gene from one locus and an HD2 gene from the other locus can be lost without loss of sexual fecundity as long as the remaining HD2 and HD1 partner genes are compatible and functional (Ku¨es and Casselton 1992, 1993)

105

protein sequence. Depending on the species, sequence differences are either found concentrated at the N-terminus or distributed over the entire protein (Gillissen et al. 1992; Stankis et al. 1992; Bakkeren and Kronstad 1993; Gieser and May 1994; Specht et al. 1994; Badrane and May 1999; Schirawski et al. 2005; James et al. 2006; Yi et al. 2009; Coelho et al. 2010). Differences in the Ntermini are responsible for mating type specificity of the proteins. The N-termini act as dimerization domains between HD1 and HD2 proteins from allelic HD1 and HD2 (HD1/HD2) gene pairs whereas HD1 and HD2 proteins from the same mating type locus allele fail to interact. Lack of HD1 and HD2 dimerization is thus the mechanism of self-discrimination (Yee and Kronstad 1993; Ku¨es et al. 1994a; Banham et al. 1995; Ka¨mper et al. 1995; Asada et al. 1997; Yue et al. 1997; Romeis et al. 1997; Yee and Kronstad 1998; Hull et al. 2005; Yi et al. 2010). Dimerization between non-self proteins yields an active heterodimeric HD1-HD2 transcription factor that is nuclear localized, binds to specific sequences in the promoters of target genes, and regulates expression of genes acting in dikaryon development and sexual reproduction (Spit et al. 1998; Schlesinger et al. 1997; Romeis et al. 2000; Brachmann et al. 2001; Robertson et al. 2002; Basse and Farsing 2006; Ekena et al. 2008; Stanton et al. 2009; Heimel

ä (Mv; translated from GenBank AEIJ01001054), Melampsora laricis-populina (Mp; protein ID 90168 and 124184 on http:// genome.jgi-psf.org/Mellp1/Mellp1.home.html), Puccinia graminis (Pg; translated from GenBank AAWC01001065), P. triticina (Pt; translated from GenBank ADAS01011712), Ustilago hordei (Uh; GenBank CAA79218 and CAJA2018), U. maydis (Um; GenBank AAA63553 and EAK81227), Sporisorium reilianum (Sr; GenBank CAI59728 and CAI59727); Malassezia globosa (Mg; GenBank EDP44402 and EDP44401), Cryptococcus gattii (Cg; GenBank AAV28797 and AAZ07733), C. neoformans var. grubii (Cng; GenBank AAN75175 and AAV98451), C. neoformans var. neoformans (Cnn; GenBank AAW42798 and AAV98474), C. heveanensis (Ch; GenBank ACZ51518 and ACZ51523), Tremella mesenterica (Tm; protein ID 57675 and a modified model of protein ID 65105 on http://genome.jgi-psf.org/Treme1/ Treme1.home.html), Coprinopsis cinerea (Cc; GenBank EAU92789, EAU92786, EAU92783, EAU92788, and EFI28677), C. scobicola (Cs; GenBank AAK17069 and AAK17010), Laccaria bicolor (Lb; protein IDs 301103 and 379291 on http://genome.jgi-psf.org/Lacbi1/Lacbi1. home.html), Pleurotus djamor (Pd; GenBank AAS46737 and AAS46736), Schizophyllum commune (Sc; protein IDs

80879, 269864, 269862, 269861, 269858, 231556, 269851, and 269863 on http://genome.jgi-psf.org/Schco1/Schco1. home.html), Coprinellus disseminatus (Cd; GenBank AAZ14916, AAZ14918, AAZ14917, and AAZ14919), Phanerochaete chrysosporium (Pc; protein IDs 7555 and 7556 on http://genome.jgi-psf.org/Phchr1/Phchr1.home. html), Pholiota microspora ¼ P. nameko (Pn; GenBank BAH22610 and BAH22609), Postia placenta (Pp; protein ID 135106 on http://genome.jgi-psf.org/Pospl1/Pospl1.home. html), Moniliophthora perniciosa (Mp; translated from GenBank ABRE01018067 and ABRE01001637). Note that the protein model for MgbW1 from GenBank EDP44402 has been altered by deleting the sequence VQHACTDNPLQ from between the amino acids W and FQNRR of the sequence of DNA-binding motif in helix 3, due to the presence of a short intron in the corresponding DNA sequence that has been overlooked in the prediction of the current gene model. An intron is usually present at this position in HD2 genes of other species (see DNA sequences to the listed HD2 proteins in the respective databases). An intron appears also to be present at this place in the T. mesenterica SXI2 gene but this has an unusual length of 620 bases (to be confirmed by cDNA analysis)

106

Ursula Ku¨es et al.

et al. 2010a, b; Kruzel and Hull 2010; Lin et al. 2010; Stajich et al. 2010; Wahl et al. 2010).

B. The Mating Type Locus Encoding Pheromone Precursors and Pheromone Receptors (P/R Locus) Basidiomycota mating type loci may encode pheromone precursors and pheromone receptors, a molecular genetic organization that is unusual in the fungal kingdom, as the corresponding genes are not part of the mating type loci of the Ascomycota or Zygomycota sensu lato (Lee et al. 2010; see Chapter 5 in this volume). Hence, this unique arrangement of the pheromone precursor and pheromone receptor genes as a molecular determinant of mating type arose within the Basidomycota, and together with the novel tetrapolar mating system are unique features that provide a window into evolutionary events that fashion sex determinants and their genomic organization. Currently, there is no uniform nomenclature for either the basidiomycete pheromone precursor or pheromone receptor genes, but much can be inferred by reference to baker’s yeast (S. cerevisiae). Pheromones and pheromone receptors from mating type loci in basidiomycetes are homologous to the lipid-modified MFa pheromones (also known as a-factors) and the G protein-coupled 7-transmembrane Ste3 pheromone receptor of the ascomycetous yeast S. cerevisiae (Casselton and Olesnicki 1998; Bakkeren et al. 2008; Xue et al. 2008; Devier et al. 2009; see Chapter 5 in this volume). Therefore, in the simplified model shown in Fig. 6.3 of a mating type locus for a pheromone–pheromone receptor system, the pheromone receptor genes are labelled STE3 and, indeed, such nomenclature is also already in use in some basidiomycetous species (e.g., Fraser et al. 2004; Coehlo et al. 2008, 2010; Niculita-Hirzel et al. 2008). In contrast, using the S. cerevisiae nomenclature (MFa and MFa) to name pheromone variants is only possible for bipolar species that possess only a pair of opposite mating types such as C. neoformans and some related heterothallic species or a few tetrapolar species having one biallelic mating type locus for pheromone communication and thus only two alternate types of pheromones such as U. maydis and most species with a haploid yeast phase (Fraser et al. 2004; Schirawski et al. 2005; Metin et al. 2010). Hence,

Ph-1

STE3-1

Ph-2

STE3-2

Fig. 6.3. Simplified model of two alleles of a basidiomycete mating type locus containing genes for pheromone precursors (called here Ph) and pheromone receptors (STE3). Different shading of genes indicates high levels of sequence divergence between allelic genes, and different numbers added with a dash to the end of the gene class names refer to different alleles of these genes. Interactions occurring during mating between foreign Ph and STE3 genes are marked by arrows. Unlike the situation of the mating type locus with a complete HD1 and HD2 gene pair for the two distinct classes of homeodomain transcription factors (Fig. 6.2), note that the two different pheromone and pheromone receptor reactions are not necessarily redundant in function, at least not in the species where pheromone-mediated courtship prior to cell fusion is part of the mating process (Bo¨lker et al. 1992; MartinezEspinoza et al. 1992; Bakkeren and Kronstad 1996; Snetselaar et al. 1996; Davidson et al. 2000; Shen et al. 2002; Chang et al. 2003; Stanton et al. 2010)

the pheromone precursor genes in the generalized scheme shown in Fig. 6.3 are called Ph (for pheromone) in a manner similar to that already applied in some species with genes for many different pheromone precursors such as in the ectomycorrhiza-forming mushroom Laccaria bicolor (Niculita-Hirzel et al. 2008) and the poplar rust Melampsora larici-populina (http://genome.jgipsf.org/Mellp1/Mellp1.home.html; Duplessis et al. 2011). The genes encoding pheromone precursors and their receptors (P/R locus) are usually linked within a defined mating type locus, and in tetrapolar species are unlinked to the homeodomain transcription factor gene locus (e.g., see Bo¨lker et al. 1992; Schirawski et al. 2005; Niculita-Hirzel et al. 2008; Metin et al. 2010; Ohm et al. 2010; Stajich et al. 2010). In bipolar and the recently described pseudo-bipolar system (see details in Section III.A), the two mating type loci are either linked or fused (Bakkeren and Kronstad 1994; Fraser et al. 2004; Coehlo et al. 2010). As with the allelic mating type genes for homeodomain transcription factors, the pheromone precursor and pheromone receptor gene alleles from different mating types of a species are generally highly

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

divergent in sequence (see data e.g., in Bo¨lker et al. 1992; Wendland et al. 1995; Bakkeren and Kronstadt 1996; Urban et al. 1996a; O’Shea et al. 1998; Halsall et al. 2000; Karos et al. 2000; Lengeler et al. 2000b; Fowler et al. 2001, 2004; Gola and Kothe 2003b; Riquelme et al. 2005; Schirawski et al. 2005). This high sequence divergence is likely related to the mechanism by which self and nonself pheromones are discriminated (Fig. 6.3). Studies of basidiomycete pheromones and pheromone receptors heterologously expressed in S. cerevisiae suggest that pheromones bind to pheromone receptors from compatible mating types but not (effectively) to pheromone receptors of the same mating type (Hegner et al. 1999; Fowler et al. 1999; Olesnicky et al. 1999, 2000). In species undergoing courtship (a mutual communication process that is mediated by secreted pheromones that prepares the mating partners for fusion, such as polarized cell growth; for examples, see Sakagami et al. 1981a; Abe 1987; Martinez-Espinoza et al. 1993; Snetselaar et al. 1996; Sherwood et al. 1998; Davidson et al. 2000; Shen et al. 2002; GarciaMuse et al. 2003) prior to cell–cell fusion, mature pheromones have in some cases been isolated and characterized from culture supernatants. In the Basidiomycota mature pheromones are lipopeptides, ~9 to 14 amino acids in length that end with a C-terminal cysteine residue that is modified by farnesylation and carboxy-methylation, rendering it more hydrophobic. The mature pheromones are derived from larger pheromone precursors by both N-terminal and C-terminal processing (Abe et al. 1975; Kamiya et al. 1978a– c; Sakagami et al. 1979, 1981b; Spellig et al. 1994; Koppitz et al. 1996; Kosted et al. 2000; Schirawski et al. 2005; see Chapter 5 in this volume), and then subsequently exported from the cell by a noncanonical secretory mechanism involving a multidrug resistance (Mdr) pump related protein known as Ste6 based on its original description as an a-cell specific sterile mutation in S. cerevisiae that was unable to export a-factors. Related pheromone export pumps have been identified in the Basidiomycota (Hsueh and Shen 2005). Pheromone precursors in the basidiomycetes are small proteins of about 40 to 100 amino acids (for examples, see Bo¨lker et al. 1992; Wendland et al. 1995; Vaillancourt et al. 1997; Halsall et al. 2000; Fowler et al. 2001; 2004; Lengeler et al. 2002; Gola and Kothe 2003b; Riquelme et al. 2005; Schirawski et al. 2005; Coelho et al. 2008; NiculitaHirzel et al. 2008; also see descriptions below). Analogously to the S. cerevisiae MFa precursor, the pheromone

107

precursors in basidiomycetes are predicted to undergo isoprenylation at the cysteine of the C-terminal CAAXmotif (C ¼ cysteine; A ¼ aliphatic residue; X ¼ any residue), then proteolytic cleavage of the motif AAX, and carboxylmethylation of the cysteine at the resulting C-terminus (Caldwell et al. 1995; Davey et al. 1998; Huyer et al. 2006). In some cases, such as C. neoformans the active pheromone has been synthesized and found to require these post-translational modifications for biological activity (Davidson et al. 2000). The farnesyl group might be further modified through alcohol groups (Caldwell et al. 1995; Davey et al. 1998; Huyer et al. 2006). At the N-terminus of the MFa precursor in the ascomycete S. cerevisiae, two successive processing events occur: the first involves N-terminal cleavage by the protease Ste24, and the second cleavage at the charged dipeptide preceding the actual pheromone peptide in the precursor, either by Axl1 or Ste23 (Caldwell et al. 1995; Chen et al. 1997; Davey et al. 1998; Huyer et al. 2006; see Chapter 5 in this volume). While similar N-terminal processing steps are hypothesized to occur in the basidiomycetes (Miyakawa et al. 1985a, b), not all pheromone precursors of basidiomycetes have a charged di-peptide preceding the pheromone peptide sequence. Others have monobasic processing sites or a dipeptide of which only the first residue is charged (e.g., see Fig. 6.4 and other figures with pheromone precursor sequences in this chapter). Actions by endopeptidases other than Axl1 and Ste23 are therefore hypothesized to participate in pheromone maturation in the basidiomycetes (Akada et al. 1989b).

Based on a plethora of earlier studies from other systems and organisms, the G protein-coupled 7-transmembrane pheromone receptors are integral membrane proteins localized on cellular membranes that span the membrane seven times with the N-terminus outside and the C-terminus inside the cell (Kothe et al. 2003; Fuchs et al. 2006; Xue et al. 2008). Through their extracellular N-terminus and/or extracellular loops, and possibly also involving the extracellular face of some transmembrane regions, these receptors bind compatible pheromones (Gola et al. 2000; Gola and Kothe 2003a; Naider and Becker 2004). Binding of a pheromone to a pheromone receptor is usually only efficient in inducing a subsequent mating reaction when the ligand and the receptor are products of nuclei of different mating type. This has been shown in various species by transformation studies of pheromone precursor and pheromone receptor genes into heterologous strains or, where possible, by gene deletion and replacement studies (see data e.g., in Bo¨lker et al. 1992; Wendland et al. 1995; Bakkeren and Kronstadt 1996; O’Shea et al. 1998; Davidson et al. 2000; Halsall et al. 2000; Fowler et al. 2001, 2004; Chung et al. 2002; Ku¨es et al. 2002b; Shen et al. 2002; Chang et al. 2003; Gola and Kothe 2003b; Riquelme et al. 2005; Schirawski et al. 2005;

Ursula Ku¨es et al.

108

A Rhodosporidium toruloides mating type A1 (a) RHA1.A1 RHA2.A1 RHA3.A1

MVAYPEISWRTNGCTVSKYPEISWRTNGCTVSKYPEISWRTNGCTVSKYPESIWTRNGCTVA MVAYPEISWTRNGCTVAKYPEISWTRNGCTVSKYPEISWTRNGCTVSKYPEISWTRNGCTVSKYPEISWTRNGCTVA MVAYPEISWTRNGCTVAKYPEISWTRNGCTVSKYPEISWTRNGCTVA

Rhodosporidium toruloides mating type A2 (A) RHA2.A2

MAPITLAANQEVAAAAPALVNEEHNPTVTSYMACTVSRKDVSVEMVDEEHNPTVTSYMACTIA

Sporidiobolus salmonicolor mating type A1 SsRHA1 SsRHA2 SsRHA3

MVAYPTIGVTTPGCTVSRTYPTIGVTTPGCTVSKTYPTIGVTTPGCTVSKTYPTIGVTTPGCIVSKTVA MVAYPTIGVTTPGCTVSRTYPTIGVTTPGCTVSKTYPTIGVTTPGCTVSRTYPTIGVTTPGCTVSKTYPTIGVTTPGCTVSKTYPTIGVTTPGCIVSKTVA MVAYPTIGVTTPGCTVSRTYPTIGVTTPGCTVSKTYPTIGVTTPGCTVA

Sporobolomyces spec. strain IAM 13481 (asexual?) SrRHA1 SrRHA2 SrRHA3

MAAYPYQGYALPGCTVSKTYPYQGYALPGCTVSKSYPYQGYALPGCTVSKSYPYQGYALPGCTVSKA MAAYPYQGYALPGCTVSKTYPYQGYALPGCTVSKSYPYQGYALPGCTVSKSYPYQGYALPGCTVSKA MAAYPYQGYALPGCTVSKTYPYQGYALPGCTVSKSYPYQGYALPGCTVSKSYPYQGYALPGCTVSKA

B Melampsora larici-populina MlpPh1 MlpPh2 MlpPh3 MlpPh4

MPPLNVDLTFVGTASSSESLLRNFGHQVGGSNHYCVIC MRRDLNTSSSHCRITSNCVRILFTAHIPGSLELIGKIPLRGVYKSCAREALLSAG.QVGGGTKYCIIS MSSSTNLKHENGSLSAIITLTSEREGQVGGGSRYCVISQRQGQV..GGGSKYCVVS..QREG.QVGGGSKYCVVS MSSATSQKHENGSPSAAIPLNSEREGQVGGGSRYCVISQRQGQV..GGGSKYCVIS..QRQG.QVGGGSKYCVVS

Fig. 6.4. Sequences for pheromone precursors as deduced (A) from cloned genes of members of the Sporidiobolales (after Coelho et al. 2008) and (B) from a larger gene family from the rust Melampsora larici-populina (Duplessis et al. 2011; UK, unpublished data). Peptide units for mature pheromones are shaded in grey and CAAX-motifs for Cterminal processing (Caldwell et al. 1995) are underlined.

Note that the residue N-terminal to the units of pheromone peptides is only in some of the cases a basic residue, and that there is in no case a charged dipeptide prior to the peptide units. Sporobolomyces spec. strain IAM 13481 refers to the yeast strain whose sequence has been published under the name Sporobolomyces roseus (http:// genome.jgi-psf.org/Sporo1/Sporo1.home.html)

Stanton et al. 2010). Binding of a pheromone stimulates conformational changes at the C-terminus of the pheromone receptor, which interacts with a heterotrimeric G protein. The activated G protein subunits then transmit the pheromone signal via the pheromone-activated signal transduction cascade (Regenfelder et al. 1997; Davidson et al. 2003; Hsueh et al. 2007; Li et al. 2007). The outcome of the activated pheromone signalling cascade is interconnected with regulation evoked by the HD1 and HD2 homeodomain transcription factors (Hartmann et al. 1996; Urban et al. 1996b; Kaffarnik et al. 2003; Lin et al. 2010; Stanton et al. 2010) and culminates ultimately in a transcriptional program in the nucleus that orchestrates the subsequent steps of sexual reproduction (Brachmann et al. 2001; Zarnack et al. 2008; Stajich et al. 2010).

arise first on autosomes and are then captured onto the sex chromosome (Ohno 1967). By comparison to certain Ascomycota where homeodomain transcription factor genes are also used to determine mating type (Casselton 2008; Dyer 2008, Lee et al. 2010), the HD locus is hypothesized to be the nascent “sex chromosome” in the basidiomycete fungi, and in an ancestral state the locus encoding the P/R genes lies on an autosome, yet to be captured onto the sex chromosome. How did this unlinked mating regulator first arise? One plausible model is that the P/R locus was originally auto-activated, either because the pheromone acted on the linked receptor gene product, or the receptor was constitutively active. This P/R locus evolved then to have more than one allele, and then recombination between two auto-activated alleles led to ones in which product of the linked pheromone gene only acts on the non-self receptor and not on self. A conceptually similar model was presented earlier (Fraser et al. 2007). While other models can certainly be envisioned or entertained, this one has several attractive features. First, a reversal of the steps in this model could lead to the collapse of a tetrapolar state back to bipolar, in which the P/R locus becomes autoactivated again and still participates in sexual

C. On the Origin of Tetrapolar Mating Type Determination Models for the origins of sex chromosomes provide a conceptual framework to understand how the tetrapolar mating system might have first evolved in the Basidiomycota. Sex determinants

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

reproduction but is no longer sex-determining (for further reading, see Section V.E.2). In multiple species, U. hordei, Malassezia globosa, C. neoformans and C. gattii, we can also observe that the unlinked P/R locus has been incorporated into the fungal sex chromosome, again leading to the collapse of an ancestral tetrapolar state to the extant bipolar one (Fraser et al. 2004; Lengeler et al. 2002; Bakkeren and Kronstad 1994; Lee et al. 1999; Xu et al. 2007; for details, see Sections IV and V.A). The key missing evidence here is to decipher gene function in extant bipolar species in which the HD locus governs mating type, and there is an unlinked P/R locus that is constitutively autoactivated and contributes to sexual reproduction but does not determine it. Coprinellus disseminatus (see Section V.E.2) is one such candidate worthy of further investigation that could emerge as a “missing link” in our genetic and molecular understanding of the key events that punctuated the bipolar-tetrapolar-bipolar evolutionary transitions (James et al. 2006).

III. Mating Type Genes in Pucciniomycotina A. Microbotryomycetes, Sporidiobolales Of all the basidiomycetes, the first molecular data on products of mating type genes were available for the saprotrophic red yeast Rhodosporidium toruloides, which possesses a biallelic one-locus mating type system with haploid cells of either the A1 (also known as A) or the A2 (also known as a) mating type (Banno 1967; Abe et al. 1975; Fell and Statzell-Tallmann 1998). A farnesylated peptide factor termed rhodotorucine A1 (or rhodotorucine A) has been isolated and characterized from A1 cells of this species (Kamiya et al. 1977, 1978a–c, 1979), and found to elicit conjugation tube formation in cells of the opposite mating type at nanomolar concentrations (Abe et al. 1975; Kamiya et al. 1979; Tsuchiya and Fukui 1978). Three structural genes (RHA1, RHA2, RHA3) encoding this pheromone were cloned and shown to be unique to the A1 mating type (Akada et al. 1987, 1989a, b). These genes encode pheromone precursors containing three to five

109

tandem repeats of the mature decapeptide pheromones (Fig. 6.4A) that are apparently released upon subsequent proteolytic processing of the precursor. Recently, a putative gene (RHA2.A2) encoding the rhodotorucine A2 pheromone was found in the alternate mating type A2. The deduced pheromone precursor produced from this A2 mating type specific gene contains two tandem repeats of a mature pheromone that differs in sequence from the A1 pheromone (Coelho et al. 2008; Fig. 6.4A). This study also identified a partial sequence for the pheromone receptor gene RtSTE3.A1 in the A1 mating type (for an analysis of the product, see Fig. 6.5) located 5’ to the rhodotorucine A1 pheromone precursor gene RHA1. Moreover for both of the alternate mating types, mating type specific genes encoding a Ste20-like p21-activated kinase (PAK) were also present. In the A1 mating type, the respective gene RtSTE20.A1 is flanked at the 5’ end by RHA3 and at the 3’ end by RHA2, respectively (Fig. 6.6). In the A2 mating type, the pheromone precursor gene RHA2.A2 is present 3’ to the RtSTE20.A2 gene encoding the Ste20-like PAK of the A2 mating type (Coelho et al. 2008). Within the Sporidiobolales, data for some other species are now also available. For the A1 mating type of the heterothallic yeast Sporidiobolus salmonicolor (strain IAM 12258), the sequences for three genes (SsRHA1, SsRHA2, SsRHA3) encoding pheromone precursors containing between four and six mature pheromone units (Fig. 6.4A) have been established. Incomplete sequences for one pheromone receptor gene (SsSTE3) and a STE20-like gene (SsSTE20) are also available. Similar to the gene arrangement in R. toruloides, the pheromone precursor gene SsRHA1 is linked to a partially sequenced pheromone receptor gene SsSTE3 whose encoded product most closely resembles RtSte3.A1 (Fig. 6.6). The SsRHA3 and SsRHA2 genes were found 5’ and 3’ of the SsSTE20 gene, respectively (Coelho et al. 2008; Fig. 6.6). Most recently, all of these genes in S. salmonicolor were shown to be linked together in an ~40 kb chromosomal region (Fig. 6.6) and the complete sequences of the pheromone receptor genes SsSTE3.A1 and SsSTE3.A2 of the two alternate mating types MATA1 and MATA2 were determined. The sequences of these gene products are considerably diverged (Fig. 6.5). Furthermore, the mating type genes

Ursula Ku¨es et al.

110

SjSte3.A1 SsSte3.A1 RtSte3.A1 SspSte3.A1 Mvpr-MATA1 RsSte3 99 CnnSte3a CngSte3a UhPra2 UmPrA2 99 SrPra2 93 LbSte3 LbSte3.5 CcSte3.2b LbSte3.2 CcSte3.2a

99 88 93 88 62

88 40 46 64

89

98 92 97 81 82 86 55

95

95 99 44 40 94 86

90 99

39 66 99 87

77

97

Tremellomycetes MATa Ustilaginomycotina a2 Agaricales B subgroup 2 Scbrl2 LbSte3.6 CcSte3-2153

92

95

Pucciniomycotina A1

LbSte3.1 CcSte3.1 ScBbr2 Scbrl1 LbSte3.9 CcSte3-7395 LbSte3.13 ScBar3 LbSte3.3 CcSte3.3 Scbrl3 LbSte3.4 CcSte3-2151 Scbrl4 TmSte3.A1 ChSte3.A1 CgSte3 99 CnnSte3a 80 CngSte3a MgSte3 CnnCpr2 UmPra1 SrPra1 UhPra1 SrPra3 ScSte3p PgSte3.1 89 PgSte3.3

Agaricales B subgroup 1

Agaricales B subgroup 3

Tremellomycetes

Ustilaginomycotina a1, a 3 Ascomycete

PgSte3.2 89 99 0.1

Mvpr-MATA2 SjSte3.A2 99 SsSte3.A2

Fig. 6.5. Phylogenetic analysis of pheromone receptor sequences from selected basidiomycetes. Sequences were taken from the complete genomes of Coprinopsis cinerea (Cc), Laccaria bicolor (Lb) and Schizophyllum commune (Sc) as previously described (Niculita-Hirzel et al. 2008; Ku¨es and Navarro-Gonza´lez 2010; Ohm et al. 2010; Stajich et al. 2010). Protein ID 28931 was taken from the JGI homepage http://genome.jgi-psf.org/Sporo1/Sporo1.home.html for the Sporidiobolus spec. strain IAM 13481 for pheromone receptor SspSTE3. PGTG_19559.2, PGTG_00333.2, and PGTG_01392.2 for the three putative Puccinia graminis (Pg) pheromone receptors were taken from the Broad homepage http://www.broadinstitute.org/annotation/ genome/puccinia_group/MultiHome.html. GenBank accession numbers for other proteins are EU401863

Pucciniomycotina A2

(partial sequence) for Rhodosporidium toruloides (Rt), ADM24789 and ADM24791 for Sporodiobolus johnsonii (Sj), ADM24772 and AMD24775 for S. salmonicolor (Ss), ABU62847 and ABU62846 for Microbotryum violaceum (Mv), AAA19965 and AAD56044 for Ustilago hordei (Uh), P31302 and P31303 for U. maydis (Um), CAI59749, CAI59755, and CAI59763 for Sporisorium reilianum (Sr), EDP44482 for Malassezia globosa (Mg), AAN75624, XP_570116 and AAW41941 for Cryptococcus neoformans var. neoformans (Cnn), AAN75156 and AAN75157 for C. neoformans var. grubii (Cng), AAV28793 for C. gattii (Cg), ACZ814463 for C. heveanensis (Ch), ABL67527 for Rhizoctonia solani (Rs), and, as an outgroup from the ascomycetes, EEU04250 for Ste3 (ScSte3p) from Saccharomyces cerevisiae (Sc). For protein comparisons, sequences

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

111

Rhodosporidium toruloides mating type A1

Sporidiobolus salmonicolor mating type A1 HD1/HD2

Sporidiobolus spec. strain IAM 13481 bE1/bW1 ~240

~27

~100

~220

~620 -1200

~60

~10

~220

Fig. 6.6. Genomic organization of DNA regions with mating type genes in the Sporidiobolales (after Coehlo et al. 2008, 2010). Mating type genes for HD1 and HD2 transcription factors are marked in blue and light blue, respectively (note that the relative orders of position of HD1/HD2 gene pairs in the species are not yet known) and mating type genes for pheromone precursors (small boxes) and pheromone receptors (large boxes) in red. Genes for products required in addition for the mating pathway

downstream to the pheromone-pheromone receptor system are shown in green. Neighbouring genes in close neighbourhood of confirmed synteny are shown in light grey, neighbouring genes for which synteny has not been analysed are depicted in white and genes linked more distantly from the mating type genes in black. Arrows indicate direction of transcription where known. Numbers beneath a map indicate distances between genes in kilobases

for pheromone receptors were found to be linked to a pair of HD1 and HD2 genes (for the homeodomains of the products, see Fig. 6.1) over an apparently large distance of several hundred kilobases (Fig. 6.6). Surprisingly, there are multiple HD1/HD2 gene pair alleles in nature (13 detected so far) of which one subset occurs in a worldwide strain collection invariably linked to SsSTE3.A1 of the A1 mating type and the other subset is linked to the SsSTE3.A2 of the A2 mating type. The sequences of the multiple HD1/HD2 gene pair alleles within and between subsets are however highly diverged in sequence suggesting that their products do not confer the same protein–protein dimerization specificity. Although the species S. salmonicolor is described as bipolar, and recombination is partially suppressed in the region between the different types of mating type

genes, laboratory experiments show that recombination can infrequently occur between the HD1/ HD2 gene pair and the SsSTE3 alleles. Delayed germination of recombinants is suggested as one possible explanation why the resulting MAT recombinant progeny produced by laboratory sexual reproduction appear to be counterselected in nature. Crosses between a recombinant and its parental strains in the laboratory revealed that both the HD1/HD2 and the SsSTE3 loci have mating type function, regulating different steps in mating. When partners in crosses differed only in the HD1/HD2 gene pairs, no reaction was seen while partners differing only in the pheromone receptor alleles formed limited pseudohyphae or conjugation tubes (Coelho et al. 2010). To describe the distinction between the purely bipolar mating behaviour of the S. salmonicolor

ä were C-terminally truncated after the seven-transmembrane regions, aligned using first ClustalX (http://wwwigbmc.u-strasbg.fr/BioInfo/ClustalX/Top.html) and then GeneDoc version 2.6.2002 (http://www.psc.edu/biomed/ genedoc/) for manual adjustments and phylogenetic trees were calculated by the neighbour joining method of MEGA version 4.0 (Tamura et al. 2007). Bootstrap values (500 replications) are shown and the bar defines the number of nucleotide substitutions per site. Clusters of pheromone receptors are underlined with a colour code to indicate the

phylogenetic positions of their organisms (coral Microbotryomycetes, orange Pucciniomycetes, green Ustilaginomycotina, pink Tremellomycetes, light blue Agaricomycetes). Clusters with pheromone receptors from mating type loci are marked by brackets and the names of their respective phylogenetic group. Unmarked pheromone receptors come either from genes that are located outside of mating type loci (C. neoformans var. neoformans, C. cinerea, L. bicolor, S. commune) or where a mating type functions remains still to be shown (P. graminis, R. solani)

112

Ursula Ku¨es et al.

strains isolated from nature and the tetrapolarity of inheritance of mating type genes generating four different mating types in the laboratory, this novel mating system has been named pseudobipolar (Coelho et al. 2010). Pseudo-bipolarity appears to be common in the heterothallic Sporidiobolales. Coelho et al. (2010) further detected in the red yeasts Sporodiobolus johnsonii and Rhodosporidium babjevae a system with multiple alleles of an HD1/HD2 gene pair (for the available homeodomain sequences, see Fig. 6.1) linked on the same chromosome in an A1- and A2-mating type specific manner to either one of two alternative pheromone receptor genes (see Figs. 6.5 and 6.7 for analyses of the mating type gene products). Most interestingly, the S. johnsonii type strain of the species (CBS 5470) is homothallic and forms a dikaryotic mycelium from a diploid yeast phase although it does not form meiotic basidia upon germination of teliospores (Fell and Statzell Tallman 1981; Coehlo et al. 2010). Inspection of the mating type genes indicates that the type strain carries both alleles for the alternate pheromone receptors as well as an HD1/HD2 gene pair. It has been suggested by Coehlo et al. (2010) that this gene pair might encode self-compatible HD1 and HD2 transcription factors due to a gene conversion event involving allelic HD1/HD2 gene pairs. Another strain of S. johnsonii (CBS 1522) is asexual (anamorphic) but has the same pair of HD1 and HD2 genes and the pheromone receptor gene SjSTE3.A1 found in three analysed heterothallic strains of the species. Furthermore, from the study of Coehlo et al. (2010) it is apparent that diverging HD1 and HD2 gene pair alleles are not reciprocally monophyletic between the very closely related species S. salmonicolor and S. johnsonii, which are able to form cross-species dikaryons and viable teliospores (Valerio et al. 2008b). Nevertheless, most extant HD1 and HD2 allele diversity seems to have been generated after speciation (Coehlo et al. 2010), and this is distinct from the evolutionary trajectory of the pheromone receptor alleles that have an ancient transspecies sequence polymorphism (Fig. 6.5). Analysis of nine HD1 alleles from S. salmonicolor and S. johnsonii indicate exceptionally high evolutionary rates, indicative of positive selection. Moreover, genes neighboring the mating type genes also exhibit degrees of allele sequence divergence and in one case (LSm7 located proximal to SsSTE3.A1; Fig. 6.6) is mating type specific. For

other genes located in the larger chromosomal area with the mating type genes, frequent past and more recent gene conversion events are evident, suggesting gene conversion is an important mechanism operating on genes in the neighbourhood of the mating type locus (Coelho et al. 2010). Self-fertile and asexual strains can be found among natural isolates of many Sporidiobolales and it is not straightforward to assign them to distinct species due to their uncertain phenotypic characters (Fell and Statzell Tallman 1981; Bai et al. 2002; Vale´rio et al. 2008a, b). Some species were reported to be asexual, e.g., Sporobolomyces roseus. However, self-fertile strains conspecific to the S. roseus type strain CBS 486 and other anamorphic isolates have recently been described under the teleomorphic name Sporidiobolus metaroseus sp. nov. (Vale´rio et al. 2008a). The DOE (US Department of Energy) Joint Genome Institute (JGI) sequenced the genome of the purportedly asexual red yeast strain IAM 13481 labelled as S. roseus (http://genome.jgi-psf.org/Sporo1/Sporo1.home. html) but the species identity is uncertain. Blastn searches with the sequences of the ITS regions (internal transcribed spacer region of the nuclear ribosomal DNA) and the D1/D2 domains of the large-subunit rDNA sequences (LSU) of the S. roseus type strain CBS 486 [GenBank accession numbers AB030351 and EU003462 at the National Center for Biotechnology Information (NCBI) http://www.ncbi.nlm.nih.gov/] against the raw genomic sequences of Sporobolomyces spec. strain IAM 13481 (deposited in the NCBI Trace Archive; http://www.ncbi. nlm.nih.gov/Traces/trace.cgi?) revealed relatedness between the strains but not identical ITS sequences, indicating that Sporobolomyces spec. strain IAM 13481 does not represent an S. roseus strain. The raw genome sequence data of Sporobolomyces spec. strain IAM 13481 contains about 250 sequences that show highest identity to the ITS sequences (97% identity) of Sporidiobolus pararoseus strain CBS 484 (GenBank AB030338) and about 250 sequences that show highest identity to the sequences of the LSU D1/D1 domains (99% identity) of the S. pararoseus type strain CBS 491 (GenBank AF189977). Sporobolomyces spec. strain IAM 13481 is thus closely related but possibly does not belong to the species S. pararoseus. S. pararoseus is reported to be a sexually reproducing yeast with the two mating types A1 and A2 (Bai et al. 2002; Vale´rio et al. 2008a). Also some isolates of S. pararoseus are self-fertile or asexual (Vale´rio et al. 2008a). In terms of mating type genes, these S. pararoseus isolates might behave in similar ways as the self-fertile and the asexual strains of S. johnsonii described by Coelho et al. (2010). In conclusion, for Sporobolomyces spec. strain IAM 13481, the species identity remains to be identified as well as whether this species also consists of self-sterile heterothallic, self-fertile homothallic, and asexual individuals.

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

Upon establishing the complete genome of Sporobolomyces spec. strain IAM 13481, mating type like genes were also found in this asexual strain (Fig. 6.6). Three related pheromone precursor genes encoding proteins with three tandem repeats of pheromone peptide motifs (Fig. 6.4A), a gene for a putative pheromone receptor (Fig. 6.5), and one complete HD1/HD2 gene pair (Figs. 6.1 and 6.7) were identified (Coelho et al. 2008; Morrow and Fraser 2009; http://genome.jgi-psf. org/Sporo1/Sporo1.home.html). The newly detected HD1 and HD2 genes were named bE (for bEast) and bW (for bWest) (http://genome.jgi-psf.org/Sporo1/ Sporo1.home.html), by analogy to the genes in the respective mating type locus in U. maydis (the b locus; Gillissen et al. 1992) and other Ustilaginomycotina (see Section IV.A). Together with the SrSTE20 gene of Sporobolomyces spec. strain IAM 13481, the three pheromone precursor genes (SrRHA1, SrRHA2, SrRHA3) and the pheromone receptor gene (SrSTE3) are found in a region of ~25 kb (on scaffold 9 of the published genome; http:// genome.jgi-psf.org/Sporo1/Sporo1.home.html), in a relative gene order similar to that in the heterothallic species R. toruloides and S. salmonicolor

a

113

(Coelho et al. 2008, 2010; Fig. 6.6). The gene arrangements in all three species suggest that in many of the Sporidiobolales a larger chromosomal region exists covering numerous distinct genes acting in the process of mating, including the typical mating type genes. For R. toruloides, the data currently available is insufficient to conclusively prove this hypothesis. However, the HD1/HD2 gene pair in Sporobolomyces spec. strain IAM 13481 is located on another scaffold in the genome assembly (scaffold 7; http:// genome.jgi-psf.org/Sporo1/Sporo1.home.html), which defines a minimum distance of ~620 kb to 1.2 Mb between the HD1/HD2 gene pair and the pheromone and pheromone receptor genes, assuming that they are located on the same chromosome (Coelho et al. 2008, 2010). A more thorough analysis of gene content and order in S. salmonicolor not only confirmed linkage of the two regions for S. salmonicolor but suggests considerable synteny between the two yeasts with several of the known genes being conserved in position (Coehlo et al. 2010; Fig. 6.6). Linkage of the two MAT loci in Sporobolomyces spec. strain IAM 13481 remains to be experimentally tested and validated.

b

0.2

Fig. 6.7. Phylogenetic analysis of the sequences of homeodomains from HD1 (A) and HD2 (B) homeodomain transcription factors of selected basidiomycetes. Sequences were obtained as depicted in Fig. 6.1 (for species codes see legend of Fig. 6.1) and aligned using ClustalW (http://wwwigbmc.u-strasbg.fr/BioInfo/ClustalX/Top.html) and a tree was constructed by the neighbour joining method of the

0.1

MEGA ver.4.0 program (Tamura et al. 2007). The bar defines the number of nucleotide substitutions per site. Clusters of pheromone receptors are underlined with a colour code to indicate the phylogenetic positions of their organisms (coral Microbotryomycetes, orange Pucciniomycetes, green Ustilaginomycotina, pink Tremellomycetes, light blue Agaricomycetes)

114

Ursula Ku¨es et al.

B. Microbotryomycetes, Microbotryales Within the Microbotryales, transcriptional profile analysis of the anther smut fungus Microbotryum violaceum identified highly divergent mating type specific pheromone receptor genes (pr-MATA1, pr-MATA2) for the alternate mating types A1 and A2 (Yockteng et al. 2007; Devier et al. 2009; Fig. 6.5). Related pairs of allelic pheromone receptor genes have subsequently been found in the alternate mating types of 12 other members of the Microbotryum species complex. The products of all of these genes clearly split into two distinct clades of pheromone receptors, each clade corresponding to one allele per species. Accordingly, the Microbotryum species complex has an ancestral state of two alternate mating types (Devier et al. 2009, 2010). Interestingly in M. violaceum, intratetrad mating (automixis) is favored: Teliospores germinate into a four-celled promycelium but produce only one type of viable sporidia (more often of the A1 mating type than of the A2 mating type) due to a high level of haplolethality genetically linked to the mating type locus. Haplolethality of sporidia however does not hinder conjugation of promycelium cells allowing intratetrad mating. In meiotic crosses, the single mating type locus is tightly linked to the centromere of chromosome VI due to suppressed recombination ( although in mitotic divisions recombination may occur). Because meiotic recombination in M. violaceum is generally low, this in combination with the high level of haplolethality ensures a high degree of heterozygosity in natural isolates of the species (Zakharov 2005).

The sequence of the complete mating type locus in M. violaceum is not yet fully established. It is known that the mating type genes are located on chromosomes of different size and structure (“sex chromosomes”) that diverged during the evolution of the A1 and the A2 mating types (Devier et al. 2009; see Chapter 12 in this volume). These dimorphic sex chromosomes are 2.8–3.1 Mb (A1) and 3.4–4.2 Mb (A2) (data for isolates from the plant Silene latifolia). The sex chromosomes are characterized by a low coding gene density, by an accumulation of transposons and repetitive elements, and by a repression of recombination over much of their length (Hood 2002, 2005; Hood et al. 2004). The non-recombining region is estimated to span 1.0–2.6 Mb (Hood and Antonovics 2004; Votinseva and Filatov 2009). Recently, Abbate and Hood (2010) demonstrated that the composition of the non-recombining region of the Microbotryum sex

chromosome appears to be expanding during the evolution of the genus, with one locus (MTL1) being captured by the recombination block multiple times independently. Furthermore, Giraud et al. (2008) reported in a minireview on the mating system of M. violaceum that sequences for HD1 and HD2 homeodomain transcription factors were detected in an EST library. Sequence AEIJ01001054 within the recently published M. violaceum genome (GenBank AEIJ01000000) appears to carry the divergently transcribed HD1/HD2 gene pair. In addition, there are sequences (AEIJ01000114, AEIJ01000884, AEIJ01000869) that appear to carry genes for pheromone precursors with tandem repeats of mature decapeptide pheromones (UK, unpublished data).

C. Pucciniomycetes, Pucciniales Rusts include plant pathogenic species with complex life cycles. The biotrophic rusts form five different types of spores on two different hosts (Buller 1950; Esser 2000; Spiers and Hopcroft 1994; Yu et al. 2009), posing challenges for the analysis of their mating system. The pioneering work by Craigie (1927, 1931) suggests that the wheat stem rust P. graminis and some related Puccinia species have two alternate mating types (known as + and –). Transfer experiments within and between species of nectar secreted by pycnidia (conidiomata forming the single-celled haploid pycniospores that act as spermatia) from one strain onto pycniospores of others induced spore cap formation on the foreign pycniospores in a mating type specific manner. These observations may confirm for a number of Puccinia and also Uromyces species the existence of a biallelic single mating type locus (Anikster et al. 1999, 2000). Mating tests on Gymnosporangium asiaticum using single-basidiospore cultures revealed a bipolar mating system with two different mating types (Tetsuka and Katsuya 1984). However, three different rusts (Cronartium quercuum, Melampsora lini, and Puccinia coronata) have so far been reported to be tetrapolar, indicating that two unlinked loci control mating (Lawrence 1980; Yamazaki and Katsuya 1988; Narisawa et al. 1994). Whether these loci are bi- or multi-allelic is not yet known and molecular data for these three species are not available. Various other species have been reported from pycniospore transfer experiments to be heterothallic (Allen 1934; Buller 1950), but the underlying genetic basis remains unclear. Insight into the genetic structure of mating type genes in the order Pucciniales is however

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

now arising from fungal genome sequencing initiatives in progress at the Broad Institute (Cambridge, Mass., USA) and at the JGI (Walnut Creek, Calif., USA). The complete genomes from the wheat stem rust P. graminis f. sp. tritici and the wheat leaf rust Puccinia triticina are available (http://www.broadinstitute.org/annotation/genome/ puccinia_group/MultiHome.html) as is the genome from the poplar rust M. larici-populina (http:// genome.jgi-psf.org/Mellp1/Mellp1.home.html). The latter species, deduced from gene flow data obtained from population studies using neutral genetic markers (Gerard et al. 2006; Pei et al. 2007; Barres et al. 2008), is thought to be heterothallic although it remains unknown whether bior tetrapolar and whether bi- or multi-allelic. An inspection of the genomes reveals all three species harbor a typical HD1/HD2 gene pair locus (for the homeodomain sequences of their products, see Fig. 6.1), and also presumptive pheromone receptor genes (Fig. 6.5) and genes for pheromone precursors with up to four tandem repeats of potential pheromone peptides (for some examples from M. larici-populina, see Fig. 6.4). Particularly the pheromone receptor genes reside in regions that are generally low in gene density and rich in transposon insertions. However, the available data do not yet allow the assembly of these different regions into one or into different chromosomes which would help to define the complete nature of mating type loci in the rusts (Duplessis et al. 2011; UK, unpublished data). It is thus unclear whether, for example, the three predicted pheromone receptor genes in P. graminis (Devier et al. 2009) all have mating type function. A full account on the mating type genes in the three sequenced species will be presented in the near future in publications prepared by members of the consortia of rust genome researchers (Duplessis et al. 2011).

D. Evolutionary Considerations The emerging data on (potential) mating type genes in species of the Pucciniomycotina suggest that the basic concept of mating type genes represented by homeodomain transcription factors and of genes for pheromone precursors and genes for pheromone receptors is also conserved in this subphylum of the Basidiomycota – as described previously in the Ustilaginomycotina and in the

115

Agaricomycotina, respectively (see Sections IV and V). Having sampled this most-basal lineage of basidiomycetes (Liu et al. 2009), we can now conclude that the occurrence of all known types of genes with mating type function (HD and P/R) is quite ancient and therefore appears to have emerged possibly once in the most recent common ancestor of the Basidiomycota, although multiple independent origins are still formally possible. Moreover, the presence of all types of genes mediating mating type function in the basidiomycetes is shared with various ascomycetous yeasts exemplified by the model species S. cerevisiae. However, compared to the Basidiomycota a major difference is that the genes for the pheromone and pheromone receptor system in the Ascomycota are not encoded by the MAT locus but instead their expression is regulated in a cell type specific manner by mating-type specific transcription factors (Herskowitz 1988, 1989; see Chapter 5 in this volume). Mating type loci of fungi other than the basidiomycetes also contain genes for transcription factors but these are in many instances of a different type than homeodomain transcription factors (Coppin et al. 1997; Kronstad and Staben 1997; Idnurm et al. 2008; Lee et al. 2010a, b; Gryganskyi et al. 2010; see Chapters 5 and 12 in this volume). By comparing the kinds of transcription factor genes (HMG box transcription factors, a-domain proteins, and homeodomain transcription factors, respectively) residing at mating type loci within different fungal phyla and subphyla [including the basal phyla Chytridiomycota, Glomeromycota, and the subphylum Mucoromycotina ¼ former Zygomycota (Hibbett et al. 2007) and the Dikarya], Dyer (2008) suggested an independent gain of mating type genes for homeodomain transcription factors in the basidiomycete lineage and in the Saccharomycotina of the ascomycete lineage, respectively. In contrast, Casselton (2008) and Lee et al. (2010) argued that HD1 and HD2 homeodomain transcription factor genes were present in the ancestor of the Dikarya based on the presence of these genes in basal lineages of both phyla of the Dikarya (in the Saccharomycotina and in the Pucciniomycotina). In this model, genes for homeodomain transcription factors must have been lost from the MAT loci in the subphyla Pezizomycotina and Taphrinomycotina of the Ascomycota.

In the baker’s yeast S. cerevisiae, in haploid MATa cells the genes for the a-pheromone receptor Ste2 and for the a-pheromone MFa are constitutively expressed. In haploid in MATa cells, transcription factors encoded by the a allele of the mating type locus control expression of a-specific genes (Herskowitz et al. 1988, 1989).

116

Ursula Ku¨es et al.

The a1-protein (belonging to a unique class of transcription factors characterized by the a1domain; Coppin et al. 1997) activates a-specific genes, including the a-pheromone receptor Ste3 and the a-pheromone MFa (also known as a-factor) while the HD1 homeodomain transcription factor a2 represses the expression of a-specific genes. After cell–cell fusion during mating, in the diploid zygote the HD1 and HD2 homeodomain transcription factors a2 and a1 (from the a allele of the mating type locus) dimerize and repress all haploid-specific genes and the mating type gene a1 while a2 continues to repress a-specific genes (Herskowitz et al. 1988, 1989; Stanton and Hull 2007). With respect to the physiological nature of mating types in basidiomycetes it is interesting to note that the Ste2 pheromone receptor of S. cerevisiae derives from a different clade of G-protein coupled receptors than Ste3 (Josefsson 1999; Graul and Sade´e 2001) and that the hydrophilic MFa pheromones are unrelated to the lipophilic MFa lipopeptides and have a completely different structure (Davey et al. 1998). Similarity searches in the genomes of the Pucciniomycotina (as also in all the available genomes of the Ustilaginomycotina and the Agaricomycotina) did not reveal any STE2-related genes nor any genes related to the two S. cerevisiae a-factor genes MFa1 and MFa2 (UK, unpublished data). Apparently, orthologues to STE2 and orthologues to MFa1 and MFa2 do not exist in any of the basidiomycetes that have been analysed at a molecular level. In a comparative analysis of G protein-coupled 7 transmembrane pheromone receptors from the Pucciniomycotina and from other basidiomycetes together with the two different yeast receptors Ste2 and Ste3, Devier et al. (2009) confirmed that all mating type receptors from basidiomycetes are evolutionary related to S. cerevisiae Ste3 (see also Fig. 6.5). An important observation regarding the Pucciniomycotina Ste3 alleles based on the alternate pheromone receptor sequences available from M. violaceum, S. salmonicolor and S. johnsonii has been the identification of two separate transspecific lineages of Ste3-like pheromone receptors (Devier et al. 2009; Coehlo et al. 2010). One of these lineages groups with the three predicted pheromone receptors from the one sequenced mating type of P. graminis and also with that from the Sporobolomyces spec. strain IAM 13481, in a clade unrelated to the pheromone receptors from the Ustilaginomycotina and the

Agaricomycotina. In contrast, the other lineage clusters with pheromone receptors from the Ustilaginomycotina and the Agaricomycotina with a strong bootstrap support (Fig. 6.5). Apparently, the genes for these two different lineages of pheromone receptors found in the Pucciniomycotina (A1 and A2) have been maintained as alleles for over 370 million years of evolution (Devier et al. 2009; see below) but only the A1 lineage has been conserved in the more derived subphyla of the Basidiomycota. Another interesting evolutionary development in the Pucciniomycotina concerns the structure of the pheromone precursor genes, specifically the structure of their primary products. Thus far, only the Pucciniomycotina appear to have evolved genes for pheromone precursors that contain (several) tandem repeats of pheromone peptide units. Precedents for pheromone precursors containing tandem repeats of pheromone units (four in total in the case of MFa1, two in the case of MFa2) that are released by enzymatic processing of precursors are found in S. cerevisiae a cells (Kurjan and Herskowitz 1982; Davey et al. 1998) and other ascomycetous yeasts (Imai et al. 1994; Panwar et al. 2003; Ongay-Larios et al. 2007; see Chapter 5 in this volume). The maturation of ascomycete MFa-type pheromone precursors for mature hydrophilic pheromones, however, involves an ordered set of events of signal sequence cleavage, glycosylation, and proteolytic processing by three different peptidases that are distinct from the events involved in processing the MFa precursors (Fuller et al. 1988; Davey et al. 1998).

IV. Mating Type Genes in Ustilaginomycotina A. Ustilaginomycetes, Ustilaginales The maize smut U. maydis is a well studied model species in the basidiomycetes with respect to mating type genes and their function (Kahmann and Ka¨mper 2005). The species is tetrapolar and the a mating type locus (P/R) has two alleles and the unlinked b mating type locus (HD) has ~25 alleles (Puhalla 1970; Silva 1972; Banuett and Herskowitz 1989). This type of mating system has been termed modified tetrapolar [Wells and Wong 1985 (or modified bifactorial in their verbage)] because it differs in terms of outbreeding potential from the

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

Ustilago maydis

Sporisorium reilianum

b1

a1

b2

a2

b1

a1

b2

a2

117

*

≈

Malassezia globosa

*

≈

≈

~ 500 kb ~ 430 kb

MAT-1

≈

Ustilago hordei

≈

a3

*

MAT-2

MAT

~ 170 kb

Fig. 6.8. Chromosomal organization of mating type loci of Ustilaginomycetes (after Gillissen et al. 1992; Urban et al. 1996; Bakkeren and Kronstad 1993, 1996; Schirawski et al. 2005; Bakkeren et al. 2006, 2008; Xu et al. 2007). Mating type genes for HD1 and HD2 transcription factors are marked in blue and light blue, respectively and mating type genes for pheromone precursors (small boxes) and pheromone recep-

tors (large boxes) in red, with a pheromone precursor pseudogene being shaded in grey. Genes found in mating type loci that lack known mating function are in light grey. Arrows indicate direction of transcription, numbers beneath a map indicate distances between genes in kilobases, and * indicates genes with unknown orientation and order

two multi-allelic mating type loci (see Section IV. C) well known from the Agaricomycotina (see Section V.E.1). The biallelic a locus of U. maydis (Fig. 6.8) encodes alternate variants of pheromone precursors (Figs. 6.9 and 6.10) and pheromone receptors (Fig. 6.5), respectively. Each of the a alleles has one active pheromone precursor gene (mfa1 or mfa2) and one active pheromone receptor gene (pra1 or pra2) but in the a2 allele there is an additional non-functional second pheromone gene (a mfa1related mfa pseudogene missing a startcodon and with a premature stop codon at the position where a codon for the cysteine of the CAAX motif is to be expected). Furthermore in the a2 allele, there are two unique genes, lga2 and rga2 (Bo¨lker et al. 1992; Spellig et al. 1994; Urban et al. 1996a) that direct uniparental inheritance of mitochondria thus inhibiting mtDNA recombination during sexual development (Fedler et al. 2009; Mahlert et al. 2009).

1991). The term idiomorph refers to alternative versions of the mating type locus on homologous chromosomes that are completely dissimilar in sequence and that encode unrelated proteins (Metzenberg and Glass 1990; see Chapter 12 in this volume). However, even though the a mating type locus in U. maydis and related species are very divergent in sequence, at least the pheromone receptor genes do share a common origin (Fig. 6.5). The cryptic pheromone gene in the 8 kb a2 locus is clearly related to the active gene in the a1 locus and there is a second block of sequence similarity between the non-coding regions of the two loci (Urban et al. 1996a). Thus, the U. maydis a mating type locus only partially fulfills the classic criteria of an idiomorph as defined by Metzenberg and Glass (1990). The two alleles of the a mating type locus of U. maydis clearly have a common origin, but during evolution these greatly diverged in sequence in addition to smaller rearrangements that occurred, and lga2 and rga2 have been incorporated into the a2 haplotype or lost from the a1 (Urban et al. 1996a).

The a mating type locus of U. maydis has been suggested to have a true idiomorphic nature (Froeliger and Leong

Compared to U. maydis, the closely related tetrapolar species Sporosorium reilianum (ITS sequences placed U. maydis and S. reilianum both into the genus Sporosorium; Piepenbring et al. 1992; Stoll et al. 2005) has three alleles of the a locus (Fig. 6.8). Each locus has one gene

118

Ursula Ku¨es et al.

Fig. 6.9. Sequences for pheromone precursors as deduced from cloned genes of members of the Ustilaginomycotina (after Bo¨lker et al. 1992; Spellig et al. 1994; Anderson et al. 1999; Kosted et al. 2000; Schirawski et al. 2005; Xu et al. 2007). Peptide units for (predicted) mature pheromones are shaded in grey and CAAX-motifs for C-terminal processing (Caldwell et al. 1995) are underlined. Note that the residue N-terminal to the pheromone peptide units is only in some of the cases a charged residue

for a unique pheromone receptor (Fig. 6.5) and two genes for pheromone precursors (Figs. 6.9 and 6.10). For each a mating type allele, one of the two encoded mature pheromones interacts specifically with only the pheromone receptor of one of the other two alleles while the other of the two pheromones interacts specifically with the pheromone receptor of the third allele (Fig. 6.10). In this way, a system of three different mating type specificities with complete intercompatibility is established (Schirawski et al. 2005). The pheromone receptors from Ustilaginales are found in two locations on the tree with the Pra1 receptors forming a clade separate from the Pra2 receptors (Fig. 6.5). Within each of these two clades, the expected species relationship is observed, i.e., U. maydis and S. reilianum are more closely related to each other than they are to U. hordei (Begerow et al. 2006). Thus, the two receptor alleles pra1 and pra2 have been inherited as a pair of compatible alleles through the speciation of the group, in a trans-specific pattern of evolution first described by Devier et al. (2009). The Pra3 receptor of S. reilianum groups with Pra1 receptors of Ustilaginales with significant support (Fig. 6.5). The origin of this third allele and the uniqueness of the S. reilianum mating

system awaits further clarification. Schirawski et al. (2005) put forward two hypotheses: (1) the a3 allele had been acquired by horizontal transfer from a related smut species (by interspecific hybridization), or (2) the a3 allele has been lost recently from U. maydis, possibly supported by the putative pheromone pseudogene (mfa) found in the U. maydis a2 locus (Fig. 6.8). Although the alleles of the a mating type loci of S. reilianum have an extra pheromone gene, the a1 and a2 loci of S. reilianum are otherwise syntenic to the a1 and a2 mating type loci of U. maydis, respectively, with respect to gene content, order, position, and genomic context (Schirawski et al. 2005; Bakkeren et al. 2008; Fig. 6.8). Other than an uncertain origin of the S. reilianum gene pra3, the pheromone precursor genes in the a3 locus of the species appear to have arisen from recombination between the a1 and a2 alleles of the species. Notably, the extra functional pheromone gene mfa2.3 in the a2 locus of S. reilianum is located at the same position as the mfa-pseudogene of the U. maydis a2 locus and might descend from a common ancestor with this pseudogene or may have been translocated to the same location as the U. maydis mfa-pseudogene (Schirawski et al. 2005). Alternatively, U. maydis

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

may have had a third mating type allele that has been lost from the population and the mfapseudogene became non-functional as divergence occurred from an ancestral allele with two active pheromone genes. In this later model, the ancestral species was tetrapolar and multi-allelic at both the HD and P/R loci, and U. maydis has lost allelic diversity at the P/R locus to become biallelic whereas S. reilianum has retained three extant P/R locus alleles and is therefore triallelic. Clearly, either evolutionary scenario is conceptually possible, and the means to distinguish these models will be to sequence additional closely related species to this species complex to examine their patterns of sexuality and mating type locus configurations and diversity. Strikingly, even though the homologous genes for the pheromone precursors that are found in the different mating type loci in S. reilianum have in some cases slightly different sequences, the mature pheromones do not differ in sequence (compare in Fig. 6.9 the predicted sequences of the mature pheromones with identical mating type specificity: SrMfa1.2 and SrMfa3.2; SrMfa2.1 and SrMfa3.1; SrMfa1.3 and SrMfa2.3). The sharing of identical mature pheromone sequences

119

explains how two mature pheromones from different mating type alleles can interact with the pheromone receptor of the third allele (Schirawski et al. 2005; Fig. 6.10). Regarding mating type genes for HD1 and HD2 homeodomain transcription factors, the multi-allelic b mating type locus of U. maydis encoding the bE and bW genes exhibits the hallmark mating type locus structure with a single divergently transcribed pair of HD1 and HD2 genes (Fig. 6.8). In this species, sequences Nterminal to the homeodomain region in the allelic bE- and bW-proteins are highly variable whereas the homeodomains and the C-terminal parts of the two protein classes are highly conserved (Kronstad and Leong 1990; Schulz et al. 1990; Gillissen et al. 1992). Similarly, divergently transcribed loci with HD1 and HD2 mating type genes exist in the tetrapolar S. reilianum where at least five allelic versions of the b mating type locus have HD1 and HD2 products characterized by a variable N-terminal dimerization domain and highly conserved C-terminal sequences (Schirawski et al. 2005; Fig. 6.8). Moreover, divergently transcribed loci with HD1 and HD2 mating type genes are also found in bipolar smuts such as in the barley smut

Sporisorium reilianum

Ustilago maydis

a1b1 Mfa1

Mfa1.2

Mfa1.3

Pra1 Pra2

a1b1

a2b2

Pra1

13aa

Mfa2.1

a2b2 Mfa2

a1 a2

14aa 9aa

Pra2

Mfa3.1

Pra3

a3b3

Mfa3.2

Mfa2.3

a1

X

X a2 a3

Fig. 6.10. Pheromone and pheromone receptor interactions between cells of different mating type in Ustilago maydis (a1b1 and a2b2) and in Sporosorium reilianum (a1b1, a2b2, and a3b3). Pheromones are indicated by filled squares, triangles and circles, pheromone receptors by

X X open cup-shaped symbols. Identical colour coding indicates compatibility between a pheromone and a receptor. The interactions between the products of pairs of different a mating type alleles are further illustrated at the bottom of the figure

120

Ursula Ku¨es et al.

U. hordei (Bakkeren and Kronstad 1993) and in the sugarcane smut Sporisorium scitamineum (Albert and Schenk 1996). In these bipolar species, the sequences of the N-terminal dimerization domains are also diverged unlike the sequences of the remaining C-terminal parts of the proteins (Bakkeren and Kronstad 1993; Fig. 6.8). Phylogenetic comparisons of HD1 and HD2 proteins from the closely related S. reilianum, U. hordei and U. maydis (Piepenbring et al. 2002; Stoll et al. 2003) suggest that allele diversification happened after speciation (Schirawski et al. 2005). In the bipolar species U. hordei, the b mating type genes are linked on the same chromosome into a single heritable unit (MAT locus) to the a mating type genes at distances of about 500 kb (MAT-1) and 430 kb (MAT-2), respectively (Bakkeren and Kronstad 1994; Lee et al. 1999; Fig. 6.8). In U. hordei in which there are two alternate mating types MAT-1 and MAT-2, there are only two alleles of the bE and bW gene pair for HD1 and HD2 transcription factors distinguished by sequences of the respective N-terminal dimerization domains (Bakkeren and Kronstad 1993). Furthermore in U. hordei, there are also only two alleles of a mating type genes that encode the alternate pheromone precursors UhMfa1 or UhMfa2 and the alternate pheromone receptors UhPra1 and UhPra2 (Bakkeren and Kronstad 1993, 1996; Anderson et al. 1999; Kosted et al. 2000; Figs. 6.5 and 6.9). In the large chromosomal region that lies between the a and b mating type genes in U. hordei (Fig. 6.8), recombination is suppressed. This ensures that this large chromosomal region is inherited as a single locus and that the same sets of allelic mating type genes remain linked. The chromosome homologues with the alternate MAT alleles thus resemble primitive sex chromosomes (Lee et al. 1999). The 527-kb MAT-1 region of U. hordei has been fully sequenced. In addition to the typical mating type genes found at the outer ends of the locus, there are 44 additional genes located between the a and the b mating type genes, only one of which (STE11 encoding a putative serine/threonine kinase component of the pheromone-response signalling pathway) might also readily function in mating. Most of the genes found in MAT-1 of U. hordei have orthologues in U. maydis in the vicinity of either the a or the b mating type locus. The gene order over several regions is largely conserved between the species

but inversions are also apparent. Unlike U. maydis, which has very few repetitive elements within the mating type syntenic regions (and also overall in the genome) the MAT-1 locus of the barley smut contains a striking accumulation of repetitive elements [often apparently remnants of (retro-)transposons with long terminal repeats], making up more than 50% of the sequence of the whole locus. Such repetitive elements accumulated in between gene-rich stretches of synteny with U. maydis, bordering regions of inversions. It is suggested that the extremely high content of repetitive elements contributed during evolution to the chromosomal rearrangements of the small islands of gene content as found in U. hordei (Bakkeren et al. 2006).

B. Ustilaginomycotina Incertae Sedis, Malasseziales The genome of an apparently haploid strain is available for the dandruff-associated fungus M. globosa. Within the genome, a canonical divergently transcribed HD1 and HD2 gene pair was found, and the specific genes were named bE1 and bW1, following the established U. maydis nomenclature. Within a region of ~170 kb downstream of the bE1 gene lies a single pheromone receptor gene pra1 and a pheromone precursor gene mfa (Xu et al. 2007; Fig. 6.8). The pheromone receptor is related to the pheromone receptors from other species of the Ustilaginomycotina (Bakkeren et al. 2008; Fig. 6.5), and the pheromone precursor shares the typical structure of precursors of other farnesylated lipopeptide pheromones (Fig. 6.9). A phylogenetic analysis of all known pheromone precursors from the Ustilaginomycotina suggests that the M. globosa protein MgMfa1 has a common origin with UmMfa2 from U. maydis, UhMfa2 from U. hordei, and SrMfa2.1 und SrMfa3.1 from S. reilianum (Bakkeren et al. 2008). Although it is not known whether the fungus undergoes mating and whether it has different mating types (Xu et al. 2007; Hsueh and Heitman 2008), the existence of genes for typical mating type proteins within one defined chromosomal region has been taken as evidence that the fungus might have an extant bipolar sexual cycle, and that this might occur on human skin. The 173 kb region spanning the HD1 and HD2 gene pair to the pheromone receptor and pheromone precursor genes has been suggested to

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

encode the MAT locus of the fungus. This region contains in total 82 different genes but there are no orthologues of the U. hordei MAT-1 locus and only one gene (CID1 for a putative nucleotidyl transferase thought to play a role in the cell cycle) is shared with the C. neoformans MATa locus (see Section V.B). Moreover, unlike the situation of MAT-1 of U. hordei, the region in between the HD1 and HD2 genes and the pheromone precursor and pheromone receptor genes in M. globosa does not contain accumulations of repetitive elements (Xu et al. 2007). A partial genome sequence has also been established for another species from human skin associated with dandruff, Malassezia restricta. In the available sequences, fragments for a pra1, for bE1 and bW1 genes were found, indicating this species also harbors genes with presumptive mating type functions and that an extant sexual cycle remains to be discovered and which could occur on the surface of human skin (Xu et al. 2007). From other Exobasidiomycetes, such as Tilletia tritici and Tilletia controversa (Tilleticeae), biallelic and multi-allelic bipolar mating type systems are known (see Table 1).

C. Evolutionary Considerations Within the Ustilaginomycotina, bipolar and tetrapolar species have been described, some of which are closely related. Where known, bipolarity has evolved by fusion of the HD1 and HD2 gene locus with the pheromone precursor and pheromone receptor genes into one larger chromosomal region in which recombination is repressed (Fig. 6.8). As a consequence, this large chromosomal region is inherited as a single mating type locus (Bakkeren et al. 1993, 1996). Other features observed in U. hordei that are similar to the chromosomal situation in M. violaceum (Pucciniomycotina; see Section II.B), include the size differences of mating type alleles, the accumulation of repetitive elements and the overall bereft gene content of about one gene every 11 kb (Bakkeren et al. 2006; Hood 2002). These observations parallel the evolution of sex chromosomes as established in metazoans and plants, particularly in mammalian systems including the human Ychromosome (Charlesworth and Langley 1989; Lahn and Page 1999; Fraser and Heitman 2005; Bachtrog 2006, 2008; see Chapter 12 in this volume). However, the example of M. globosa with linked a and b mating type genes (and probably also S. salmonicolor, Pucciniomycotina; see Section III.A) contradicts at least some of the

121

predictions posited for sex chromosome evolution. Not only does the region with the mating type genes in M. globosa lack repetitive DNA elements, the region also has a comparatively high gene content – there is an average of 2.07 kb/gene – but the region harbours none of the genes required downstream to the mating type genes in the process of mating (hence the region lacks gene congruence; Xu et al. 2007). Because it is not known whether M. globosa is sexual, this region may serve other roles, or may be in the very early stages of evolution following a more recent fusion event (Fraser et al. 2004; Bakkeren et al. 2006). Different theories have been advanced to explain the widespread occurrence of sexual reproduction in nature. These include: generating genetic variability to adapt to heterogeneous environments, spreading of beneficial mutations, eliminating deleterious mutations (including those due to invading transposable elements) and the facilitation of DNA repair (Burt 2000; Butlin 2002). Sexual recombination contributes to species cohesion and generates more diverse genomes relative to species that lack recombination (Heng 2007). Asexual lineages, in contrast, are predicted to ultimately end in extinction, with the proliferation of deleterious transposable elements being one major driving factor (Judson and Normark 1996; Arkhipova and Meselson 2005). However, putative ancient asexual lineages are known throughout the eukaryotic kingdoms (Kirk et al. 2008; Neiman et al. 2009; Dunthorn and Katz 2010), confounding this prediction. Survival of ancient asexual rotifers of the class Bdelloidea has been attributed to the absence of deleterious retrotransposons (Arkipova and Meselson 2005). Asexuals might only survive in the long term after evolving mechanisms to prevent transposition and to completely eliminate the vertically transmitting transposable elements (Wright and Finnegan 2001; Dolgin and Charlesworth 2006). However, examples of sexually reproducing fungi largely devoid of transposable elements are U. maydis (Ka¨mper et al. 2006) and the filamentous ascomycete Neurospora crassa that developed a special inactivation mechanism [repeat induced point (RIP) mutation] for repetitive DNA active in the sexual stage (Galagan et al. 2003). Thus, an apparent lack of transposable elements is neither a unique hallmark of either sexual or asexual fungi.

Fusion of two mating type loci into one chromosome has apparently occurred more than once in the Ustilaginomycotina based on the lack of shared genes other than mating type genes within the enlarged mating type loci in U. hordei and in M. globosa (Bakkeren et al. 2006), and apparently at least once in the Pucciniomycotina (see Section III). In this scheme, the two-locus tetrapolar configuration is an ancestral character state and the bipolar one locus configuration a derived state

122

Ursula Ku¨es et al.

(see Section II.C). This interpretation is again supported by arguments such as the massive accumulation of repetitive elements and rearrangements with the added effect of increased recombinational suppression and thus lack of purifying recombination, which in turn further drives the accumulation of repetitive elements and rearrangements (Fraser et al. 2004; Bakkeren et al. 2006). Coelho et al. (2010), in contrast, argues from the apparently broad distribution of pseudo-bipolarity within the Sporidiobolales that the ancestral basidiomycete had possibly just one mating type region in which both groups of mating type genes were present. The Pucciniomycotina, as the earliest derived basidiomycete lineage (Liu et al. 2009), are proposed to have retained the mating system most similar to the ancestral basidiomycete system. Tetrapolarity, as it is known in the Ustilaginomycotina as well in the Agaricomycotina (see Section V), would then be the derived system with the few examples of bipolarity in these subphyla mediated by linkage of two groups of mating type genes being a recent derivation of the ancestral state. Confounding this assessment of the ancestral mating system is the observation that in the Pucciniomycotina there are evidently tetrapolar species, at least among the rusts whose mating type genes have yet to be analysed (see Section III.C). From a broader phylogenetic perspective, the fact that bipolarity is extant in the Ascomycota, Basidiomycota, and Zygomycota sensu lato argues that the ancestral fungal mating type configuration was bipolar. In this view, the emergence of tetrapolar sex determination is unique and restricted to the Basidiomycota and may have occurred only once, with secondary derivations of return to a bipolar state. In S. salmonicolor and in Sporobolomyces spec. strain IAM 13481 the genes STE20, KAP95, Lsm7, RibL18ae, RibL6, RNAPOL and GPD are closely linked to the genes of the pheromone system and gene AKOR to the HD1 and HD2 genes (Coehlo et al. 2010; Fig. 6.6), whereas in M. larici-populina and in P. graminis they are scattered over the whole genome with no linkage to any potential mating type gene with the possible exceptions of gene STE20 and of a copy of GPD that are linked at large distances to putative pheromone genes in M. larici-populina (UK, unpublished data). These facts contradict a structure of a larger chromosomal region with mating type genes shared between the Sporidiobolales and the Pucciniales.

Within the Dikarya, genes for the pheromonepheromone receptor communication system have mating type function only in the Basidiomycota, whereas the use of transcription factors as mating type functions is shared with the Ascomycota and other phyla of fungi (Dyer 2008; Casselton 2008; Lee et al. 2010b). This suggests that even in Basidiomycota, the ancestral species likely had homeodomain genes as a mating type locus, and that the pheromone receptors became mating type specific during the early evolution of the group (see Section II.C). One important evolutionary question that arises is when did the pheromone system become a mating type locus by itself? A crucial clue to this question of what evolved first – a system of two different loci linked tightly together on one chromosome or a system of two freely recombining loci – must consider also the evolution of multiple mating type alleles. At first glance, fusion of the two mating type loci into a single inheritable unit might have selective benefits: Fusing the two types of genetic loci together can favor inbreeding. With only two mating type alleles such as in the bipolar biallelic U. hordei, in a population at equilibrium there is a 50% chance of compatibility among random individuals. This corresponds to a 50% outbreeding potential (Figs. 6.11 and 6.12) and a 50% inbreeding avoidance among full siblings. A system with two types of genetic loci that are independently heritable traits each with two different alleles reduces the chance of both inbreeding and outbreeding to 25% since only individuals that differ at both mating types are inter-fertile. This scenario is not as favorable for mixing the population’s gene pool although inbreeding is reduced (Casselton and Ku¨es 1994; Hsueh and Heitman 2008). Such a system might be especially disfavored for species that are limited in the chance of meeting appropriate mating partners. Cells of different mating type of biotrophic pathogenic species such as U. hordei for example need to germinate together on a host’s leaf in order to infect the plant; a higher chance for inbreeding may favour successful infection (Bakkeren et al. 2006). Under such circumstances, there may have been no driving force to develop or maintain multiple mating type alleles in the bipolar U. hordei, in contrast to the closely related tetrapolar species U. maydis that also requires two compatible mating types to germinate on the same leaf (Banuett 1995;

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

123

A Cryptococcus neoformans var. neoformans MATα MFα1 MFα2 MFα3

MDAFTAIFTTFTSAATSSSEAPRNQEAHPGGMTLCVIA MDAFTAIFTTFTSAATSSSEAPRNQEAHPGGMTLCVIA MDAFTAIFATFTSAATSSSEAPRNQEAHPGGMTLCVIA

Cryptococcus neoformans var. neoformans MATa MFa1 MFa2 MFa3

MDAFTAIFSTLSSSVASSTTDAPRNEEAYGSGQGPTYSCVIA MDAFTAIFSTLSSSVASSTTDAPRNEEAYGSGQGPTYSCVIA MDAFTAIFSTLSSSVASSTTDAPRNEEAYGSGQGPTYSCVIA

Cryptococcus neoformans var. grubii MATα MFα1 MFα2 MFα3 MFα4

MDAFTAIFTTSISTVTSSSEVPRNQEAHPGGMTLCVIA MDAFTAIFTTSISTVTSSSEVPRNQEAHPGGMTLCVIA MDAFTAIFTTSISTVTSSSEVPRNQEAHPGGMTLCVIA MDAFTAIFTTSISTVTSSSEVPRNQEAHPGGMTLCVIA

Cryptococcus neoformans var. grubii MATa MFa1 MFa2 MFa3

MDAFTAIFSTLSSSVASKTDAPRNEEAYSSGNSPTYSCVIA MDAFTAIFSTLSSSVASKTDAPRNEEAYSSGNSPTYSCVIA MDAFTAIFSTLSSSVASKTDAPRNEEAYSSGNSPTYSCVIA

Cryptococcus gattii MATα MFα1 MFα2 MFα3 MFα4

MDAFTAIFTTFTSAAASSSEVPRNQEAHPGGMTLCVIA MDAFTAIFTTFTSAAASSSEVPRNQEAHPGGMTLCVIA MDAFTAIFTTFTSAAASSSEVPRNQEAHPGGMTLCVIA MDAFTAIFTTFTSAAASSSEVPRNQEAHPGGMTLCVIA

Cryptococcus gattii MATa MFa1 MFa2 MFa3

MDAFTAIFSTLSSSVASSTDAPRNEEAYGSGHGITYSCVIA MDAFTAIFSTLSSSVASSTDAPRNEEAYGSGHGITYSCVIA MDAFTAIFSTLSSSVASSTDAPRNEEAYGSGHGITYSCVIA

B Cryptococcus heveanensis MATα MFa1

MDAFTAIFTTFTSAAASSSEVPRNQEAHPGGMTLCVIA

Cryptococcus heveanensis MATa MFa2

MDAFTAIFTTLSSSASGNTESPRDQEYGSSGGGYSCIIA

Tremella mesenterica mating type a Tremerogen a-13

MDAFTISSPTPVNEETIIVFGPVNEEGGGNRGDPSGVCVIA

C Coprinopsis cinerea (tetrapolar) B43 mating type CcPhB1 Ccphb143 CoPhB3 Ccphb2.343 Ccphb2.243 Ccphb2.143 Ccphb3.143 Ccphb3.243 Ccphb3.343

MDAFLDLSEYIAQCDMAETVEVSENVPRNEDTRSRPSTICTIA MDDLTVILNLSELPPEFTAMMNSSPEFPSNTTDVPIDEDTPYRPVTACVIS MDSFTTLTAQSVFDELNLDVQDDVSHHFGFPSEEEAATEVPADQERPGDGRKAMGAFCVIV MDSFSSLDAFVVDEELLQLAIDIPQPIPGQDQQPPINEERPGAGTMGAFCIIN MASTTDTFTSLDAIRNDETLTLTLTPIPTNEPAAAPSTAAASSEVPRNFERPGSGKLSAFCIIA MDNFSTIDLAALCDELDFVQDVEALMASSTMGDAFVAGEDGPPANHERPGAGVNRAFCVIA MSDLFASLDLFLSSTEDNGCVCFDTNLSATTESQGCEILSKQASISTQELDGVLADFERRSGGGASWFCTIA MCDTFNTLTLNAYAEDPTPRLSNSTTNEITSTAPSDHTSLTLEELIDLPVDHERRTAGSSGQTWFCVIA MDSPLIPLDRVVLVLPDQMIDTDSSSFDECLLEFRLAVSSDERIAILDKQRLGIKAQAEMQAAGKNEERRTVNKPIKQSTLCPII

Coprinopsis cinerea non-mating type pheromone-like peptides Ccphl1 Ccphl2 Ccphl3 Ccphl4 Ccphl5 Ccphl6 ... Ccphl29 Ccphl30 Ccphl31 Ccphl32 Ccphl33

MDSFTTITSFTNKPSVEVPSDLETGGGSGNGAYCVIA MDSFATITSFTNKPSVEVPSDLETGGGSGNGAYCVIA MDSFTTIASFTNKPTVEVPSDLETGGGSGNGAYCVIA MDSFTTIASFTNQPSVEVPSDLETGGGSGNGAYCVIA MDSFTAITSFTNQPSVEVPSDLETGGGSGNGAYCVIA MDSFTTIASFTNQPSVEVPSDLETGGGSGNGAYCVIA ... MDASITSGNIPLDKETGGGAGNGAHCIIA MDSFTTVLSIFTTSQVEDVFVPVEEESGKKGGSGAYCTIA MDSFTIFSALFTSSSATSGAPATAEAVSTILQVPTDFETGGGSGNGAYCVIA MDSFTTINIPTNFETGGGSGNGAYCVIA MDSFQTICFSSEIAAAPNASANVDSSTTGAGVPVVAAAAPVIEIPVDEERNPWAGIWGGKMMCTIA

Fig. 6.11. Sequences for pheromone precursors as deduced from cloned genes of members of the Agaricomycotina. Pheromone precursors from Filobasidiales (A, Lengeler et al. 2002; Fraser et al. 2004), Tremellales (B, Metin et al. 2010), and tetrapolar (C, James et al. 2004b; Niculita-Hirzel et al. 2008; Ohm et al. 2010; Stajich et al. 2010; UK, unpblished observations), bipolar (D, James

et al. 2006, 2011; Martinez et al. 2009; UK, unpublished data), and homothallic species (E, Ku¨es and NavarroGonza´lez 2010) of the Agaricomycetes. The putative mature pheromones and pheromone-like peptides are shaded in grey and the CAAX-motifs for C-terminal processing (Caldwell et al. 1995) are underlined

Ursula Ku¨es et al.

124 Laccaria bicolor (tetrapolar) B mating type H82 Lbphb1 Lbphb2 Lbphb3

MDEFTTLFSDSLSEHACNGNDSIPSDIPLDEDSQYRPAIWCTIS MDSFTSLESLCESVDFITFQEDFSSSCSSHQPISSDLPIDQERYNAGNTGAFCVIS MDSFSTFDLSTFEPTSSSLPISSAHQTEDDDILRQLVDEDTKLGYAGYCVIA

Laccaria bicolor non-mating type pheromone-like peptides Lbphl1 Lbphl2 Lbphl3 Lbphl4 Lbphl5 Lbphl6 ... Lphhl39 Lbphl40 Lbphl41 Lbphl42 Lbphl44

MDAFTFYITSVDAIPIDEKTEVESLPTDEESGGSGNNAYCVIA MDAFTFYITSGVDAIPINEKTDIESAPTDEDNGGSGNNAYCVIA MDAFTFFVDIETTFTHHQTQGDSIPTNGESGGSGNNAYCVIA MDAFTLFFNVDTVFTNEQTQGDSIPTSEESGGSGNNAYCVIA MDAFAELVDYLSTPTVTTVGGDVASVPTNEEDRGTGNNAYCVVA MDAFTLFANIDTVFTNEQTQGDTIPTNEESGGSGNNAYCVIA ... MDSFTTISNTSNIPTNSDDGGSGNNAYCVVA MDSFSTISTSTDSIPTNYEGGGSGNNAYCVIA MDSFSTISTSTDSIPTNYEGGGSGNNAYCVIA MNSFSTISTSTGSIPTNYEGGGSGNNAYCVIA MFSLKFNHARAGFNRQNSAKFIQYSQHYLPPCTQPSLQVTRDHHLNRLIIHEPHETFLAEVGALCTIYEGGGSGNNAYCVIA

Schizophyllum commune (tetrapolar) B41 mating type Bap3-1 Bap3-2 Bap3-3 BBP2-1 BBP2-2 BBP2-3 BBP2-4 BBP2-5 BBP2-6 BBP2-7 BBP2-8 Bpl1 Bpl2 Bpl3 Bpl4 Bpl5

MDAFQSILDVLSAALDEPVDAPLTAVAQHPDADAVFDTPTDFERVGTGGTATAFCVVA MDVFAELFPTLVVDTLGAGDAPLGGPSPLAGPSSSGDSSFSILAGPAVYPSPVEGRHLHGPRWCVVA MDSFATLPALEDTLLQALLDACAVPEDDALDAMLSSSRPSSDAVVSDAERHGSGNMTYFCVVA MDSFDDLLDFLGAGTSRPAEASARNPLGSSSASSSSASLAASTSDLLSASPSSAPTSPDDVIMSILADAEHGYGGSNVHGWCVVA MDAQTTAPPTRPASPSPFAPSTAPTTVTPPASAAPPTNDTILALLANLEHEEDTDSNVHGWCIIA MDTFTYVDLAAVAAAAVADEVPRDFEDQITDYQSYCIIC MDDFITLDFLEDTTPVFDFAPPTPNELTPEGYDEFMRMVANSDSPDGYFGGYCVVA MAAFSARTSSPAPVASTIAFPSTRTKDTAPSTSGSSTPTRPASATPRAPDADLLRLLADARSAIARPPQDADEPDGYFAGYCVVM MSAAAPTTANQPKRNPAIQALVTSAPSDLSSSSTPSDLSSSSDAPQSPALTPRGGYYVSRTTGRAELKDAEREGDGNMTYFCEVM MSTGLSDGAGECIAYRTRPAQQTSPSCSSLSRRNHRARPRGRGFHRSRAYGPKDKLTNPLGSNDRPSTKPADADVRRALASGDKPCGYGGGYCVVG MDSFTTLSLLDGTMPTFEDDMPVSFLDAALSLSGDCFSSQSSASSSRSTPFASPSPSSPSPNLLSSAKAATDPHLVVNADSPCGFGGGYCVVA MDSFDTLLTAFVDGSLDGETAIEHLILAAASNSTISSHGGVSPTHGASSVTQHLSAHGSPNDGSPLPVNADTIEDYGSYCTIA MHPEHCAPPVGLQLRFICPCRLT MVILATARSTQVSNNFNTYIDECQLT MPSCFRYYYSSLCCGTIPRCAVVLFCVML MFPWSFPFGCFLWSFPFGVIVSYSRLRGLEFRQCISVTVCSMSFGMLDDHHCASR

Schizophyllum commune non-mating type pheromone-like peptides Phl251488 Phl111927 Phl253173 Phl249814 Phl269841 Phl270035 Phl270037

MDHFTILPTALATESHPIVIDEEPLAVFELDHPELGAGVPVNEEHHGSCHTQNQYVWCIVC MDHTTTPATTLATDSTNHPSDPLNEAPLATPEPTNDGVTNKTDDSADVPSNKEHHGTHHTQSAFAWCIVC MDFFREFSLSSVNDSSASLSSPAPSEQTSSSHNTSARHDAPLPVSDASLDDVENAPVDYERSTVAFYCIVA MDSFDLDLAALIASAPTVPTMAPTQTTNAEGMTYIDLSTFVEPGQLPVDEERSHGSFTGFYCIVS MDAFYTILSVLNLTGTSQEEDWDLEFPADYEGNSTTGNNGSSGTGMYCVVA MDAFSMAAIPDIEEIVNVPEDFESGGGTTGVYCVVA MDMFSPVVEIDVEDITTVDEEQQQQHNSSGFYCVVA

Pleurotus djamor (tetrapolar) B mating type RV95/957.30 PDphb1

MDAFFDFLPSISSSPSTTSAEELGSDIPLDEDTPWSPVSWCTIA

D Coprinellus disseminatus (bipolar) B-mating-type-like locus CdPHB1 CdPHB2.1 CdPHB2.2

MISTEESGLFLSQPRDSFSTLSLEEILTDGTFHEVFVAMAAEQPTEPDVLDPPRDQERDRALPGALCTIC MNTLSTITYTSDDFGIELDAATISTLLEGEMAQLLEASARGGDESEAIPYEMERNPNPSPIALCTLC MSEVQTEPGVRQVLADTSIYNSATKGLVDRSVGFPKHILPSPSSFIVSQSNEGDSIPTNEEKDRGADQSMKAFCVIA

Postia placenta (bipolar) B-mating-type-like locus Ppl135016 Ppl135014 Ppl135010 Ppl135011 Ppl135012 Ppl135013 Ppl135015 Ppl135043 Ppl135017 Ppl135031

MDAFTIAAPVPSEETEQIPADYDTGNGTTNYSCTIA MDAFFAIAAPVPSEETEQQIPADFDTGNGTTNYSCIIA MDAFFAIANPVPAEETEQVPVNHDTGGGSTSSTCTIA MDALITFDQIITSAQCAPDVDDYIACSIPVDEEHIGNTTSHSWCTIA MDDFASFEPSTAFIMEESIRSFDIPVDEDSCEPRPGLYCVVT MDEFSSESLQIDTNHITIIEGDRQFLVDAPPVDLDHWGSGNSSIFCTIV MDIFAEPFDSSLSTMTHHSSDLSQTLPNDTDIPLDFEHSGGGVIFYCVIS MDAFLTLAAPVPSDETDTSRVPSDYEWQTNTTNFSCIIA MDAFFVIANPVPAEETEQVPVNFDTGGGSTSSSCTIA MDALVDVFEPLSATPASAAERVNHPMPPQTDIPVDYEFQRDYGSTFQCCIIA

Phanerochaete chrysosporium (bipolar) RP78 non-mating type pheromone-like peptides Pcphl1 Pcphl2 Pcphl3 Pcphl4 Pcphl5

MRFTARFVGRYTMSDSGSGAVAARRRRSHRPSVLALEEGTPSRPAASSFRSLSSLGGSGAYCVIA MDAYTTIALAPAVPEEGTPVDFEGGGGPPTNPGGSCVIA MDAFAPLAEPFPADQVPRDAEGGSTSQNNGASCVIA MDTFVALADPIAGVEQVPRDAETGSSNNSHNCVVA MDAFFTLASPIATEQPEVPTDEETVSSNTNRASCVIA

E Moniliophthora perniciosa (homothallic) MVIRPFSRRWEMVSQPARRVMSYGQGKRVVTHRCRLNLHTTPCDYSRYEIFLRRSLCLIS MDSFENFDFLALASDESPIHNPFASFSTSLSEDASEVPSAALFNIESCRSRSFELQTDSQSLPTNHERAEAAAVHGGFCVIA MDSFTTITSSVEDITVSLPVPVDEELKGGGWAYCVIA ...VISLPVPVDEEQNGGGWAYCVIA ...IPINEEQQGGGWCYCVIA

Fig. 6.11. (continued)

Kahmann and Ka¨mper 2005; Brefort et al. 2009; Fig. 6.12). Increasing the number of alleles for one of two completely linked mating type loci as described for the pseudo-bipolar system found in the

Sporidiobolales would not change the potential for inbreeding. The set of genes with only two alleles will still determine the outbreeding frequency which will be 50% as is the case of typical bipolar species, e.g., U. hordei (Fig. 6.12). If

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

125

Fig. 6.12. Outbreeding potential as a function of number of mating type alleles. Bipolar biallelic species: Cryptococcus neoformans (Cn), Ustilago hordei (Uh); bipolar multiallelic species: Coprinellus disseminatus (Cd), Fomitopsis palustris (¼ Polyporus palustris; Fp); modified tetrapolar: Ustilago maydis (Um); tetrapolar species: Coprinopsis cinerea (Cc), Schizophyllum commune (Sc), Sporisorium

reilianum (Sr). The number of mating type alleles refers to the number of alleles at the single mating type locus or to the sum of alleles of two mating type loci if present in a species. For comparison, the outbreeding potential of a unipolar system is also indicated, such as occurring during same-sex mating in C. neoformans

recombination between the two loci is allowed, as is the case in S. salmonicolor, again the evolution of multiple mating type alleles at one locus does not facilitate an increase in outbreeding efficiency. A full release of linkage of the two mating type loci defines the condition known as modified tetrapolarity. This would result in the maximal realization of outbreeding potential but in the best case with a high number of mating type alleles at one locus this is still not higher than 50%. The outbreeding potential could increase above 50% only if the second locus would also develop multiple alleles (Fig. 6.12). Is the mating system of the pseudo-bipolar S. salmonicolor an intermediate state on the evolutionary path to tetrapolarity? Ongoing evolution of new HD1 and HD2 alleles in S. salmonicolor and in S. johnsonii after separation of the two species (Coelho et al. 2010) suggests this could be the case, rather than degeneration to a bipolar state. Or, as suggested by Coelho et al.

(2010), this species may combine the advantages of bipolarity and tetrapolarity. A growth impairment of mating type-recombined progeny (Coelho et al. 2010) however would favor the evolution of a reduced recombination (i.e., bipolarity), rather than increasing recombination. Progressive evolution of a bipolar mating system from a tetrapolar situation has been considered (Hsueh et al. 2008; see also Section V.A). As soon as linkage between the two loci was established in one genome, mating with another strain with compatible unlinked mating type genes might nevertheless have yielded recombinants among sexual progeny in which a new combination of mating type genes resulted in a second mating type specificity becoming linked together. With the achievement of two different specificities mediated by different alleles of linked mating type genes, this doubles the potential frequency of inbreeding and resets the allele number to two (Hsueh et al. 2008).

126

Ursula Ku¨es et al.

Developing multiple mating types in tetrapolar species is an expedient way out of the dilemma of low outbreeding efficiency (Fig. 6.12). By increasing the allele number for one of the loci as occurred in the tetrapolar U. maydis (modified tetrapolar), the overall probability of finding a mate increases proportionally with the number of different alleles being available. As calculated for U. maydis by using the formula (nA  nB – nA – nB + 1)/(nA  nB) given by Koltin et al. (1972), with 2 a mating type alleles and 25 b mating type alleles the outbreeding potential increases to 48% and the chance of finding an appropriate mate in the whole population is thus nearly as high as in the case of the bipolar biallelic species U. hordei (Fig. 6.12). Inbreeding restriction between siblings however remains high and only 25% of sibling pairs are compatible. Increasing the allele numbers at the second mating type locus further overcomes the problem of low outbreeding potential (Fig. 6.12) and still restricts inbreeding to 25%. With the three known a mating type alleles and five known b mating alleles of the tetrapolar S. reilianum, already an outbreeding potential of 53% is achieved (Fig. 6.12). The study by Schirawski et al. (2005) was likely not exhaustive in identifying b alleles in nature. Therefore, the actual outbreeding potential of the species might be much higher. Assuming ten different b alleles for S. reilianum, together with the three different alleles of the a mating type locus, the calculated outbreeding potential could be as high as 60%. A tetrapolar system with multiple alleles at each locus is thus superior to a tetrapolar system that is biallelic at one locus and multi-allelic at the second locus (modified tetrapolar) in terms of mediating a high outbreeding potential with minimal inbreeding restriction. However, only in the Agaricomycotina with dozens to hundreds of alleles at each of the mating loci does outbreeding efficiency reach values over 90% (Fig. 6.12). The outbreeding efficiency is achieved merely by having all mating type loci multi-allelic; for example, bipolar species such as Fomitopsis palustris (formerly Polyporus palustris) with an estimated 20 mating type alleles (Raper 1966) can also approach outbreeding potentials above 90% (Fig. 6.12). It is interesting to consider how the expansion from two to three a mating type alleles occurred in S. reilianum because the evolution of multiple alleles at a single pheromone receptor locus is

one of the evolutionary innovations only witnessed the basidiomycetes, and it appears to be an event that is independent of the evolution of multi-allelism in the Agaricomycotina. In the phylum Ustilaginomycotina, based on currently available data, there appear to be no recent duplications of pheromone receptor genes within a species (Devier et al. 2009; Fig. 6.5). Pra3 of S. reilianum clusters in phylogenetic analysis in a clade with Pra1 of the species and each one of the two alternate pheromone receptors of other Ustilaginales (Fig. 6.5) indicating a common origin for two of the three a mating type alleles in S. reilianum that might have been arisen by divergence from a common ancestral allele. The third pheromone precursor Mfa3 identified in S. reilianum (Figs. 6.8 and 6.9) is not as closely related to other known functional pheromone precursors in the Ustilaginales. An origin from another distantly related species has been considered (Bakkeren et al. 2008). However, translation of the U. maydis mfa pseudogene in all three reading frames indicates two blocks of high sequence similarity to Mfa3 of S. reilianum in two different reading frames (QAPVQEERPNQQF and SRPWTG*VIA; compare Fig. 6.9) suggesting that a three-allelic situation for the P/R locus was possibly found in the shared ancestor of U. maydis and S. reilianum. Generally in this context it is interesting to note that, DNA hybridization studies with a range of Ustilago and Sporisorium species revealed sequence conservation in a 1:1 manner to the a1 and a2 mating type genes of U. maydis, respectively (Bakkeren et al. 1992, Bakkeren and Kronstad 1996), suggesting low allele turnover in the younger evolution of the extant species and a broad stabilization of the biallelic stage within the Ustilagomycetes.

V. Mating Type Genes in Agaricomycotina A. Tremellomycetes, Filobasidiales Mating type control of sexual reproduction has been extensively characterized in the Cryptococcus species complex spanning the closely related human fungal pathogens C. neoformans var. neoformans (serotype D), C. neoformans var. grubii (serotype A) and C. gattii (serotypes B and C).

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

These species have a well established bipolar mating type system consisting of two opposite mating types, a and a. The species belong to the teleomorphic genus Filobasidiella and exist as a haploid yeast form and in a hyphal growth mode during sexual development (Kwon-Chung 1975, 1976a, b; Nielsen et al. 2003; Hull and Heitman 2002; Fraser et al. 2003; Lin and Heitman 2006; Lin 2009). The alternate alleles of the mating type locus of the species (MATa and MATa) have been found to be very distinct, spanning >100 kb and encompassing some 25 genes which in C. neoformans var. neoformans represents 6% of the total length of the chromosome on which the mating type locus resides (Karos et al. 2000; Lengeler et al. 2000, 2002; Fraser et al. 2004, 2005, 2007; Ren et al. 2005; Figs. 6.13 and 6.14). Mating type specific and unique genes are SXI1a encoding an HD1 homeodomain transcription factor unique to a strains (Fig. 6.1) or the SXI2a gene encoding an HD2 homedomain transcription factor unique to a strains (Fig. 6.1). Sxi1 and

127

Sxi2 together form a heterodimeric complex that orchestrates dikaryotic hyphal formation (Hull et al. 2002, 2005). Accordingly, the HD1 and HD2 gene pair has been reduced to the minimum needed for functionality with one type of gene in one locus and the complementary type in the alternate locus (compare Figs. 6.13 and 6.2). This is in contrast to other basidiomycetes in which a pair of divergently oriented homeodomain transcription factor genes are present in the MAT locus. SXI1a and SXI2a have as yet no known roles in haploid cells (including virulence) but function post-fusion. They are necessary and sufficient to undergo dikaryon formation and sexual reproduction following the fusion of a and a cells and thus establish the a/a cell identity (Hull et al. 2002, 2004, 2005). In accord, haploid transformed C. neoformans strains expressing both compatible homeodomain transcription factor genes are self-fertile and form monokaryotic hyphae with unfused clamp cells (Hull et al. 2005; Idnurm and Heitman 2005; Hsueh et al. 2008).

SX I1 CA P SP 1 O1 4 RP L2 2 BS P1 RU M 1 GE F1 CI D1

LP D1 BS P RP 2 O4 ST 1 E1 2 ST E3 ST E2 M 0 YO 2 ET F1 ST E1 1 M Fα 1 PR T1 ZN F1 RP L M 39 F M α3 Fα 2

Cryptococcus neoformans var. neoformans

MATa

b1 -2 b2 -2 d1 -1 bfg

18 32

m ip

a1 -2 a2 -2

RP L2 PR 2 T1 ZN F1 ST E1 M 1 Fa 2 ST E1 2 ST E3 M Fa M 3 Fa ST 1 E2 0 M YO 2 ET F1

SX I2 SP O RP 14 O4 BS 1 P2 RP L3 9 CA P1 LP D1 CI D1 GE F1 RU M 1 BS P2

Coprinopsis cinerea

A43 locus

Ab Q6

≈

Ab T6 Ab S6 Ab R6

21 41

.2 21 40

Ab V Ab 6 U6

ip m

Aα Z Aα 4 Y4 ge ne X

m sf1

Ph B1 ST E3 ph .1 b1 4

ST E3 ST -21 E3 51 -2 15 3

~28

3

Ph B ph 3 b ph 2.3 43 b2 ph .2 43 b2 .1 43 ST E3 ST .2a E3 ST .2b E3 ph .3 b ph 3.1 4 b3 3 ph .2 43 b3 .3 43

≈

~6

Schizophyllum commune

Fig. 6.13. Chromosomal organization of exemplified mating type loci of Agaricomycotina (after Fraser et al. 2004; Niculita-Hirzel et al. 2008; Stajich et al. 2010; Ohm et al. 2010). Mating type genes for HD1 and HD2 transcription factors are marked in blue and light blue, respectively (note that the relative order of HD1/HD2 gene pair positions in the species are not yet known) and mating type genes for pheromone precursors (small boxes) and pheromone receptors (large boxes) in red. Genes located within

B43 locus

A43 locus

≈

B41 locus l2

br

br

l3

~20

Lb Ph B Lb 4 Ph B6

Lb a1 Lb -1 a2 -1 bfg

ip m

ST E3 Lb .1 Ph B1 ST E3 LB .2 Ph B2 ST E3 Lb .3 Ph B3

Laccaria bicolor

6

ph p ba l5 p3 ba (3) p3 ba (2) p3 (1 ) ba r3

l1 br

bb pe (2 ) bb r2 bb p2 bp (1) l2 bp l bb 1 p bb 2(4 p2 ) ( bp 7) l3 bb p2 bb (3) p bb 2(5 p ) bb 2(8) p2 bp (6) l4

≈

~550 ~25

MATα

A1 locus B1 locus

mating type loci that encode components of the pheromone response pathway (downstream of the pheromonepheromone receptor system) are shown in green, and pheromone receptor genes located outside of defined mating type loci are illustrated in lilac. Other genes within the mating type loci are shown in light grey. Genes within the black border define the mating type loci. Arrows indicate direction of transcription, numbers beneath a map indicate distances between genes in kilobases

128

Ursula Ku¨es et al.

Fig. 6.14. Model for the evolution of the MAT locus of the pathogenic bipolar, biallelic Cryptococcus species from the Filobasidiales, starting from an ancestral tetrapolar system with a locus containing a complete HD1 and HD2 gene pair and an unlinked locus containing a pheromone receptor gene and pheromone precursor genes (after Hsueh et al.

2011). HD1 and HD2 mating type genes are indicated by blue letters, and pheromone precursor and pheromone receptor genes by red letters. Arrows indicate direction of transcription of genes. The resulting extant MATa and MATa loci shown in the figure represent Cryptococcus neoformans var. grubii

Two types of a/a cells can be distinguished: dikaryotic filaments (a þ a) with fused clamp cells at the hyphal septa or monokaryotic diploid (a/a) filaments with unfused clamp cells at the hyphal septa. Formation of these two types of filaments is dependent on temperature. At 25  C, upon mating of a and a cells, cell–cell fusion produces a zygote from which dikaryotic hyphae are produced. Diploid strains are thermally dimorphic and at 37  C produce yeast cells, whereas at 25  C differentiation

occurs to produce monokaryotic hyphae with unfused clamp connections and basidia decorated with four chains of basidiospores (Hull and Heitman 2002).

In contrast, recent studies reveal that pheromones and pheromone receptors determine haploid a and a cell identity (Stanton et al. 2010). Interestingly, haploid a and a cells differ in behaviour.

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

Nitrogen limitation causes MFa pheromone secretion from a cells and a cells respond to MFa pheromone by producing conjugation tubes. The responses of a cells to mating pheromone can involve either conjugation tube formation (polarized cell growth) or the formation of enlarged cells (isotropic cell growth) depending on the variety or species. Following cell–cell, conjugation tube–cell, or conjugation tube–conjugation tube fusion, the nucleus migrates unidirectionally from the a cell towards the a cell nucleus (Chang et al. 2000; Wang et al. 2000; McClelland et al. 2002, 2004), suggesting a kind of primitive dioecism for this group of basidiomycetes (Lee et al. 2005). In turn, nitrogen limitation in the presence of a cells induces MFa pheromone production in a cells and the presence of MFa pheromone initiates production of enlarged, round, refractile a cells (Moore and Edman 1993; Davidson et al. 2000; Shen et al. 2002). Exchanging the pheromone receptor and the pheromone precursor genes in a cells with those from the opposite mating type changes the identity of the resulting MATaa cells, which now respond to and mate with a cells. These cell identity altered cells mate but do not produce a conjugation tube, suggesting the MATa pheromone receptor Ste3a is responsible for the different morphological behaviour of the cells (Stanton et al. 2010). In addition to paracrine signalling during mating reactions, MFa pheromone may also stimulate same-sex mating of a cells via autocrine signalling (Shen et al. 2002; see below). MFa pheromone production is tightly controlled by the DEAD box protein Vad1 that ensures low steady transcript levels of MFa pheromone precursor transcripts by RNA degradation, unless environmental conditions favour mating and increase pheromone expression (Nielsen et al. 2007; Park et al. 2010). Inactivation of the gene for the global repressor Tup1 derepresses MFa and MFa production and impairs mating efficiency (Lee et al. 2005, 2009), indicating a complex regulatory network governs pheromone production as a key early step in mating. In addition to the HD1 and HD2 SXI1/SXI2 genes in the alternate mating type locus alleles, there are allelic genes for mating type specific Ste3-like pheromone receptors of divergent sequence (STE3a and STE3a, respectively; Chung et al. 2002; Chang et al. 2003; Stanton et al. 2010; for an analysis of their products, see Fig. 6.5), and three (or four) individual pheromone precursor genes

129

for mating type specific lipopeptide pheromones (MFa1, MFa2, MFa3 and MFa4 and MFa1, MFa2, and MFa3, respectively; Davidson et al. 2000; Shen et al. 2002; Stanton et al. 2010; for their products, see Fig. 6.11A) as well as alleles of other genes that are predominantly mating type specific (Figs. 6.13 and 6.14). This includes the STE12a and STE12a genes encoding Ste12-like transcription factors that appear not to function downstream of the pheromone-sensing MAP kinase cascade based on epistasis analysis (Davidson et al. 2000; for a review of this signal transduction cascade, see Lengeler et al. 2000a; Zhao et al. 2007) but do function in regulating aspects of mating (Wickes et al. 1997; Alspaugh et al. 1998; Yue et al. 1999; Chang et al. 2000; 2001, 2004; Ren et al. 2006). Other mating type specific genes include STE20a and STE20a for p21-activated protein kinases (Karos et al. 2000; Wang et al. 2002; Nichols et al. 2004) and STE11a and STE11a for kinases operating in the MAP (mitogen-activated protein) kinase pathway (Karos et al. 2000; Clarke et al. 2001; Davidson et al. 2003). Furthermore, the alleles of SPO14, RUM1, and BSP1 within the MAT loci may function in meiosis and sporulation (Lengeler et al. 2002; Fraser et al. 2004). A summary of potential gene functions of other genes is given by Hsueh et al. (2011). Other than the allelic STE3 genes for pheromone receptors and the allelic MF genes for pheromones, 15 of the genes within the biallelic MAT loci of the three Cryptococcus species (amongst STE12, STE11, STE20) display a mating type specific phylogeny, indicating an ancestral association of the genes with the MAT loci. Five other genes exhibit species-specific rather than mating type specific phylogenies (LPD1, CID1, RPO41, BSP2, GEF1), indicating either a recent recruitment to the MAT locus or ongoing gene conversion that has reset the evolutionary clock for one or more of these highly conserved genes. These five genes are largely syntenic in the three species in contrast to the genes of ancestral MAT association that have undergone considerable rearrangment in their gene order (Fraser et al. 2004, 2007; Metin et al. 2010; Rodriguez-Carres et al. 2010; Figs. 6.13 and 6.14). The large biallelic mating type locus in the Cryptococcus species is reminiscent of sex chromosomes of multi-cellular eukaryotes: the locus contains genes of sex-related functions, recombination in the locus is suppressed during meiosis,

130

Ursula Ku¨es et al.

and the locus is rich in transposable elements and remnants (in C. neoformans var. neoformans: 17.3% of total length in MATa and 13.2% of total length in MATa; Lengeler et al. 2002; Fraser et al. 2004). Between the different species, transposon remnants and repetitive sequences are highly variable in position suggesting that they were independently acquired during different time points in evolution (Fraser et al. 2005). The MAT loci of the three pathogenic Cryptococcus species consist of four gene strata of different ages (Fig. 6.14). The first strata represents an ancient group of genes that share little allelic DNA sequence, show strong clustering into MATa and MATa genes and include the genes involved in the MAP kinase signalling pathway. The second and the third strata (intermediate I and intermediate II) represent genes whose alleles have progressively higher nucleotide identity and less discrete but still MATspecific phylogenetic patterns, indicating that they were more recently incorporated into the mating type locus. The fourth gene strata, possibly the youngest genes within the MAT locus, are those that show a species-specific phylogenetic pattern and allele identity
90% (Fraser et al. 2004, 2005). From this line of investigation, Fraser, Heitman and colleagues proposed models to explain the evolution of the large mating type regions in the genome of bipolar biallelic Cryptococcus species (Fraser et al. 2004, 2006, 2007; Hsueh and Heitman 2008; Metin et al. 2010; Hsueh et al. 2011; Fig. 6.14). Assuming the tetrapolar situation to be the ancestral state in the Filobasidiales, the authors postulated: In the first evolutionary step, gene cluster formation occurred in parallel into the two independent mating type loci, one of which contained a pair of divergently transcribed HD1 and HD2 genes (HD locus) and the other of which contained the pheromone receptor and pheromone precursor genes (P/R locus). Among the genes acquired to the MAT loci were sexrelated genes such as STE12, STE11, and STE20 for the pheromone response MAP kinase pathway (ancient class of genes). In the second evolutionary step, the MAT loci serially expanded and underwent diversification of allele sequences of genes captured into the MAT clusters (intermediate I and intermediate II). Subsequently in a third step, the unlinked MAT loci then fused to a single larger mating type-determining locus. This step happened in one of the mating types and gave rise to an unstable transient tripolar system.

In this step, flanking genes may have been incorporated into the fused locus, as the region between the two ancestral HD and P/R regions that now lie adjacent (recent genes). Based on the analysis of species aligned with the pathogenic Cryptococcus species complex (including Cryptococcus heveanensis, Cryptococcus amylolentus, and Tsuchiyaea wingfieldii; see below and Section V.B.1) the ancestral state can be inferred to have had paired divergently oriented HD genes, and thus one or the other was lost in the evolutionary transitions to the extant bipolar state in C. neoformans and C. gattii. This may have occured before, after, or concomitant with the step during which the ancestral unlinked loci became fused. Stabilization was achieved in a fourth stage via gene conversion or recombination and collapse of the tripolar intermediate to the bipolar state. A series of more recent gene conversions and inversions gave rise to the extant bipolar mating type system with MAT alleles with genes of divergent allellic sequences. Remnants of transposons might have facilitated the observed inversion events by providing homologous sequences as templates for recombination. As a further consequence, the rearrangements together with sequence diversification provide the mechanistic basis for suppression of recombination between the MAT locus alleles, in particular repressing recombination in the region in between the unique genes for homeodomain transcription factors and the group of genes for the pheromone-pheromone receptor system. Experimental evidence has been presented to support the proposed scenario of evolution. First, strikingly within the MAT loci the genes for the homeodomain transcription factors localize to the left flank and the genes for the pheromone receptors and for the pheromone precursors and the genes for the pheromone response pathway localize to the right border (Fraser et al. 2004; Metin at al. 2010). Second, the MAT locus is flanked by recombinational activators that may have driven the expansion and fusion of the loci and the multiple rearrangements within the fused locus. Recombination within the MAT locus in meiosis is largely suppressed. In the MAT flanking regions, recombination occurs 10- to 50-fold more frequently than the genome-wide average. Furthermore, due to negative interference double-strand crossovers at both sides of the MAT locus are about four times more often than expected and hotspots at both

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

sides of MAT are present within distinct GC-rich regions (Hsueh et al. 2006). Third, fertile Cryptococcus strains were constructed in which the HD1 and the HD2 genes were relocated from the MAT locus into another unlinked genomic location (URA5). Upon crossing, the modified strains of opposite mating types now with two mating type loci completed a tetrapolar sexual cycle. In tripolar crosses between a normal MAT strain and a constructed tetrapolar strain of opposite mating type, the resulting progenies were of decreased fertility – 50% of the progeny were sterile and among progeny the chance of inbreeding is only 12.5%. Transitions between outcrossing and inbreeding preference might in such a situation select for stabilization of a bipolar system with finally only two mating types (Hsueh et al. 2008). Fourth, as a further important line of evidence for the proposed MAT evolutionary model, the MAT loci of a non-pathogenic tetrapolar Cryptococcus species (C. heveanensis) from a sister clade (sensu lato Kwoniella clade) have recently been described, indicating that the (multi-allelic) tetrapolar situation with a divergently transcribed HD1 and HD2 gene locus and an unlinked locus with pheromone receptor and pheromone precursor genes exists also in the Tremellomycetes (Metin et al. 2010; see Section V.B.1). Furthermore, a complete HD1 and HD2 SXI1/SXI2 gene pair has been found in T. wingfieldii (Hsueh et al. 2011), another species from the Filobasidiales clade (Findley et al. 2009) for which only a single isolate is known and which thus remains currently uncharacterized in terms of sexual reproduction. SXI1 and SXI2 reside in a subcloned DNA region of 56 kb (HD locus), and a second subcloned region of 70 kb (P/R locus) contains the pheromone receptor gene STE3 and three pheromone precursor genes MFa1, MFa2, and MFa3. Importantly, it was shown that the two subcloned regions localize to different chromosomes. In both subcloned DNA regions, more genes that are known from the MAT loci of the three Cryptococcus species were found to be present, providing an insight into the locations of genes in the two ancestral MAT gene clusters. The overall gene content of the two subcloned regions is quite similar to that of the Cryptococcus species, with the exception that the STE11 gene is missing and apparently lies elsewhere in the genome. The five most recently acquired genes known from the MAT loci of the three pathogenic Cryptococcus varieties/species (LPD1, RPO41, BSP2,

131

CID1, and GEF1) are already present in the vicinity of the genes for the pheromone signalling system providing evidence that these were likely acquired into the fused MAT locus via an earlier acqusition into the ancestral P/R loci rather than captured and entrapped during the translocation event that fused the HD and P/R loci in the tripolar intermediate (Hsueh et al. 2011). Overall from the available data, species comparisons imply that the first steps in the model of the genomic events leading from a tetrapolar to a bipolar mating type system occurred prior to the split of the Filobasidiales, and the sensu lato Kwoniella clade and other steps occurred after this split but prior to speciation within the Filobasidiales (Fraser et al. 2004; Metin et al. 2010; Hsueh et al. 2011; see Section V.B.1). Data for mating type loci from further species of the Filobasidiales are accumulating: C. amylolentus, originally described to be asexual, has now been found to have an extant sexual cycle (teleomorphic name Filobasidiella amylolenta). Two DNA regions for the HD and P/R loci have been subcloned, and the initial analysis suggests these appear to be similar to those of T. wingfieldii, also with a complete HD1 and HD2 gene pair in the HD locus and with the STE11 gene missing from the P/R locus (Hsueh et al. 2011). Filobasidiella depauperata is a non-pathogenic, homothallic, exclusively filamentous species of the Filobasidiales clade (Findley et al. 2009; Rodriguez-Carres et al. 2010). Of the genes found within the MAT loci of the heterothallic Cryptococcus species, three of the species-specific genes (LPD1, CID1, and RPO41) and five of the ancestral MAT-associated genes (STE20, MYO2, PRT1, ZNF1, and RPL39) are in F. depauperata within a contiguous cluster cloned from the species. Moreover, STE11 is found translocated to another cloned gene cluster of F. depauperata that otherwise shares no genes with the MAT locus of the three fertile Cryptococcus species (Rodriguez-Carres et al. 2010). Of the cloned genes, STE20 and MYO2 show an evolutionary origin common to the MATa mating type of the Cryptococcus species of the clade of Filobasidiales and for STE11 an origin common to the MATa mating type whereas all other genes exhibit a species-specific phylogeny (RodriguezCarres et al. 2010; Hsueh et al. 2011). Neither the pheromone receptor and the pheromone precursor genes, nor the HD1/HD2 genes were, however,

132

Ursula Ku¨es et al.

found thus far in F. depauperata. The mechanism of homothallism in F. depauperata is unknown and awaits a full genome sequence to clarify the status of the typical basidiomycetous mating type genes in this species (Rodriguez-Carres et al. 2010), i.e. also whether there are possibly no such genes. Alternatively, conversion of mating type genes into constitutively active gene versions, formation of self-compatible loci due to incorporation of compatible mating type genes, or mutations in genes downstream of the mating type loci may have bypassed the need for mating type genes. All of these types of mutations, with the exception of the loss of mating type genes, have been found to mediate self-fertility in other Agaricomycotina (see Section V.F). Based on the recent discovery of unisexual mating of Cryptococcus species, which occurs without a requirement for the HD1/HD2 genes, it is also possible that the extant homothallic sexual cycle for F. depauperata is a form of same-sex mating (Wickes et al. 1996; Fraser et al. 2005; Lin et al. 2005, 2007). Heterothallic mating of haploid yeast cells (opposite-sex mating) of the three fertile Cryptococcus varieties gives rise to dikaryotic filaments with fused clamp connections. On the dikaryotic filaments, ultimately basidia with sexual basidiospores are formed (Kwon-Chung 1975, 1976a, b; Hull and Heitman 2002; Nielsen et al. 2003; Fraser et al. 2003; Lin and Heitman 2006; Lin 2009). In natural populations of C. neoformans var. grubii and of C. gattii, there is a strong predominance of a strains, and in many populations a strains are rare or do not appear to be present (Kwon-Chung and Bennett 1978; Franzot et al. 1997; Lengeler et al. 2000b; Viviani et al. 2001, 2006; Yan et al. 2002; Litvintseva et al. 2003, 2006; Kidd et al. 2005; Okabayashi et al. 2005; Saracli et al. 2006; Randhawa et al. 2006; Hiremath et al. 2008). As another specific feature, the three Cryptococcus species are also able to undergo unisexual reproduction (in the literature referred to as same-sex mating, also haploid fruiting, or monokaryotic fruiting) and this is likely related to the bias of a strains in nature (Hull and Heitman 2002; Lin et al. 2007). Unisexual reproduction of C. neoformans is a novel form of fungal homothallism and occurs either with an individual obtained from a single spore undergoing sex in solo culture or can involve same-sex mating between two genetically

distinct individuals of the same mating type (Fraser et al. 2005; Lin et al. 2005). Same-sex mating in Cryptoccocus involves fusion of cells of the same mating type, diploidization, and meiosis. Same-sex mating results in septated diploid monokaryotic hyphal fragments (monokaryotic filaments) but with unfused clamp cells and culminates in basidia and basidiospore development (Hull and Heitman 2002; Idnurm et al. 2005; Lin et al. 2005, 2007, 2009). Evidence for unisexual reproduction in nature along with genetic recombination between nuclei of non-isogenic strains is available for C. neoformans var. neoformans, C. neoformans var. grubii, and C. gattii (Fraser et al. 2005; Hiremath et al. 2005; Lin et al. 2007, 2009; Bui et al. 2008; Saul et al. 2008). Same-sex mating is usually readily observed with a strains (Wickes et al. 1996; Lin et al. 2005, 2006) although some a strains are able to undergo same-sex mating (Hull and Heitman 2002; Tscharke et al. 2003; Lin et al. 2006). The difference in degree of same-sex mating resides in the alleles of the MAT locus (Lin et al. 2006) which may be linked to the pheromone response pathway: For unisexual reproduction the pheromone response pathway contributes and also the meiotic regulators Dmc1 and Spo11 (Yue et al. 1999; Davidson et al. 2003; Hsueh and Shen 2005; Lin et al. 2005) as well as the specific transcription factors Mat2 and Znf2 that function downstream of the pheromone response pathway (Lin et al. 2010). Same-sex mating is positively influenced by pheromones – reduction of hyphal differentiation in a cells of C. neoformans var. neoformans deleted for the MFa1, MFa2 and MFa3 genes provides evidence for autocrine pheromone signalling. Enhanced same-sex mating of MATa cells upon transformation with a MFa gene and stimulation by the presence of MATa cells at a distance implicates paracrine pheromone signalling in the process (Wang et al. 2000; Shen et al. 2002). Samesex mating in response to nitrogen starvation is defective in a cells when the GPB1 gene for a G protein b-subunit essential for opposite-sex mating is deleted (Wang et al. 2000). In a cells, overexpression of the Ste12a transcription factor enhances same-sex mating (Tscharke et al. 2003). Deletion of STE12a from the MATa locus reduces but does not abolish same-sex mating (Yue et al. 1999; Ren et al. 2006, Lin et al. 2010). Deletion of STE11a results in filamentation defects during

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

133

same-sex mating (Chung et al. 2002; Davidson et al. 2003; McClelland et al. 2004). In contrast to opposite-sex mating, same-sex mating is independent of the homeodomain transcription factors Sxi1a and Sxi2a (Hull et al. 2002, 2005; Lin et al. 2005; Stanton et al. 2010). The pheromone-transporter Ste6 that functions during opposite-sex mating appears to be dispensable for same-sex mating (Hsueh and Shen 2005). Furthermore, in the genome of C. neoformans var. neoformans, there is a non-mating type gene for a novel pheromone-receptor-like protein Cpr2 (CnnCpr2). This protein is a constitutively active G proteincoupled receptor with ligand-independent activity that functions in both opposite and same-sex mating to activate the pheromone response pathway. Cpr2 (CnnCpr2) is sufficient but not necessary for same-sex mating (Hsueh et al. 2009). The available data define complex regulatory networks that exist in Cryptococcus species to control the pathways of opposite- and of same-sex mating, and these pathways overlap but are not fully identical (Idnurm et al. 2005).

late re-examination of the mating type configuration of other closely aligned species, which may therefore be similarly tetrapolar, or have undergone a recent transition to bipolarity from a tetrapolar ancestral state. The C. heveanensis A mating type locus has two known alleles with one pheromone receptor gene STE3 (for an analysis of the pheromone receptor, see Fig. 6.5), one pheromone gene, MFA1 or MFA2 (for the product, see Fig. 6.11B), a copy of STE12 and a new gene CNG04540. In the same chromosomal region of ~180 kb, the STE11 and STE20 genes and several other genes known from the MAT loci of the two C. neoformans varieties and C. gattii reside, adjacent to novel genes not present in the other species within or in the vicinity of the MAT loci, some of which have no apparent homologues in these other species. The high level of sequence identity of STE11 and other genes (99–100%) in different C. heveanensis isolates suggests that these are flanking the MAT-specific P/R locus region but are not part of the MAT locus (Metin et al. 2010; Hsueh et al. 2011).

B. Tremellomycetes, Tremellales

This finding provides robust support for the second hypothesized step in the proposed evolutionary model for the origin of a fused MAT locus in the three bipolar pathogenic Cryptococcus species from the Filobasidiales involving capture and acquisition of neighbouring genes into an expanding ancestral tetrapolar MAT locus (Hsueh et al. 2011; Fig. 6.14).

1. Mitosporic Tremellales/Sensu Lato Kwoniella Clade The mitosporic Tremellales are part of a poorly resolved clade (Statzell-Tallman et al. 2008), many of which have been given the genus names Cryptococcus. This clade also includes the anamorph genus Bullera and the newly described teleomorph genus Kwoniella. The clade is clearly distinguished from the Filobasidiales to which the pathogenic C. neoformans var. neoformans, C. neoformans var. grubii and C. gattii belong, and also from the sexually proficient Tremellaceae. This group of fungi has been assigned to the sensu lato Kwoniella clade (Findley et al. 2009). The first sexual species in this clade, Kwoniella mangroviensis, was recently reported to be bipolar with two mating types, A and a (Statzell-Tallman et al. 2008). The non-pathogenic C. heveanensis has now been shown to have a sexual state, leading to a teleomorphic assignment as Kwoniella heveanensis. Sexual reproduction is apparently controlled by two unlinked mating type loci in a classic tetrapolar mating system (Metin et al. 2010). This discovery will stimu-

The unlinked B mating type locus lies in an ~80 kb chromosomal region containing a typical HD1 and HD2 gene pair, and the mating type specific region is restricted to a region of only ~2 kb involving the HD1/HD2 genes. This is a key finding that within the Tremellomycetes the HD1 and HD2 divergent gene pair was present prior to the divergence of the Filobasidiales and the Tremellales and represents the most parsimonious ancestral state. Moreover, in the tetrapolar species C. heveanensis a total of six different alleles of the HD1 and HD2 gene pair have been described, giving rise to multiple mating types. As in U. maydis, within the divergently described gene pair the 5’ region for the putative N-terminal dimerization domains of the proteins is of low sequence conservation, in contrast to the more conserved regions encoding the homeodomains and the C-termini of the proteins (Metin et al. 2010).

134

Ursula Ku¨es et al.

2. Tremellacecae Mating type lipophilic pheromones (Tremerogens) with an S-polyisoprenyl cysteine residue at the carboxy-terminus of the peptide have been isolated and sequenced from the yeast-like jelly fungi Tremella mesenterica (Tremerogen a-13: EGGGNRGDPSGVC from the a mating type and Tremerogen A-10: EHDPSAPGNGYC from the A mating type) and Tremella brasiliensis (Tremerogen A-I: DSGSSRDPGASSGGC and Tremerogen A-9291-I: DSGGSRDPGASSGGC occuring in relative proportions of 1 : 1 in the A mating type; Sakagami et al. 1979, 1981b; Ishibashi et al. 1984). The release of the genome of T. mesenterica by the JGI (http://genome.jgi-psf.org/Treme1/Treme1. home.html) allowed identification of one gene for a small pheromone precursor of a single mature pheromone unit (Tremerogen a-13; see Fig. 6.11B) present in the genome (on scaffold 7) close to the TmSTE12, TmSTE3 and TmSTE20 genes. The genome also contains a divergently transcribed HD1 and HD2 gene pair (on scaffold 13). Although there are also a considerable number of genes unique to the species, many other genes found in the C. neoformans varieties, C. gattii, C. amylolentus, and C. heveanensis within or linked to the mating type loci are also present in T. mesenterica in the vicinity of the mating type genes, indicating a common evolutionary origin (Metin et al. 2010). Notably on scaffold 7 at 60 kb distance to TmSTE12, TmSTE3, and TmSTE20, there is also in T. mesenterica a copy of the gene TmSTE11 (http://genome.jgi-psf.org/Treme1/Treme1.home. html; UK, unpublished data). Most genes are shared with C. heveanensis and phylogenetic analysis of eight different genes indicates that, of the Cryptococcus species, it is the closest relative to T. mesenterica. However, various rearrangements in gene order and gene directions and speciesspecific gene insertions or losses are apparent for example between T. mesenterica and C. heveanensis (Metin et al. 2010). Furthermore, also a gene for a putative pheromone receptor highly similar (77% amino acid similarity) to the nonmating type pheromone receptor CnnCpr2 of C. neoformans var. neoformans (see Section IV.A) is found in T. mesenterica (on scaffold 4, http://genome.jgi-psf.org/ Treme1/Treme1.home.html; UK, unpublished data).

The localization of the HD1 and HD2 gene pair and the pheromone precursor and pheromone

receptor genes to different scaffolds is consistent with T. mesenterica being tetrapolar with multiple mating types. Two A mating type alleles (A and a or A1 and A2) and 68 different B mating type alleles (B1, B2, B3, etc.) were identified from isolates from the wild (Bandoni 1963; Wong and Wells 1985). The A mating type locus defines courtship and conjugation tube formation prior to mating whilst the B mating type locus is required for formation of dikaryotic hyphae after fusion (Bandoni 1963) which suggests that the locus with the HD1 and HD2 genes is the B locus with the multiple alleles for defining mating types. The situation resembles thus the modified tetrapolar (Wong and Wells 1985) cases of C. heveanensis and U. maydis with a biallelic locus encoding the pheromone precursors and the pheromone receptors and a multi-allelic locus encoding the two types of homeodomain transcription factors. The implications are that the evolution of a multi-allelic P/R locus was associated with the origin of the Agaricomycetes (where it is the predominant pattern; see Section V.E) and not Agaricomycotina (that includes Tremellales), and thus the biallelic P/R locus is more common among dimorphic fungi that have a haploid yeast phase. Multi-allelism of the P/R locus might have evolved more than once, because a triallelic P/R locus in S. reilianum is existent within the Ustilagomycetes (see Section IV.A). Exhaustive analyses suggested that the B mating type locus of T. mesenterica is not undergoing recombination between different alleles although dual mater strains might be formed in crosses (Wong and Wells 1985). Lack of recombination to produce new mating type alleles is easily explained by the structure of the locus with a single pair of divergently transcribed genes of low DNA homology. As deduced by Wong and Wells (1985) from further crosses of dual maters to their parental strains, their production is best explained by an abberant disomic condition for the B locus which can easily be resolved in meiosis back to the original alternative single copy situations.

C. Agaricomycetes, Auriculariales The Auriculariales are another ancient lineage in the Agaricomycetes (Hibbett 2006). First molecular data are available from this fungal order by partial gene sequences for STE3-like pheromone receptors of the species Auricularia polytricha (GenBank AY226009) and Auricularia auriculajudae (GenBank DQ303127; FJ756941; FJ756942).

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

Two of the sequences available from different mating types of A. auricula-judae differ significantly from each other (65% sequence identity, 80% sequence similarity), supporting that they may be part of a mating type system that involves pheromone signalling. Reports on matings of A. polytricha from different parts of the world described the species to be tetrapolar with the typical reactions of the Agaricomycetidae A and B mating type loci in dikaryon formation with the A mating type locus controlling clamp cell formation and the B mating type locus controlling clamp cell fusion (Wong and Wells 1987). However, a report from China and a report from Malaysia deduced A. polytricha to be bipolar (Luo and Cheng 1989; Daud and Napiah 1991) and A. auricula-judae was also reported to be bipolar (Whitehouse 1949; Luo and Cheng 1989; Wu et al. 2004). Overall, the data are currently too little and contrasting to get a clear view on the situation.

D. Agaricomycetes, Cantharellales Within the ancient lineage Cantharellales within the Agaricomycetes (Hibbett 2006), species are reported such as Sistotrema brinkmannii in which homothallic strains are found in the same population as bipolar heterothallic individuals. It is expected that there should be mating type genes in all of these strains since mating between homothallic and heterothallic individuals is possible (Lemke 1969; Ullrich and Raper 1975). While inbreeding should be the rule in the homothallic strains, this species may possess a mixed mating system allowing both selfing and outcrossing. The co-existence of two contemporary mating type systems may be analogous to the recent appreciation that C. neoformans retains extant heterothallic and homothallic unipolar modes of sexual reproduction (see Section V.A). (Partial) sequences for pheromone receptor genes from Thanetophorus cucumeris (Rhizoctonia solani) are available (GenBank ABL67527; AAP57503). Despite the phylogenetic position of the organism, a pheromone receptor of R. solani clusters with pheromone receptors of the Pucciniomycotina, pheromone receptors of the Ustilaginomycotina and pheromone receptors of the Tremellomycetes (Fig. 6.5). T. cucumeris is an incompletely resolved species complex of at least twelve different anastomosis groups (vegetative incompatibility groups) with a intertwined homothallic and heterothallic population structure. At least some anastomosis groups of this species

135

complex appear to be bipolar but they tend to also contain homothallic individuals (Cubeta and Vilgalys 1997; Julian et al. 1997; Toda and Hyakumachi 2006; Qu et al. 2008a, b). The molecular nature of these mixed phenotypes of control of sexual propagation is currently unknown.

E. Agaricomycetes, Agaricales, Polyporales, and Russulales 1. Tetrapolar Species Generally for the ease of understanding the comparison of the mating type loci from the tetrapolar Agaricomycetes, a reader should note that for historial reasons the HD locus found to encode the HD1 and HD2 homeodomain transcription factors is known in the tetrapolar species as the A mating type locus, and the locus that encodes pheromone precursors and pheromone receptors (P/R) is known as the B mating type locus (Casselton and Olesnicky 1998). Accordingly, the alphabetical nomenclature of the two loci as compared to most Pucciniomycotina and Ustilaginomycotina is therefore unfortunately reversed. A novel unifying nomenclature for mating type loci is lately in use for all the basidiomycetes with HD as the “homeodomain” locus encoding HD1 and HD2 homeodomain transcription factors and P/R as the “pheromone” locus encoding pheromone precursors and pheromone receptors (Coehlo et al. 2010; Metin et al. 2010). This nomenclature has been adopted also in this review (see Sections IIA and II.B). It has the additional advantage that it can be applied to loci found together in one larger mating type region inherited as a single unit and to loci which during evolution lost mating type function.

Two properties are uniquely characteristic of the mating type loci of many heterothallic Agaricomycetes. First, a single locus may be hyperpolymorphic, determining the specificity of hundreds of mating types (Whitehouse 1948; Raper 1966; Table 6.1). Second, this hyperallelism is generated by duplication of mating type loci followed by the parallel evolution of subloci that perform redundant function. The process of recombination between subloci (each encoding dozens of specificities) then allows the true realization of hundreds of mating types to be achieved by combinations of alleles. The best studied species in the Agaricomycetes with regards to mating types are the two model species C. cinerea and

136

Ursula Ku¨es et al.

S. commune, where various allelic and paralogous mating type genes of highly diverse sequence have been cloned and functionally analysed by transformation into monokaryotic strains of compatible mating type (for the various A mating type genes, see Mutasa et al. 1989; May et al. 1991; Ku¨es et al. 1992, 1994a, b, c; Specht et al. 1992, 1994; Pardo et al. 1996; Shen et al. 1996, 2001; for the various B mating type genes, see Specht 1995; Wendland et al. 1995; Vallaincourt et al. 1997; O’Shea et al. 1998; Fowler et al. 1998, 2001, 2004; Halsall et al. 2000; Riquelme et al. 2005). In both species, a complex genetic structure with multiple HD1 and HD2 genes of paralogous function was established for the A mating type locus, and a complex genetic structure with multiple pheromone and pheromone receptor genes of paralogous function for the B mating type locus. With a completed genome now for both species, at least one (nearly) complete copy for both mating type loci is available (Ohm et al. 2010; Stajich et al. 2010; Fig. 6.13). Furthermore, a full set of sequences for a complete A and a complete B locus is available from the tetrapolar L. bicolor (Niculita-Hirzel et al. 2008), but a functional gene analysis is so far lacking. A partial A mating type locus from Coprinopsis scobicola with functional analysis of HD1 and HD2 genes has also been described (Ku¨es et al. 2001a). In both C. cinerea and S. commune, it was observed that directly linked to the A mating type locus is a gene mip for a mitochondrial intermediate peptidase of conserved sequence (Specht et al. 1994; Ku¨es et al. 1992; Isaya et al. 1995; Ku¨es et al. 2001a; Fig. 6.13). Classical genetic studies indicated that in many other Agaricomycotina, mip is also closely linked to the A mating type locus (James et al. 2004a) and molecular data from cloning and sequencing confirmed the direct linkage of A mating type loci to mip (Ku¨es et al. 2001a; James et al. 2004b, 2006; Martinez et al. 2004, 2009; Aimi et al. 2005; Niculita-Hirzel et al. 2008; UK, unpublished data). The proximal location of this gene helped in identifying the A mating type locus of C. scobicola (Ku¨es et al. 2001a) and provides a strategy for cloning A mating type loci for more species in the future (van der Nest et al. 2008). Whilst the easy to detect mip marks one border of A mating type loci, at the other end there is typically also a conserved gene, the intron-rich gene b-fg for the Ab-flanking gene (in C. cinerea it flanks the b sublocus of the subdivided A mating type locus; Ku¨es et al. 1992, 1994c; Fig. 6.13), for an unknown fungal protein (James et al. 2004b, 2006; Aimi et al. 2005; NiculitaHirzel et al. 2008; Martinez et al. 2009; Stajich et al. 2010; UK, unpublished data). The only known exception is S. commune where by chromosomal rearrangements the two subloci of the A mating type locus are separated from

each other by much larger distances (Ohm et al. 2010). Apparently during the course of extensive chromosomal rearrangements in S. commune, the b-fg gene has been translocated to another place in the same chromosome 840 kb downstream of the Aa sublocus (UK, unpublished data). Unfortunately for the chromosomal region surrounding the B mating type locus, their is little evidence of synteny that could be used for positional cloning (Niculita-Hirzel et al. 2008). For cloning pheromone receptor genes from B mating types (see the NCBI database for sequences of the Agaricomycetes mentioned in Table 1), one option is the use of degenerate primers targeting regions of higher sequence conservation. However, the PCR primers do not work on all species (James et al. 2004b; Li et al. 2007; van der Nest et al. 2008). Even between very closely related species, the outcome of such approach can however be in one case positive and in the other negative (van der Nest et al. 2008).

For the A mating type locus in the saprotrophic dung fungus C. cinerea, an idealized archetype has been postulated with three different functionally redundant and divergently transcribed HD1 and HD2 gene pairs of positional conservation: the a gene pair, the b gene pair and the d gene pair (Ku¨es and Casselton 2003; Pardo et al. 1996; for sequences of encoded homeodomains, see Fig. 6.1). The a gene pair is separated from the other gene pairs by 7 kb non-coding DNA of conserved sequence in which recombination leads to new gene pair combinations of novel mating type specificity, indicating the functional redundancy of paralogous gene pairs. The a pair corresponds to the classical Aa sublocus and the b and d gene pairs to the classical Ab sublocus of C. cinerea (Ku¨es et al. 1992, 1994c; May and Matzke 1995; Lukens et al. 1996). So far, the idealized archetype has not been identified in any strain. The total number of homeodomain genes in characterized C. cinerea A mating type loci varies between four and seven that arise from the presence of between two to four HD1 genes and between two to three HD2 genes (Casselton and Ku¨es 1994; Ku¨es et al. 1994a–c; Pardo et al. 1996; Stajich et al. 2010; Fig. 6.13). All analysed loci appear to have at least one functional A mating type gene at the a, the b and the d gene pair (Pardo et al. 1996). In both the a and the d gene pairs, cases are known where the HD2 gene has been lost (Ku¨es et al. 1992, 1994c; Pardo et al. 1996). Moreover, the HD2 gene of the a gene pair has been inactivated by mutation in the A43 mating type locus (Ku¨es et al. 1994c) which is present in the sequenced C. cinerea strain Okayama 7 (Stajich et al. 2010). Functional redundancy between

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

different gene pairs and within a gene pair (Fig. 6.2) allows such losses of genes and gene functions without general loss of mating type function (Ku¨es and Casselton 1993). On the contrary, C. cinerea A mating type loci may also acquire additional genes. In the A42 mating type locus, an extra HD2 gene has been found next to the HD2 gene of the a gene pair (referred to as an e gene) and an extra HD1 gene (referred to as an c gene) in between the b gene pair and the d gene (Ku¨es et al. 1992; Casselton and Ku¨es 1994; Ku¨es et al., unpublished data). Apparently, the A mating type locus is a dynamic entity, growing and shrinking throughout the evolution of mating types. The observed patterns possibly reflect a balance between selection to increase mating type allele variation and selection to lower numbers of incompatible proteins that need to be produced. In the wood-rot S. commune, traditional genetics also established that the A locus consists of two subloci called Aa and Ab (Raper et al. 1960; Raper 1966; Fig. 6.13). Aa was analysed first and found to contain one HD1 and HD2 gene pair (Stankis et al. 1992). There are nine different alleles of the Aa in nature and all but one (Aa) contain the complete gene pair. Aa1 has only one HD1 gene but this one gene is enough to allow compatible interactions with the HD2 products of all other Aa alleles (Specht et al. 1992, 1994). From the Ab sublocus, only one HD2 gene has been cloned thus far (Shen et al. 1996). With the available S. commune genome (A43 mating type allele), in silico analysis revealed, however, a structure that is far more complex than the A locus of C. cinerea with at least three HD1 and five HD2 genes (Fig. 6.13). The structure of the Aa sublocus with one HD1 and HD2 gene pair was shown to be as predicted while the new genes were all situated at the Ab sublocus in a region of about 24 kb. At Ab, only one divergently transcribed gene pair is seen, one pair of genes is transcribed convergently and two HD2 genes are unpaired (Ohm et al. 2010). A region of the Ab sublocus is however still incomplete in sequence (http://genome.jgipsf.org/cgi-bin/browserLoad/?db¼Schco1) and it is thus still possible that more genes are present. The Aa and Ab subloci are 550 kb apart from each other (Ohm et al. 2010) allowing frequent recombination between the two A subloci (Raper 1966). In other tetrapolar species, the organization of the A mating type locus is less complicated. There is a single HD1-HD2 gene pair in the A locus of

137

the ectomycorrhizal species L. bicolor (NiculitaHirzel et al. 2008; Fig. 6.13) and in the A locus of the white rot fungus P. djamor (James et al. 2004b) and there are two or possibly three (incomplete) gene pairs at the A locus of the secondary homothallic species C. scobicola (Ku¨es et al. 2001a). Of these species, estimates of mating types have been given for P. djamor with 58 different A loci and 231 different B loci (James et al. 2004b). With only one gene pair, for multiple A mating type specificities P. djamor is thus forced to distinguish fully 57 heteroallelic combinations of HD1 and HD2 proteins as functional interactions while preventing the dimerization of homoallelic combinations. In comparison, in C. cinerea analysis of alleles for all gene pairs present in nature defined four different a gene pairs, 9–10 different b gene pairs and three different d gene pairs (May and Matzke 1995; Pardo et al. 1996; Badrane and May 1999; May et al. 1999). Due to the free recombination in between the gene pairs, this leads to a combination of 108–120 different A mating type specificities, which come close to the 160 different A mating type alleles estimated for C. cinerea to exist in nature (Raper 1966). With only one to two more a pair alleles or d pair alleles or three more b pair alleles, the estimated number of A mating type specificities would be reached. Thus, at most, only 13 different allelic gene pair combinations must functionally be distinguished in this species. The situation at the B mating type locus of tetrapolar species is also complex; based on genome sequences, one complete B mating type copy is available for C. cinerea, S. commune and L. bicolor (Niculita-Hirzel et al. 2008; Stajich et al. 2010; Ohm et al. 2010; Fig. 6.13). The B mating type locus of C. cinerea (B43 mating type allele) contains four different pheromone receptor genes (CcSte3.1, CcSte3.2a, CcSte3.2b, CcSte3.3) and two related putative pheromone receptor genes (CcSte3-2151, CcSte3-2153) are linked at a distance of 40 kb – whether these have B mating type function or reside outside the B mating type locus is not established (Stajich et al. 2010; Figs. 6.5 and 6.13). Genome sequencing revealed thus once again a structure that turned out to be more complex than a previous model published for a B locus that suggested only three paralogous groups of genes for a pheromone–pheromone receptor system. In this archetypal model, each group consists of one pheromone receptor linked

138

Ursula Ku¨es et al.

with two or three pheromone precursor genes (Halsall et al. 2000; Brown and Casselton 2001; Riquelme et al. 2005). In the sequenced B43 locus, a total of nine pheromone precursor genes were detected and these are linked to the group of four pheromone receptor genes (Stajich et al. 2010; Figs. 6.11C and 6.13), for three of which functional B mating type activity has previously been shown (Riquelme et al. 2005). Apparently, the gene order in the B43 locus is not found in easily recognized distinct chains of subloci that would allow easy prediction of affiliation of pheromone precursor genes to a pheromone receptor gene as previously envisaged in the model by Casselton and colleagues. Interestingly enough, however, the hypothesized archetypal structure of the C. cinerea B locus is observed at the B locus of L. bicolor (NiculitaHirzel et al. 2008). L. bicolor has three pheromone receptor genes (LbSte3.1, LbSte3.2, LbSte3.3) each linked with a single pheromone gene and at a distance of 6 kb there are two further pheromone receptor genes (LbSte3.4, LbSte3.6) that lack associated pheromone genes (Fig. 6.13). Phylogenetic analyses of the pheromone receptors (Fig. 6.5) suggests that in the paralogous genes in the B mating type loci of C. cinerea and L. bicolor arose before the species diverged as well as the two mating type linked pheromone receptor genes that are conserved in relative position (NiculitaHirzel et al. 2008). In S. commune, the B mating type locus is subdivided into the recombining subloci Ba and Bb (Raper 1966). Within the sequenced locus (B41 mating type specificity), there are only two pheromone receptor genes (Scbar3, Scbbr2), one in each sublocus. However, 20 kb upstream of the Ba locus there are two extra pheromone receptor genes (Scbrl2, Scbrl3) and another extra receptor gene (Scbrl1) is 25 kb downstream of the Bb locus (Ohm et al. 2010; Fig. 6.13). In phylogenetic analysis of the proteins, ScBar3 clusters with CcSte3.3 and LbSte3.3, ScBbr2 with CcSte3.1 and LbSte3.1, ScBrl2 with CcSte3-2151 and LbSte3.4 and ScBrl3 with CcSte3-2153 and LbSte3.6, respectively (Fig. 6.5). These pairs of related receptors between S. commune and C. cinerea and L. bicolor further push back the origin of these paralogues to before the Schizophyllum–Coprinopsis split. This clustering supports further that two pheromone receptor genes outside the defined B mating type loci have a common origin with the mating type

genes. Other pheromone precursor in the three species also seem to have a common origin with the mating type pheromone receptors but are more various in sequence (Fig. 6.5). In the Ba locus of S. commune are four postulated pheromone precursor genes (Ohm et al. 2010; Fig. 6.13), three of which are known to be functional (Fowler et al. 2004) and in the Bb locus are twelve postulated pheromone precursor genes (Ohm et al. 2010; Fig. 6.13), eight of which are known to be functional (Fowler et al. 2001, 2004). Intriguently, functional studies of S. commune pheromones indicated that several of them can activate both a Bb receptor and a Ba receptor originating from B loci of other specificity (Fowler et al. 2004). Of the products of the five newly postulated pheromone genes in the sequenced genome of S. commune, only one pheromone precursor (BP11) however resembles in length and general structure other fungal pheromone precursors (PF08015; http://pfam.sanger.ac.uk/family? entry¼PF08015) above the required CAAX motif at the C-terminus (see Fig. 6.11C).

From P. djamor, thus far one receptor gene (PdSTE3.3) has been fully sequenced that is linked to B mating type function and accompanied by a pheromone precursor gene (Fig. 6.11C). Partial sequences for two other genes (PdSTE3.1, PdSTE3.2) and progeny analysis for mating type function helped to establish that the B mating type locus of P. djamor is subdivided as in S. commune but with one pheromone receptor gene in one sub-locus and two pheromone receptor genes in the other sub-locus (James et al. 2004b). PdSTE3.3 is linked to the p21-kinase gene PdCLA4 (James et al. 2004b), which in S. commune lies downstream of the Scbrl1 gene (U. Ku¨es, unpublished data). The orthologous gene in C. cinerea is linked to the B mating type locus while in L. bicolor the orthologous gene is linked to a non-mating type pheromone receptor gene but not on the B mating type locus chromosome (Niculita-Hirzel et al. 2008). A thorough analysis of the genome structures of the sequenced species could reveal the origin and evolution of pheromone receptor genes outside of defined B mating type loci. C. cinerea and S. commune each have one complete extra pheromone receptor gene (CcSte3-7395, Scbrl4) that lie on a chromosome other than those that carry the B mating type locus, and L. bicolor has in total eight further pheromone receptor genes (LbSte3.5, LbSte3.7 to LbSte3-13) unlinked to the

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

B mating type locus, five of which appear to be nonfunctional because they encode truncated proteins. Some of these genes were shown to be expressed suggesting they may have a biological function and this may involve roles other than meditating mating type (Niculita-Hirzel et al. 2008; Ohm et al. 2010; Stajich et al. 2010). For such apparently non-mating type receptor genes, a function in vegetative mycelial interactions has for example been hypothesized (Ku¨es and Navarro-Gonza´lez 2009; for a further discussion, see Section V.E). As the example of C. neoformans var. neoformans showed, non-mating type pheromone receptors might be constitutively active (Hsueh et al. 2009; see Section V.A.) or might be activated by pheromones. Activating pheromones could come from the mating type loci or might be mating type-independent. Congruent with this latter assumption, gene families for pheromone-like peptides have been detected in all available Agaricales genomes (Niculita-Hirzel et al. 2008; Ku¨es and Navarro-Gonza´lez 2010; UK, unpublished data) but not yet in all of the other genomes currently available for other Agaricomycetes (UK, unpublished data). Lack of success in finding such genes in the genome of a species might not confirm a complete absence of such genes. Genes for small pheromone-like peptides can elude detection in automated genome annotations. Accordingly, many of the pheromone genes in the B mating type loci of the Agaricomycetes were manually annotated by methods such as translation of the DNA of the B mating type regions in all six possible frames and visual inspection of the resulting predicted amino acid sequences. Likewise, in the Agaricales the families of non-mating type pheromone precursor genes were manually identified by a Blast search the genomes with known basidiomycete pheromones at the lowest stringency and filtering out hundreds of false positives prior to de novo gene definition (Niculita-Hirzel et al. 2008; Ku¨es and Navarro-Gonza´lez 2010; UK, unpublished data). The recent definition of a Pfam domain for fungal pheromone precursors at the Sanger Institute (Hinxton, UK; PF08015; http://pfam.sanger.ac.uk/family? entry¼PF08015) should help future gene identifications in genomes via ab initio gene annotations. Ongoing manual analysis in S. commune revealed the presence of some genes for pheromone-like proteins that follow the PF08015 structure (Fig. 6.11C). However, they are highly divergent in sequence from the precursors of non-mating type pheromone-like peptides found in species of the Agaricaceae (UK, unpublished data; Fig. 6.11C).

2. Bipolar Species Considerable genomic and molecular data are available for four bipolar species, C. disseminatus, Phanerochaete chrysosporium, Pholiota microspora

139

(Pholiota nameko), and Postia placenta (Martinez et al. 2004, 2009; Aimi et al. 2005; James et al. 2006, 2011; Yi et al. 2009). Uncertainity exists in the literature as to status of breeding behaviour in P. chrysosporium and there are contradicting reports that the species is homothallic or bipolar heterothallic (Alic and Gold 1985; Alic et al. 1987; Thompson and Broda 1987; Krejcˆ´ı and Homolka 1991; Dhawale and Kessler 1993). A recent report by James et al. (2011) suggests a bipolar mating system in P. chrysosporium based on population genetic data and pairing studies; however facultative homothallism may exist because self-fertile single basidiospore isolates have been reported (Alic et al. 1987; Krejcˆ´ı and Homolka 1991). The ability for P. chrysosporium to fruit could be akin to monokaryotic fruiting observed in monokaryotic individuals in other heterothallic Agaricomycetidae without any outcrossing (Raper and Krongelb 1958; Esser and Stahl 1976; Esser and Meinhardt 1977; Esser et al. 1979; Julian et al. 1997), but whether this homothallic behaviour could occur in nature has little support given that wild isolates are consistently heterokaryotic (Gold and Alic 1993).

Unlike the situation in dimorphic fungi (i.e., those with a haploid uninuclear yeast phase and a filamentous dikaryotic phase; e.g., U. hordei described in Section IV.A, or C. neoformans described in Section V.A), bipolarity in the analysed filamentous Agaricomycetes has arisen from a tetrapolar system not by the fusion of two ancestral independent loci, but by the loss of the mating type determining function of the pheromone receptor and pheromone precursor genes. In C. disseminatus, sequences of multiple strains of P/R loci confirmed that this species is not polymorphic at any P/R locus, which would allow discrimination among different mating types (James et al. 2006). In three of the bipolar species, C. disseminatus, P. chrysosporium, and P. microspora, the functional mating type loci contain HD1 and HD2 genes with allele-specific sequence divergence and specific HD1 and HD2 alleles co-segregating in populations with control of mating function and dikaryon formation (James et al. 2006, 2011; Aimi et al. 2005). A genetic study from P. placenta linking specific genes to mating type behaviour is as yet not available. In C. disseminatus, the A mating type locus contains two pairs of mating type specific HD1 and HD2 genes, and these are directly flanked by the conserved mip and b-fg genes (James et al. 2006). C. disseminatus has at least four different pheromone receptor genes, all of which do not cosegregate with the mating type locus. CDSTE3.1 and CDSTE3.3 are clearly linked with about 15 kb

140

Ursula Ku¨es et al.

in between, and there is one pheromone precursor gene (CDPHB1) upstream of CDSTE3.1 and two pheromone precursor genes (CDPHB2.2, CDPHB2.1) in proximity to CDSTE3.3 (James et al. 2006; Fig. 6.11D). In cluster analysis, all four C. disseminatus pheromone receptors group with B mating type pheromone receptors of tetrapolar species (James et al. 2006; NiculitaHirzel et al. 2008), strongly suggesting for the C. disseminatus genes a common origin with B mating type genes of the tetrapolar species. Genes of the C. disseminatus B mating type-like locus are active in B-regulated development when heterologously expressed in C. cinerea, further supporting a common origin and proving that the genes are likely still involved in sexual development (James et al. 2006). In the published genome of P. chrysosporium strain RP78, there is an HD1 and HD2 gene pair that lies between the mip and b-fg genes on scaffold 13. Four potential pheromone receptor genes (three in a cluster) are on scaffold 9, and an additional one lies on scaffold 5 (Martinez et al. 2004; Niculita-Hirzel et al. 2008; James et al. 2011). Scaffold 5 also bears a sequence for a putative pheromone precursor but at a distance of ~500 kb from the pheromone receptor gene. Four genes for pheromone-like peptides are found on scaffold 3 (Fig. 6.11D). Phylogenetic analysis of the sequences of the pheromone receptors indicated a relationship between P. chrysosporium receptors to both B mating type specific and non-mating type specific pheromone receptors of the tetrapolar species C. cinerea and L. bicolor (NiculitaHirzel et al. 2008). In P. placenta, a dikaryon was used for the genome sequence and thus allelic loci are available (Martinez et al. 2009). A DNA region found in two allelic forms of low sequence homology is present that resembles the A mating type locus of tetrapolar species. This region is flanked by the mip and b-fg genes. Evidence for both an HD1 gene and an HD2 gene can be deduced from the sequences of both of the allelic regions but complete HD1 gene models are not available due to poor sequence quality due to transposon insertions and extreme sequence divergence within the candidate A mating type locus. Usually, mip is located convergent to the adjacent HD1 gene and this arrangement is likely to be the ancestral arrangement. Exceptions from the rule are P. placenta

(Martinez et al. 2009) and P. chrysosporium (NiculitaHirzel et al. 2008) where the single HD1 and HD2 gene pair has been inverted with respect to the neighbouring mip and b-fg genes.

Another P. placenta chromosomal region present in two allelic forms resembles the B mating type loci of tetrapolar species (Martinez et al. 2009). In this ~55 kb region, there are six and seven pheromone receptor genes and ten and 13 pheromone precursor genes within the two allelic regions, respectively. Most, but not all, allelic pheromone receptor genes in this region are quite similar to each other in sequence as are many of the pheromone precursors. In addition, two other pheromone receptor genes in the P. placenta genome are unlinked to this locus. Comparison of all the pheromone receptors of P. placenta with those from C. cinerea revealed that the larger P. placenta locus with the several pheromone receptor genes encodes homologues of the B mating type locus of C. cinerea, suggesting that the genes represent ancestral duplications prior to speciation in the Agaricomycetes (Martinez et al. 2009). According to Nobles (1943), P. placenta is bipolar. Without knowledge on the segregation of the A and B mating type-like chromosomal regions, the current sequence data showing sequence dissimilarity at the putative mating type loci does not allow the resolution of the mechanistic basis of bipolarity in this fungus. In P. microspora, the structure of the A mating type locus is known in addition to a pheromone receptor gene that does not co-segregate with mating type. The A locus consists of a single HD1 and HD2 gene pair flanked by a mip gene and a b-fg gene (Aimi et al. 2005; Yi et al. 2009). An interesting physiological study on dosage effects of the two different A mating type genes in transformants suggests that A mating type pathway activation by one compatible gene pair is sufficient for production of clamp cells, but clamp cell fusion requires activation by both the alternate compatible HD1 and HD2 pairs (Fig. 6.2) or higher expression levels of one gene as shown by altering its promoter (Yi et al. 2010). Apparently, clamp cell fusion requires a certain threshold of HD proteins. However, this result does not mean that the A mating type proteins in the bipolar species now have the same function that pheromones and pheromone receptors exhibit in the tetrapolar species. B mating type genes in

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

tetrapolar species regulate clamp cell fusion (Raper 1966; Casselton and Olesnicky 1998), but it is unknown whether clamp cell fusion in bipolar species is affected directly by pheromone receptors or indirectly via activation of the HD1 and HD2 genes. Indeed, B-regulated clamp cell fusion always occurs after A-induced clamp cell production in tetrapolar species (Ku¨es et al. 2001b, 2002; Badalyan et al. 2004), and it is therefore an open question whether the A mating type proteins possibly contribute to clamp cell fusion in either tetrapolar or bipolar species. In U. maydis, the pheromone and pheromone receptor system regulates the expression of the HD1 and HD2 genes via a signalling chain ending at the transcription factor Prf1 (Hartmann et al. 1996; Kaffarnik et al. 2003; Mendoza-Mendoza et al. 2009). In C. cinerea, the activated B mating type pathway controls the strength of functions exerted by the A pathways. The activated B pathway enhances light-regulated initiation of A mating type-induced fruiting body initiation and counteracts the negative lighteffect on A mating type-mediated repression of asexual sporulation (Ku¨es et al. 2002a, b, 2004). The data suggest a fine-tuning of control of A mating type functions by the B mating type pheromone response pathway, possibly by regulation of their transcription. In P. microspora, another mechanism of ensuring correct HD1 and HD2 product levels within dikaryotic cells (other than the normal pheromone-response pathway) may be one solution to the loss of B mating type function for pheromone receptor and pheromone precursor genes. Alternatively, the B pathway in bipolar species could function by constitutive expression or by differential and transient expression of the pheromones and receptors that allow communication between different mating type nuclei. 3. Homothallic Species A partial genome sequence is available (Mondego et al. 2008) for the homothallic cacao pathogen Moniliophthora perniciosa (Griffiths and Hedger 1994). Inspection of the partial genome revealed sequences for one or possibly two HD1 genes, seven or possibly eight genes for potential pheromone receptors and five genes for putative pheromone precursors (Mondego et al. 2008; Ku¨es and Navarro-Gonza´lez 2010). As much as the incomplete sequences for the products of pheromone

141

receptor genes allow an analysis by phylogenetic comparisons, each protein clusters with pheromone receptors from the B mating type gene subgroup 1 or from the B mating type gene subgroup 3 of the Agaricales. Five others cluster independently from mating type gene products in a manner suggesting that they all have a common origin with the putative non-mating type pheromone receptor LbSTE3.4 of L. bicolor. Of the pheromone precursors, two variants resemble more the mating type pheromone precursors of other Agaricales and three variants resemble the precursors of non-mating-type pheromone-like peptides of the Agaricales (Ku¨es and Navarro-Gonza´lez 2010; Fig. 6.11E). Currently, it is unknown whether the mating type orthologues in M. perniciosa have a post-mating function, such as in the establishment of the dikaryon-like mycelium upon basidiospore germination and in mushroom production (Griffiths and Hedger 1994). Homothallism is unlikely to be generated by just loss of mating type genes since in C. cinerea a mutant missing A mating type genes and in S. commune a mutant without the typical B mating type locus have been generated but are self-sterile and morphologically similar to monokaryons (Pardo et al. 1996; Fowler et al. 2001, 2004). Instead, constitutive self-compatibility in M. perniciosa might be mediated by mutations within the mating type genes that unlink mating, i.e., fusion of different individuals and incidental outcrossing, from sexual development, i.e., mushroom and basidiospore development. Rare cases of such mutations in A or B mating type loci of C. cinerea and S. commune are reported (Raper et al. 1965; Swamy et al. 1984): HD1-HD2 fusion genes combine incompatible HD1 and HD2 proteins into a functional pseudo-heterodimer (Ku¨es et al. 1994b; AsanteOwusu et al. 1996; Pardo et al. 1996). Certain mutations in pheromone precursor genes or pheromone receptor genes allowed their products to react with normally incompatible homoallelic products encoded in the same haploid genome while other mutations in pheromone receptors made them independent from any pheromone stimulation (Olesnicky et al. 1999, 2000; Fowler et al. 2001). Moreover, while in C. cinerea mating type genes control mating and sexual development (Tymon et al. 1992; Ku¨es et al. 1998, 2002), mutations in certain other genes downstream of the mating type genes can make sexual

142

Ursula Ku¨es et al.

reproduction independent from mating (e.g., see Murata et al. 1998; Murata and Kamada 2009). Unpublished work from Burrow, Casselton and Challen suggests that mating type-like genes are also present in homothallic Agaricus species (Casselton and Challen 2006).

F. Evolutionary Considerations In the Agaricomycotina, there are two interesting evolutionary routes leading from tetrapolarity to bipolarity: One route, known already from several species in the Pucciniomycotina and the Ustilaginomycotina, involves fusion of two formerly independent loci into one contiguous locus with both HD and P/R genes remaining functional in mating type determination. The other route proceeds via inactivation of the mating type function at the P/R locus, i.e., reducing the number of genes needed for mating type determination. From the perspective of outcrossing potential, the two situations may appear to yield the same outcome. However, the two routes may have different consequences because the ability to maintain multi-allelism in the fused situation is more costly than the situation in which one locus is liberated from mating type function. In the fused case, each component of the fused locus would be required to evolve in concert, necessitating parallel mutations in HD1, HD2, pheromone and pheromone receptor genes. In line with this consideration, the characterized bipolar species of the basidiomycetes with fused functional loci are all biallelic, excepting the pseudo-bipolar Sporidiobolales (Table 6.1) which may be a transitory mating system or may be an evolutionary stable system reaping the benefits offered from both tetrapolar and bipolar systems (see Sections III.D and IV.D). The fusion of two independent mating type loci together into one locus requires blocks to recombination and ultimately the evolution of sex chromosomes, as seen in a number of Filobasidiales (see Section V.A), certain Microbotryales (see Section III.B), and Ustilaginales (see Section IV.A). As a consequence, inbreeding potential increases but also outbreeding potential will actually increase. The reason for an increase in outbreeding potential is that the bipolar, biallelic mating system of the fused HD and P/R loci has an outbreeding potential of exactly 50%. In con-

trast, the modified tetrapolar mating system possessed by the ancestors of those bipolar fungi with unfused loci is less than 50% (Fig. 6.12; see Section IV.C). Whether bipolarity predisposes a lineage to further evolving homothallic mating where inbreeding potential greatly increases has long been speculated (Raper 1966), but there are no clear demonstrations that this is the case. Whether generation of homothallism is a one step development from bipolarity or whether it also can directly come in one step from tetrapolarity is not known. As available genome data show, homothallism as well as asexuality do not essentially require that there are no mating type-like genes (see Sections III.A and V.E.3). It remains to be elucidated in these homothallic species whether previous mating type genes maintain similar functions as in heterothallics. In the bipolar Agaricomycetes characterized thus far, in all cases the P/R locus (corresponding to the B mating type locus) is unlinked to the A mating type locus and has no mating type function. This might, however, be a coincidence attributable to the low number of systems analysed to date. A genetic analysis of P. placenta, for example, might either reveal a bipolar system based on linkage of the A and B mating type genes of different allelic sequences or it might reveal the P/R locus to encode mating incompatibility rather than the HD locus. Regardless of whether bipolar Agaricomycetes species with fused A and B mating type loci might exist, inactivation of one mating type locus with a second locus having multiple specificities at a smaller number of genes minimizes the number of compatible gene interactions that must be maintained for mediating outcrossing and restricting inbreeding. Principally, such inactivation should be possible for both types of loci with similar costs, but from the perspective of biochemical interactions the HD and P/R gene products may have very different constraints on numbers of allelic interactions. Surprisingly, genomics has revealed that Agaricomycotina harbor non-mating type pheromone receptor genes and, at least in the Agaricales, non-mating type pheromone precursor genes are often also present (V.E). Several are known to be expressed (Niculita-Hirzel et al. 2008; Hsueh et al. 2009; Ohm et al. 2010), suggesting that these indeed have a function. In C. neoformans var. neoformans the constitutive non-mating type pheromone receptor CnnCpr2 can drive an

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

inbreeding-homothallic unisexual reproduction process between strains of the same mating type (Hsueh et al. 2009). By analogy, the non-mating type pheromone receptors and pheromones in Agaricales might play novel roles during monokaryotic fruiting. However, where monokaryotic fruiting has been described in species of Agaricales with a multi-allelic tetrapolar mating type system, these were due to mutations in individual strains (Raper and Krongelb 1958; Esser and Stahl 1976; Esser and Meinhardt 1977; Esser et al. 1979). Nonmating type pheromone receptors and pheromones might also operate during general cellular communication pathways similar to quorum sensing in bacteria and fungi (Ku¨es and NavarroGonza´lez 2009). In the filamentous Agaricomycetes, cellular fusion can occur independently of mating type and the functions of mating type genes only become apparent following cell fusion (Raudaskoski 1998; Hiscock and Ku¨es 1999). Nevertheless, some type of recognition process must occur between different individuals of a species (or between hyphae fusing within mycelial colonies) to ensure species discrimination, and these novel non-mating type specific pheromone receptors may function in this process (Ku¨es and NavarroGonza´lez 2009). Alternatively, in the complex interspecies milieu found in natural niches, these non-mating type specific receptors may allow detection of pheromones from closely or distantly related species as a form of cross-species eavesdropping, and this may involve pheromones that act as receptor agonists of inverse agonists that reduce signalling for example by constitutively active receptors such as CnnCpr2. Precedents for cross-species inhibition by pheromones have so far been reported within the Ustilagomycetes and Exobasidiomycetes (Sherwood et al. 1998; Kosted et al. 2002). In the saprotrophic C. cinerea and the ectomycorrhizal species L. bicolor, the A mating type locus is found on the largest chromosome I. Large regions of this chromosome, including that with the A mating type locus, are highly conserved in gene order between the two species. The genes in the conserved region are mostly found in a single copy within the respective genomes unlike genes located in non-conserved regions. Gene distances are very short and many of the genes have housekeeping functions. Moreover, recombination is reduced in this large region of synteny. All these observations together argue for an

143

ancient and stable gene content on chromosome I of the two species in which the A mating type genes have been embedded in and are protected from frequent rearrangements (Niculita-Hirzel et al. 2008; Stajich et al. 2010). In additional species for which data on the genes surrounding the A mating type loci are available (P. djamor; C. disseminatus, P. chrysosporium), a similar synteny is seen beyond the two genes mip and b-fg (James et al. 2004b, 2006; Niculita-Hirzel 2008). Only S. commune deviates from the gene order found in C. cinerea and L. bicolor due to massive chromosomal rearrangements as suggested from classical genetics (Raper 1966; Ku¨es et al. 2001b). Despite the massive rearrangements, most genes have remained on the same chromosome as the A mating type locus (ongoing analysis of the S. commune genome by UK). The chromosomal arrangement of B mating type loci in Agaricales is quite different. Comparisons of B mating type locus chromosomal regions between C. cinerea and L. bicolor show only a few common genes, a low gene content, high frequency of gene duplications, high frequency of inactivated genes (pseudogenes) and a high frequency of transposon insertions and transposon remnants (Niculita-Hirzel et al. 2008). Although not yet thoroughly analysed for other species, a low overall level of conservation is also suggested among other species (UK, unpublished data). Data suggest that the B mating type locus is generally of younger age (Niculita-Hirzel et al. 2008) or subject to higher rates of evolutionary change. This is consistent with the recent observation that close to the B mating type locus in C. cinerea is a recombination hotspot (Srivilai 2006) and that the B mating type chromosome is large based on physical map size (North 1990) but comparably small by total base length (O’Shea et al. 1998; Stajich et al. 2010). In S. commune, based on weak linkage to the ura1 gene observed in classical genetic analyses, the B mating type was assigned together with ura1 onto a large chromosome based on recombination units (Raper 1988, 1990). In contrast, in another mapping study these genes were assigned to different chromosomes (Frankel and Ellingboe 1977), suggesting that the B mating locus might be localized close to a chromosomal end that has been recently translocated. Hybridization studies with subcloned genes confirmed the localization to different chromosomes, and that the B mating type

144

Ursula Ku¨es et al.

locus is localized on small-sized chromosomes ´ sgeirsdo´ttir et al. 1994). Moreover, highly vari(A able recombination frequencies (between 0–8.9%) are observed between the two B mating type subloci of S. commune (Raper et al. 1958; Koltin et al. 1967) within a short DNA region characterized by repeated sequences (Fowler et al. 2004; Ohm et al. 2010). Due to increased rates of genome rearrangement in the region, the B mating type genes might more frequently become inactive or self-compatible by linkage of pheromones with compatible receptors in the same haplotype. Recombination within subdivided B mating loci also contributes to multiplication of mating type specificities similar to subdivided A mating type loci (Raper et al. 1958, 1960; Day 1960; Koltin et al. 1967). In terms of evolutionary constraints, the divergence of sequences resulting in different mating type specificities is a daunting process. The maximum possible number of functional alleles per mating type gene is naturally limited because: (1) each allele must be capable of recognizing self from non-self, and (2) the number of non-self alleles that must each be recognized is potentially more than a dozen other alleles. Moreover, at the same time divergence of gene sequences must preserve functionality of encoded products and not result in functional interaction between products of the same mating type that would cause self-fertility. The available molecular data show that different trajectories evolved among basidiomycetes in order to increase the number of different mating type specificities and to bring variety into the system: diversification of the alleles of given genes and the acquisition of paralogous genes with diverged alleles. In C. cinerea and S. commune, there are an estimated 160 and 288 different functional versions of the A mating type locus, respectively and in both about 81 different functional versions of the B mating type locus (Raper 1996). Taking the homeodomain transcription factors as an example, with the N-terminal specificity domains of ~120 amino acids in length (Ku¨es et al. 1994a; Banham et al. 1995; Asada et al. 1997; Yue et al. 1997) and the possibility to incorporate at each position 20 different amino acids, mathematically many different versions can be envisaged. However, there are two strong constraints on variability that arise from the need to: (1) conserve a specific domain structure (which is predicted to be helical; Banham et al. 1995; Niculita-Hirzel et al. 2008),

and (2) ensure that appropriate amino acids are presented at the interfaces of the dimerization domains to enable interaction of the potential partner proteins (Banham et al. 1995). Therefore, duplication of mating type genes and divergence into functionally independent genes (paralogous genes) is a recent evolutionary innovation so far specific to Agaricales as an efficient mechanism for dramatically increasing mating type specificities. In this manner the mushroom mating system ensures a high level of outbreeding (Fig. 6.12) while maintaining an inbreeding restriction of 25%.

VI. Conclusions In all three subphyla of the Basidiomycota, heterothallic bipolar, heterothallic tetrapolar, and homothallic species have been described, and in all subphyla, there are evidently also anamorphic (asexual) species. In every species investigated, genes for HD1 and HD2 homeodomain transcription factors are present as are genes for pheromone receptors and pheromone precursors homologous to the a-factor and Ste3 receptor of S. cerevisiae. From this observation, it may be inferred that the ancestor of the basidiomycetes possessed all these genes, but lacked the a-factor and Ste2 receptor. Casselton (2008) and Lee et al. (2010) hypothesized that the acquisition of the HD1 and HD2 homeodomain transcription factors occurred in the ancestor of the Dikarya before the split of the asco- and basidiomycetes. This scenario appears indeed less demanding since it requires only once the evolution of two types of homeodomain transcription factors, classical homeodomain transcription factors and TALE proteins that are of different ancient lineage (Derelle et al. 2007; Mukherjee et al. 2009). Ascomycetes encode Ste3-type pheromone receptors and lipopeptide pheromone precursors, although in the ascomycetes their genes are not mating type genes (see Chapter 5 in this volume). Thus, after the divergence of the Dikarya into Ascomycota and Basidiomycota, the basidiomycete lineage appears to have recruited the Ste3 pheromone receptor genes and pheromone precursor genes into a second mating type locus. Given its ubiquity, the ancestral mating type configuration in the Fungal Kingdom is likely to have been the bipolar state, although arguments

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality

might also be marshalled in favour of an ancestral unipolar state. For example, there is no report of mating types in the basal-most phylum of fungi, the Chytridiomycota (Idnurm et al. 2007). Thus far, the unusual tetrapolar state appears to be unique to the Basidiomycota, and thus the most parsimonious model is that it arose once within this phylum, although independent derivations are formally possible. It is currently equivocal whether the common ancestor of the Basidiomycota had a tetrapolar mating system, as is the deep branching of the Basidiomycota phylogeny (Liu et al. 2009; McLaughlin et al. 2009). Understanding whether tetrapolarity was ancestral in Basidiomycota will require identifying the mating type genes in the tetrapolar Pucciniomycotina (the putative basal lineage), and reconstructing the phylogenetic history of these mating type locus arrangements. If tetrapolarity evolved before the evolution of multiple alleles (greater than two alleles per locus), selection pressure for increasing outbreeding efficiency may have driven the fusion of the two loci into a bipolar system and/or in producing multiple mating type specificities in one or in both of the unlinked mating type loci. Only in the more derived lineage with the larger fruiting bodies (Agaricomycotina) has the evolution of two multi-allelic mating type loci been observed along with concomitant extreme outcrossing efficiency. Bipolarity has clearly been derived several times independently within different lineages of the Ustilaginomycotina and the Agaricomycotina from tetrapolarity and possibly also in the Pucciniomycotina (Bakkeren et al. 2008; see earlier in this chapter). Mechanisms by which bipolarity evolved include fusion of the P/R locus with the HD locus in all species that had a modified tetrapolar mating system. Alternatively in the Agaricomycotina that have two multi-allelic mating type loci, no fusion is observed but rather a release of the P/R locus from encoding mating type specificity. As shown in C. disseminatus (James et al. 2006), loss of mating type function in a bipolar species does not automatically abolish all cellular functions for these genes: despite loss of mating type function, pheromone receptor genes and pheromone precursor genes could still promote dikaryon stability, fructification, and sporulation when expressed in a heterologous host. While in such a bipolar system one locus lost mating type function, this route allows the continued

145

inheritance of multiple mating types without the collapse suggested for the fusion origin of bipolarity (Hsueh et al. 2008). This ensures still a very high outbreeding potential and 50% inbreeding avoidance, unlike the bipolar–biallelic situation that has an outbreeding potential of only 50% and shows inbreeding avoidance of also 50% (see Fig. 6.12 for the outbreeding potentials). Coehlo et al. (2010) recently defined pseudobipolarity from a basal subphylum of the Pucciniomycotina (Sporidiales). In the species with pseudo-bipolarity, all mating type genes are linked on one chromosome, but recombination between the HD1 and HD2 genes and the pheromone receptor and pheromone precursor genes is allowed. Strikingly, also in the system of pseudobipolarity, there are only two versions of pheromone precursor and pheromone receptor genes but multiple alleles of the HD1/HD2 gene pair. Coehlo et al. (2010) discuss their data in view of bipolarity being the ancient situation within the basidiomycetes. However, emerging data for the Pucciniomycotina suggest that also in this subphylum bipolarity has evolved more than once. Generally, a multitude of different insights on mating type genes and their evolution have now been revealed in different basidiomycete species using both model and non-model species and molecular and evolutionary genetics. Most of this has been facilitated by the ability to generate large amounts of sequence data that have revealed unforeseen insights into basidiomycete mating genes. For example, genome sequencing has revealed large MAT loci in U. hordei, C. neoformans and other bipolar species that contain mating type genes as well as other genes acting in mating type pathways and are analogous to rapidly evolving sex chromosomes; these mating type loci, like sex chromosomes, are also subject to pseudogene formation and invasion by transposable elements because of suppression of recombination (see Sections III.B and V.A). In the tetrapolar species in contrast, the genomic context of the HD and P/R loci are free to evolve with respect to the association with conserved genes, but appear to do so in rather different ways. It is striking that in the Agaricomycetidae, apparently the A mating type locus with the HD1 and HD2 genes has a very stable chromosomal location (typically the largest chromosome) and conserved synteny, whereas the B mating type locus can be found in

146

Ursula Ku¨es et al.

genome regions of low overall gene conservation, low gene density, high level of defective genes and high frequency of transposon insertions. Loss of mating type function for the B mating type locus in bipolar Agaricomycetides might thus be related to the frequency of recombination in this region. The existing data suggest that multiple mating types have evolved only once in fungi, being restricted to Basidiomycota. This plethora of mating type genes has led to a long unresolved question regarding how exactly do novel mating type specificities evolve? Many researchers have tried to generate novel mating types through mutagenesis with no success (Casselton and Ku¨es 2007). What they were able to generate were self-compatible alleles, however, and this led to important inroads to understanding the evolution of bipolarity and homothallism (Ku¨es et al. 1994b; Olesnicky et al. 1999). The role of recombination versus point mutation (i.e., punctuated versus gradual) in generating novel alleles is unclear. All species investigated have mating type alleles that differ greatly in sequence, thus obscuring the evolutionary events that led to the diversification of alleles (Stankis et al. 1992; Specht et al. 1994; May et al. 1999). So little is known that even in the case of S. reilianum with only three alleles, horizontal transfer has been evoked to explain a pattern that is clouded in rearrangement, gene loss, and large sequence divergence (Schirawski et al. 2005; Bakkeren et al. 2008). Knowledge of the diversification process and its clear relationship with mitochondrial inheritance (Fedler et al. 2009) has great promise to unravel one of the great mysteries in evolutionary biology: the origin of more than two sexes (Hurst 1996; Billiard et al. 2011). Here we have introduced a new term, unipolar, to describe the unisexual mode of fungal reproduction that is distinguished from the bipolar and tetrapolar configurations. Unipolar sexual reproduction is homothallic and involves a single mating type and the production of meiotic progeny that are also all of the one parental mating type. Thus, all of the progeny are interfertile with each other as well as all other members of the population because mating type compatibility is not required for mating. Unipolarity, whether facultative or strict, could allow the maximal outbreeding potential of 100% to be achieved (Fig. 6.12). Unipolar species may include those that are classically considered homothallic as well as species such as C. neoformans that is

facultatively unipolar. The existence of facultative unipolarity within heterothallic species may have been largely ignored in the past but merits further investigation. Acknowledgements Marco A. Coelho is gratefully acknowledged for providing data on Sporidiobolales prior to publication and Marianna Feretzaki for generating Fig. 6.10. The JGI and the Broad-Institute and their staff are thanked for providing genome sequences of various basidiomycetes to the public and all colleagues in the genome annotation teams for any input in definition of mating type genes. This effort was supported in part by R01 grants AI39115 and AI50113 from the NIH/NIAID to J.H.

References Abbate JL, Hood ME (2010) Dynamic linkage relationships to the mating-type locus in automictic fungi of the genus Microbotryum. J Evol Biol 23:1800–1805 Abe K (1987) Differentiation of mating tubes induced by the mating hormone Rhodotorucine A during the mitotic cycle of Rhodosporidium toruloides. J Gen Appl Microbiol 33:437–444 Abe K, Kusaka I, Fukui S (1975) Morphological change in the early stages of the mating process of Rhodosporidium toruloides. J Bacteriol 122:710–718 Aimi T, Yoshida R, Ishikawa M, Bao DP, Kitamoto Y (2005) Identification and linkage mapping of the putative homeodomain protein (hox1) and the putative pheromone receptor homologue (rcb1) in a bipolar basidiomycete, Pholiota nameko. Curr Genet 48:184–194 Akada R, Minomi K, Yamashiata I, Miyakawa T, Fukui S (1987) Cloning of a gene coding for Rhodotorucine A, a farnesyl peptide mating pheromone of Rhodosporidium toruloides. Agric Biol Chem 51:1211–1215 Akada R, Kaj J, Yamashita I, Miyakawa T, Fukui S (1989a) Genomic organization of multiple genes coding for Rhodotorucine A, a lipopeptide mating pheromone of the basidiomycetous yeast Rhodosporidium toruloides. Arch Microbiol 152:484–487 Akada R, Minomi K, Kai J, Yamashita I, Miyakawa T, Fukui S (1989b) Multiple genes coding for precursors of Rhodotorucine A, a farnesyl peptide mating pheromone of the basidiomycetous yeast Rhodosporidium toruloides. Mol Cell Biol 9:3491–3498 Albert HH, Schenck S (1996) PCR amplification from a homolog of the bE mating-type gene as a sensitive assay for the presence of Ustilago scitaminea DNA. Plant Dis 80:1189–1192 Alby K, Schaefer D, Bennett RJ (2009) Homothallic and heterothallic mating in the opportunistic pathogen Candida albicans. Nature 460:890–893 Alic M, Gold, MH (1985) Genetic recombination in the lignin-degrading basidiomycete Phanerochaete chrysosporium. Appl Environ Microbiol 50:27–30 Alic M, Letzring C, Gold MH (1987) Mating system and basidiospore formation in the lignin-degrading

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality basidiomycete Phanerochaete chrysosporium. Appl Env Microbiol 53:1464–1469 Allen RF (1932) A cytological study of heterothallism in Puccinia triticina. J Agric Res 44:733–753 Allen RF (1934) A cytological study of heterothallism in Puccinia sorghi. J Agric Res 49:1047–1069 Alspaugh JA, Perfect JR, Heitman J (1998) Signal transduction pathways regulating differentiation and pathogenicity of Cryptococcus neoformans. Fungal Genet Biol 25:1–14 Anderson CM, Willits DA, Kosted PJ, Ford E, MartinezEspinoza AD, Sherwood JE (1999) Molecular analysis of the pheromone and pheromone receptor genes of Ustilago hordei. Gene 240:89–97 Anikster Y, Eilam T, Mittelman L, Szabo LJ, Bushnell WR (1999) Pycnial nectar of rust fungi induces cap formation on pycniospores of opposite mating type. Mycologia 91:858–870 Anikster Y, Eilam T, Bushnell WR (2000) Interspecific transfer of pycnial nectar induces pycniospore caps in rust fungi in a manner related to mating type within species. Mycol Res 104:311–316 Arcangioli B, Thon G (2004) Mating-type cassettes: structure, switching and silencing. In: Egel R (ed) The molecular biology of Schizosaccharomyces pombe. Springer, Berlin Heidelberg New York, pp 129–148 Arkhipova I, Meselson M (2005) Deleterious transposable elements and the extinction of asexuals. BioEssays 27:76–85 Asada Y, Yue CL, Wu J, Shen GP, Novotny CP, Ullrich RC (1997) Schizophyllum commune Aa mating-type proteins, Y and Z, form complexes in all combinations in vivo. Genetics 147:117–123 Asante-Owusu RN, Banham AH, Bo¨hnert HU, Mellor EJ, Casselton LA (1996) Heterodimerization between two classes of homeodomain proteins in the mushroom Coprinus cinereus brings together potential DNAbinding and activation domains. Gene 172:25–31 ´ sgeirsdo´ttir SA, Schuren FHJ, Wessels JGH (1994) A Assignment of genes to pulse-field separated chromosomes of Schizophyllum commune. Mycol Res 98:689–693 Bachtrog D (2006) A dynamic view of sex chromosome evolution. Curr Opin Genet Dev 16:578–585 Bachtrog D (2008) The temporal dynamics of processes underlying Y chromosome degeneration. Genetics 179:1513–1525 Badalyan S, Polak E, Hermann R, Aebi M, Ku¨es U (2004) Role of peg formation in clamp cell fusion of homobasidiomycete fungi. J Basic Microbiol 44:167–177 Badrane H, May G (1999) The divergence-homogenization duality in the evolution of the b1 mating type gene of Coprinus cinereus. Mol Biol Evol 16:975–986 Bai F-Y, Zhao J-H, Takashima M, Jia J-H, Boekhout T, Nakase T (2002) Reclassification of the Sporobolomyces roseus and Sporidiobolus pararoseus complexes, with the description of Sporobolomyces phaffii sp. nov. Int J Syst Evol Microbiol 52:2309–2314 Bakkeren G, Kronstad JW (1993) Conservation of the b-mating-type gene-complex among bipolar and tetrapolar smut fungi. Plant Cell 5:123–136

147

Bakkeren G, Kronstad JW (1994) Linkage of mating-type loci distinguishes bipolar from tetrapolar mating in basidiomycetous smut fungi. Proc Natl Acad Sci 91:7985–7089 Bakkeren G, Kronstad JW (1996) The pheromone cell signalling components of the Ustilago a mating-type loci determine intercompatibility between species. Genetics 143:1601–1613 Bakkeren GM, Kronstad JW (2007) Bipolar and tetrapolar mating systems in the Ustilaginales. In: Heitman J, Kronstad JW, Taylor JW, Casselton LA (eds) Sex in fungi. Molecular determination and evolutionary implications. ASM Press, Washington, DC, pp 389–404 Bakkeren G, Gibbard B, Yee A, Froeliger E, Leong S, Kronstad J (1992) The a and b loci of Ustilago maydis hybridize with DNA sequences from other smut fungi. Mol Plant Microb Interact 5:347–355 Bakkeren G, Jiang G, Warren RL, Butterfield Y, Shin H, Chiu R, Linning R, Schein J, Lee N, Hu G, Kupfer DM, Tang Y, Roe BA, Jones S, Marra M, Kronstad JW (2006) Mating factor linkage and genome evolution in basidiomycetous pathogens of cereals. Fungal Genet Biol 43:655–666 Bakkeren G, Ka¨mper J, Schirawski J (2008) Sex in smut fungi: Structure, function and evolution of mating– type compleses. Fungal Genet Biol 45:S15–S21 Banham AH, Asante-Owusu RN, Go¨ttgens B, Thompson SAJ, Kingsnorth CD, Mellor EJC, Casselton LA (1995) An N-terminal dimerization domain permits homeodomain proteins to choose compatible partners and initiate sexual development in the mushroom Coprinus cinereus. Plant Cell 7:773–783 Bandoni RJ (1963) Conjugation in Tremella mesenterica. Can J Bot 41:467–474 Banno I (1967) Studies on sexuality of Rhodotorula. J Gen Appl Microbiol 13:167–196 Banuett F (1995) Genetics of Ustilago maydis, a fungal pathogen that induces tumors in maize. Ann Rev Genet 29:179–208 Banuett F (2007) History of mating types in Ustilago maydis. In: Heitman J, Kronstad JW, Taylor JW, Casselton LA (eds) Sex in fungi. Molecular determination and evolutionary implications. ASM Press, Washington, DC, pp 351–388 Banuett F, Herskowitz I (1989) Different a alleles of Ustilago maydis are necessary for maintenance of filamentous growth but not for meiosis. Exp Mycol 18:247–266 Barres B, Halkett F, Dutech C, Andrieux A, Pinon J, Frey P (2008) Genetic structure of the poplar rust fungus Melampsora larici-populina: Evidence for isolation by distance in Europe and recent founder effects overseas. Infect Genet Evol 8:577–587 ˚ stro¨m SU (2010) Ume6 is Barsoum E, Sjo¨strand JOO, A required for the MATa/MATa cellular identity and transcriptional silencing in Kluyveromyces lactis. Genetics 184:999–1011 Basse C, Farsing JW (2006) Promoters and their regulation in Ustilago maydis and other phytopathogenic fungi. FEMS Microbiol Lett 254:208–216

148

Ursula Ku¨es et al.

Bensaude M (1918) Recherches sur le cycle e´volutif et la sexualite´ chez les Basidiomyce`tes. PhD thesis, Faculte´ des Sciences de Paris, Imprimerie Nemourienne, Henri Bouloy, Nemours, France Billiard S, Lo´pez-Villavicencio M, Devier B, Hood ME, Fairhead C, Giraud T (2011) Having sex, yes, but with whom? Inferences from fungi on the evolution of anisogamy and mating types. Biol Rev 86:421–442 Blakeslea AF (1904) Zygospore formation a sexual process. Science 3:864–866 Bloomfield G, Skelton J, Ivens A, Tanakqa Y, Kay RR (2010) Sex determination in the social amoeba Dictyostelium discoideum. Science 330:1533–1536 Boidin J, Lanquentin P (1984) Le genre Amylostereum (Basidiomycetes) incompatibilite´s partielles entre espe`ces allopartriques. Bull Trimest Soc Mycol Fr 100:211–236 Bo¨lker M (1998) Sex and crime: Heterotrimeric G proteins in fungal mating and pathogenesis. Fungal Genet Biol 25:143–156 Bo¨lker M, Urban M, Kahmann R (1992) The a mating type locus of U. maydis specifies cell signalling components. Cell 68:441–450 Boulton MD, Kolmer JA, Garvin DF (2008) Wheat leaf rust caused by Puccinia triticina. Mol Plant Pathol 9:563–575 Brachmann A, Weinzierl G, Ka¨mper J, Kahmann R (2001) Identification of genes of the bW/bE regulatory cascade in Ustilago maydis. Mol Microbiol 42:1047–1063 Brefort T, Doehlemann G, Mendoza-Mendoza A, Reissmann S, Djamei A, Kahmann R (2009) Ustilago maydis as a pathogen. Ann Rev Phytopathol 47:423–445 Brown AJ, Casselton LA (2001) Mating in mushrooms: increasing the chances but prolonging the affair. Trends Genet 17:393–400 Bui T, Lin X, Malik R, Heitman J, Carter D (2008) Isolates of Cryptococcus neoformans from infected animals reveal genetic exchange in unisexual, a mating type populations. Eukaryot Cell 7:1771–1780 Buller AHR (1950) Researches on fungi. Vol. VII. The sexual process in the Uredinales. University of Toronto Press, Toronto, Canada Bu¨rglin T (1997) Analysis of TALE superclass homeobox genes (MEIS, PBC, KNOX, Iroquois, TGIF) reveals a novel domain conserved between plants and animals. Nuc Acids Res 25:4173–4180 Burt A (2000) Sex, recombination, and the efficiency of selection – was Weismann right? Evolution 54:337–351 Butler G, Kenny C, Fagan A, Kurischko C, Gaillardin C, Wolfe KH (2004) Evolution of the MAT locus and its HO endonuclease in yeast species. Proc Natl Acad Sci USA 101:1632–1637 Butlin R (2002) Evolution of sex: the costs and benefits of sex: new insights from old asexual lineages. Nat Rev Genet 3:311–317 Byrnes EJ III, Li W, Lewit Y, Perfect JR, Carter DA, Cox GM, Heitman J (2010) First reported case of Cryptococcus gattii in the Southeastern USA: Implications for travel-associated acquisition of an emerging pathogen. PLoS ONE 4:5851

Caldwell GA, Naider F, Becker JM (1995) Fungal lipopeptide mating pheromones: a model system for the study of protein prenylation. Microbiol Rev 59: 406–422 Casselton LA (2008) Fungal sex genes – searching for the ancestors. BioEssays 30:711–714 Casselton LA, Challen MP (2006) The mating types of the basidiomycetes. In: Ku¨es U, Fischer R (eds) The Mycota, vol. 1, 2nd edn. Growth, differentiation and sexuality. Springer, Berlin Heidelberg New York, pp 357–374 Casselton LA, Ku¨es U (1994) Mating-type genes in homobasidiomycetes. In: Wessels JGH, Meinhard F (eds) The Mycota, vol. 1. Growth, differentiation, and sexuality. Springer, Berlin Heidelberg New York, pp 307–321 Casselton LA, Ku¨es U (2007) The origin of multiple mating types in the model mushrooms Coprinopsis cinerea and Schizophyllum commune. In: Heitman J, Kronstad JW, Taylor JW, Casselton LA (eds) Sex in fungi. Molecular determination and evolutionary implications. ASM Press, Washington, DC, pp 283–300 Casselton LA, Olesnicky ND (1998) Molecular genetics of mating type recognition in basidiomycete fungi. Microbiol Mol Biol Rev 62:55–70 Chang YC, Wickes BL, Miller GF, Penoyer LA, KwonChung KJ (2000) Cryptococcus neoformans STE12a regulates virulence but is not essential for mating. J Exp Med 191:871–881 Chang YC, Penoyer LA, Kwon-Chung KJ (2001) The second STE12 homologue of Cryptococcus neoformans is MATa-specific and plays an important role in virulence. Proc Natl Acad Sci USA 98:3258–3263 Chang YC, Miller GF, Kwon-Chung KJ (2003) Importance of a developmentally regulated pheromone receptor of Cryptococcus neoformans for virulence. Infect Immun 71:4953–4960 Chang YC, Wright LC, Tscharke RL, Sorrell TC, Wilson CF, Kwon-Chung KJ (2004) Regulatory roles for the homeodomain and C2H2 zinc finger regions of Cryptococcus neoformans STE12ap. Mol Microbiol 53:1385–1396 Charlesworth B, Langley CH (1989) The population genetics of Drosophila transposable elements. Ann Rev Genet 23:251–287 Chaturvedi V, Fan J, Stein B, Behr MJ, Samsonoff WA, Wickes BL, Chaturvedi S (2002) Molecular genetic analyses of mating pheromones reveal intervariety mating of hybridization in Cryptococcus neoformans. Infect Immun 70:5225–5235 Chen P, Sapperstein SK, Choi JD, Michaelis S (1997) Biogenesis of the Saccharomyces cerevisiae mating pheromone a-factor. J Cell Biol 136:251–259 Chung S, Karos M, Chang YC, Lukszo J, Wickes BL, KwonChung KJ (2002) Molecular analysis of CPRa, a MATa-specific pheromone receptor gene of Cryptococcus neoformans. Eukaryot Cell 1:432–439 Clarke DL, Woodlee GL, McClelland CM, Seymour TS, Wickes BL (2001) The Cryptococcus neoformans STE11a gene is similar to other fungal mitogen-activated protein kinase kinase (MAPKKK) genes but is mating type specific. Mol Microbiol 40:200–213

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality ´ , Gonc¸alves P Coelho MA, Rosa A, Rodriguez N, Fonseca A (2008) Identification of mating type genes in the bipolar basidiomycetous yeast Rhodosporidium toruloides: First insight into the MAT locus structure of the genus Sporidiobolales. Eukaryot Cell 7:1053–1061 Coelho MA, Sampaio JP, Gonc¸alves P (2010) A deviation from the bipolar-tetrapolar mating paradigm in an early diverged basidiomycete. PLoS Genet 6:e1001052 Coppin E, Debuchy R, Arnaise S, Picard M (1997) Mating types and sexual development in filamentous ascomycetes. Microbiol Mol Biol Rev 61:411–428 Craigie JH (1927) Discovery of the function of the pycnia of the rust fungi. Nature 120:765–67 Craigie JH (1931) An experimental investigation of sex in the rust fungi. Phytopathol 21:101–1040 Cubeta MA, Vilgalys R (1997) Population biology of the Rhizoctonia solani complex. Phytopathol 7:480–484 Davey J, Davis K, Hughes M, Ladds G, Powner D (1998) The processing of yeast pheromones. Sem Cell Dev Biol 9:19–30 Davidson RC, Moore TDE, Odom AR, Heitman J (2000) Characterization of the MFa pheromone of the human fungal pathogen Cryptococcus neoformans. Mol Microbiol 38:1017–1026 Davidson RC, Nichols CBN, Cox GM, Perfect JR, Heitman J (2003) A MAP kinase cascade composed of cell type specific and non-specific elements controls mating and differentiation of the fungal pathogen Cryptococcus neoformans. Mol Microbiol 49:469–485 Daud F, Napiah I (1991) Mating systems of two Auricularia spp. in Malyasia. Malyasian Appl Biol 20: 87–91 Day AW (1979) Mating type and morphogenesis in Ustilago violacea. Bot Gaz 140:94–109 Day PR (1960) The structure of the A mating type locus in Coprinus lagopus. Genetics 45:641–650 Derelle R, Lopez P, Le Guyaser H, Manuel M (2007) Homeodomain proteins belong to the ancestral toolkit of eukaryotes. Evol Dev 9:212–219 Devier B, Aguileta G, Hood M, Giraud T (2009) Ancient trans-specific polymorphism at pheromone receptor genes in basidiomycetes. Genetics 181:209–223 Devier, B, Aguileta G, Hood ME, Giraud T (2010) Using phylogenies of pheromone receptor genes in Microbotryum violaceum species complex to investigate possible speciation by hybridization. Mycologia 102: 689–696 Dhawale SS, Kessler K (1993) Alternative methods for production and staining of Phanerochaete chrysosporium basidiospores. Appl Environ Microbiol 59: 1675–1677 Dhitaphichit P, Nawawongwiwat J (2006) Mating system of five edible species of the mushroom, genus Lentinus. Songklanakarin J Sci Technol 28:285–292 Dolgin ES, Charlesworth B (2006) The fate of transposable elements in asexual populations. Genetics 174:817–827 Duboule D (1994) Guidebook to homeobox genes. Oxford University Press, Oxford, UK Dunthorn M, Katz LA (2010) Secretive ciliates and putative asexuality in microbial eukaryotes. Trends Microbiol 18:183–188

149

Duplessis S, Cuomo CA, Lin Y-P, Aerts A, Tisserant E, Veneault-Fourrey C, Joly DL, Hacquard S, Amselem J, Cantarel BL, Chiu R, Coutinho PM, Feau N, Field M, Frey P, Gelhaye E, Goldberg J, Grabherr MG, Kodira CD, Kohler A, Ku¨es U, Lindquist EA, Lucas SM, Maho R, Mauceli E, Morin E, Murat C, Pangilinan JL, Park R, Pearson M, Quesneville H, Rouhier N, Sakthikumar S, Salamov AA, Schmutz J, Selles B, Shapiro H, Tanguay P, Tuskan GA, Henrissat B, Van de Peer Y, Rouze´ P, Ellis JG, Dodds PN, Schein JE, Zhong S, Hamelin RC, Grigoriev IV, Szabo LJ, Martin F (2011) Obligate biotrophy features inraveled by the genomic analysis of rust fungi. Proc Natl Acad Sci USA, doi:10.1073/pnas.1019315108 Dyer PS (2008) Evolutionary biology: Genomic clues to original sex in fungi. Curr Biol 18:R207–R209 Ekena JL, Stanton BC, Schiebe-Owens JA, Hull CM (2008) Sexual development in Cryptococcus neoformans requires CLP1, a target of the homeodomain transcription factor Sxi1a and Sxi2a. Eukaryot Cell 7:48–57 Esser K (2000) Kryptogamen 1. Cyanobakterien Algen Pilze Flechten, 3rd edn. Springer, Berlin Heidelberg New York Esser K, Meinhardt F (1977) Common genetic control of dikaryotic and monokaryotic fruiting in the basidiomycete Agrocybe aegerita. Mol Gen Genet 155:113–115 Esser K, Saleh F, Meinhardt F (1979) Genetics of fruit body production in higher basidiomycetes. 2. Monokaryotic and dikaryotic fruiting in Schizophyllum commune. Curr Genet 1:85–88 Fedler M, Luh KS, Stelter K, Nieto-Jacobo F, Basse CW (2009) The a2 mating-type locus genes lga2 and rga2 direct uniparental mitochondrial DNA (mtDNA) inheritance and constrain mtDNA recombination during sexual development of Ustilago maydis. Genetics 181:847–860 Feldbru¨gge M, Ka¨mper J, Steinberg G, Kahmann R (2004) Regulation of mating and pathogenic development in Ustilago maydis. Curr Opin Microbiol 7:666–672 Feldbru¨gge M, Bo¨lker M, Steinberg G, Ka¨mper J, Kahmann R (2006) Regulatory and structural networks orchestrating mating, dimorphism, cell shape, and pathogenesis in Ustilago maydis. In: Ku¨es U, Fischer R (eds) The mycota. Vol. 1, 2nd edn. Growth, differentiation and sexuality. Springer, Berlin Heidelberg New York, pp 375–392 Fell JW, Statzell Tallman A (1981) Heterothallism in the basidiomycetous yeast genus Sporidiobolus Nyland. Curr Microbiol 5:77–82 Fell JW, Statzell-Tallman A (1998) Rhodosporidium Banno. In: Kurtzman CP and Fell JW (eds) The yeasts, a taxonomic study, 4th edn. Elsevier, Amsterdam, pp 678–692 Findley K, Rodriguez-Carres M, Metin B, Kroiss J, Fonseca ´ , Vilgalys R, Heitman J (2009) Phylogeny and pheA notypic characterization of pathogenic Cryptococcus species and closely related saprobic taxa in the Tremellales. Eukaryot Cell 8:363–361 Fowler TJ, Vaillancourt LJ (2007) Pheromones and pheromone receptors in Schizophyllum commune mate recognition: Retrospective of a half-century of progress and a look ahead. In: Heitman J, Kronstad JW, Taylor

150

Ursula Ku¨es et al.

JW, Casselton LA (eds) Sex in fungi. Molecular determination and evolutionary implications. ASM Press, Washington, DC, pp 301–316 Fowler TJ, DeSimone SM, Mitton MF, Kurjan J, Raper CA (1999) Multiple sex pheromones and receptors of a mushroom-producing fungus elicit mating in yeast. Mol Biol Cell 10:2559–2572 Fowler TJ, Mitton MF, Vaillancourt LJ, Raper CA (2001) Changes in mate recognition through alterations of pheromones and receptors in the multisexual mushroom fungus Schizophyllum commune. Genetics 158:1491–1503 Fowler TJ, Mitton MF, Rees EI, Raper CA (2004) Crossing the boundary between the Ba and Bb mating-type loci in Schizophyllum commune. Fungal Genet Biol 41:89–101 Fox HM, Burden J, Chang S-T, Pederby JF (1994) Matingtype incompatibility between commercial strains of Lentinula edodes. Exp Mycol 18:95–102 Frankel C, Ellingboe AH (1977) New mutations and a 7chromosome map of Schizophyllum commune. Genetics 85:417–425 Franzot SP, Hamdan JS, Currie BP, Casadevall A (1997) Molecular epidemiology of Cryptococcus neoformans in Brazil and the United States: evidence for both local genetic differences and a global clonal population structure. J Clin Microbiol 35:2243–2251 Fraser JA, Heitman J (2005) Chromosomal sex-determining regions in animals, plants and fungi. Curr Opin Genet Dev 15:645–651 Fraser JA, Subaran RL, Nichols CB, Heitman LJ (2003) Recapitulation of the sexual cycle of the primary fungal pathogen Cryptococcus neoformans var. gattii: Implications for an outbreak on Vancouver Island, Canada. Eukaryot Cell 2:1036–1045 Fraser JA, Diezmann S, Subaran RL, Allen A, Lengeler KB, Dietrich FS, Heitman J (2004) Convergent evolution of chromosomal sex-determining regions in the animal and fungal kingdoms. PLoS Biol 2:2243–2255 Fraser JA, Giles SS, Wenink EC, Geunes-Boyer SG, Wright JR, Diezmann S, Allen A, Stajich JE, Dietrich FS, Perfect JR, Heitman J (2005) Same-sex mating and the origin of the Vancouver Island Cryptococcus gattii outbreak. Nature 437:1360–1364 Fraser JA, Hsueh Y-P, Fundley KM, Heitman J (2007) Evolution of the mating-type locus: the basidiomycetes. In: Heitman J, Kronstad JW, Taylor JW, Casselton LA (eds) Sex in fungi. Molecular determination and evolutionary implications. ASM Press, Washington, DC, pp 19–34 Froeliger EH, Leong SA (1991) The a mating-type alleles of Ustilago maydis are idiomorphs. Gene 100:113–122 Fuchs U, Hause G, Schuchardt I, Steinberg G (2006) Endocytosis is essential for pathogenic development in the corn smut fungus Ustilago maydis. Plant Cell 18:2066–2081 Fuller RS, Sterne RE, Thorner J (1988) Enzymes required for yeast prohormone processing. Ann Rev Physiol 50:345–362

Galagan JE, Calvo SE, Borkovich KA, Selker EU, Read ND, Jaffe D, FitzHugh W, Ma LJ, Smirnov S, Purcell S, Rehman B, Elkins T, Engels R, Wang S, Nielsen CB, Butler J, Endrizzi M, Qui D, Ianakiev P, Bell-Pedersen D, Nelson MA, Werner-Washburne M, Selitrennikoff CP, Kinsey JA, Braun EL, Zelter A, Schulte U, Kothe GO, Jedd G, Mewes W, Staben C, Marcotte E, Greenberg D, Roy A, Foley K, Naylor J, Stange-Thomann N, Barrett R, Gnerre S, Kamal M, Kamvysselis M, Mauceli E, Bielke C, Rudd S, Frishman D, Krystofova S, Rasmussen C, Metzenberg RL, Perkins DD, Kroken S, Cogoni C, Macino G, Catcheside D, Li W, Pratt RJ, Osmani SA, DeSouza CP, Glass L, Orbach MJ, Berglund JA, Voelker R, Yarden O, Plamann M, Seiler S, Dunlap J, Radford A, Aramayo R, Natvig DO, Alex LA, Mannhaupt G, Ebbole DJ, Freitag M, Paulsen I, Sachs MS, Lander ES, Nusbaum C, Birren B (2003) The genome sequence of the filamentous fungus Neurospora crassa. Nature 422:859–868 Garcia-Muse T, Steinberg G, Perez-Martin J (2003) Pheromone-induced G2 arrest on the phytopathogenic fungus Ustilago maydis. Eukaryot Cell 2:494–500 Gerard PR, Husson C, Pinon J, Frey P (2006) Comparison of genetic and virulence diversity of Melampsora larici-populina populations on wild and cultivated poplar and influence of the alternate host. Phytopathology 96:1027–1036 Gieser PT, May G (1994) Comparison of 2 b1 alleles from within the A-mating-type of the basidiomycete Coprinus cinereus. Gene 146:167–176 Gillissen B, Bergemann J, Sandmann C, Schro¨er B, Bo¨lker M, Kahmann R (1992) A 2-component regulatory system for self non-self recognition in Ustilago maydis. Cell 68:647–657 Giraud T, Yockteng R, Lo´pez-Villavicencio M, Refre´gier G, Hood ME (2008) Mating system of the anther smut fungus Microbotryum violaceum: Selfing under heterothallism. Eukaryot Cell 7:765–775 Gola S, Kothe E (2003a) The little difference: in vivo analysis of pheromone discrimination in Schizophyllum commune. Curr Genet 42:276–283 Gola S, Kothe E (2003b) An expression system for the functional analysis of pheromone genes in the tetrapolar basidiomycete Schizophyllum commune. J Basic Microbiol 43:104–112 Gola S, Hegner J, Kothe E (2000) Chimeric pheromone receptors in the basidiomycete Schizophyllum commune. Fungal Genet Biol 30:191–196 Gold MH, Alic M (1993) Molecular biology of the lignindegrading basidiomycete Phanerochaete chrysosporium. Microbiol Rev 57:605–622 Golubev W (1993) Rhodosporidium babjevae, a new heterothallic yeast species (Ustilaginales). Syst Appl Microbiol 16:445–449 Graul RC, Sade´e W (2001) Evolutionary relationships among G protein-coupled receptors using a clustered database approach. AAPS PharmSci 3:12 Griffiths GW, Hedger JN (1994) The breeding biology of biotypes of the witches´ broom pathogen of cocoa, Crinipellis perniciosa. Heredity 72:278–289

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality Groth JV (1975) Two additive, independent genes for mycelial growth versus sporidial growth of haploid cultures of Ustilago hordei. Can J Bot 53:2233–2239 Gryganski AP, Lee SC, Litvintseva AP, Smith ME, Bonito G, Porter TM, Anishchenko IM, Heitman J, Vilgalys R (2010) Structure, function, and phylogeny of the mating locus in the Rhizopus oryzae complex. PLoS ONE 5:e12273 Haber JE (2007) Decisions, decisions: Donor preference during budding yeast mating-type switching. In: Heitman J, Kronstad JW, Taylor JW, Casselton LA (eds) Sex in fungi. Molecular determination and evolutionary implications. ASM Press, Washington, DC, pp 159–170 Halsall JR, Milner MJ, Casselton LA (2000) Three subfamilies of pheromone and receptor genes generate multiple B mating specificities in the mushroom Coprinus cinereus. Genetics 154:1115–1123 Hartmann HA, Kahmann R, Bo¨lker M (1996) The pheromone response factor coordinates filamentous growth and pathogenicity in Ustilago maydis. EMBO J 15:1632–1642 Hegner J, Siebert–Bartholmei C, Kothe E (1999) Ligand recognition on multiallelic pheromone receptors from the basidiomycete Schizophyllum commune studied in yeast. Fungal Genet Biol 26:190–197 Heimel K, Scherer M, Schuler D, Ka¨mper J (2010a) The Ustilago maydis Clp1 protein orchestrates pheromone and b-dependent signalling pathways to coordinate the cell cycle and pathogenic development. Plant Cell 22:2908–2922 Heimel K, Scherer M, Vranes M, Wahl R, Pothiratana C, Schuler B, Voncon V, Finkernagel F, Flor-Parra I, Ka¨mper J (2010) The transcription factor Rbf1 is the master regulator for b-mating type controlled pathogenic development in Ustilago maydis. PloS Pathog 6: e1001035 Heitman J (2009) Microbial genetics: Love the one you’re with. Nature 460:807–808 Heng HHQ (2007) Elimination of altered karyotypes by sexual reproduction preserves species identity. Genome 50:517–524 Herskowitz I (1988) Life-cycle of the budding yeast Saccharomyces cerevisiae. Microbiol Rev 52:536–553 Herskowitz I (1989) A regulatory hierarchy for cell specialization in yeast. Nature 342:749–757 Hibbett DS (2006) A phylogenetic overview of the Agaricomycotina. Mycologia 98:917–925 Hibbett DS, Binder M, Bischoff JF, Blackwell M, Cannon PF, Eriksson OE, Huhndorf S, James T, Kirk PM, Lu¨cking R, Lumbusch HT, Lutzoni F, Matheny PB, McLaughlin DJ, Powell MJ, Redhead S, Schoch CL, Spatafora JW, Stalpers JA, Vilgalys R, Aime MC, Aptroot A, Bauer R, Begerow D, Benny GL, Castlebury LA, Crous PW, Dai YC, Gams W, Geiser DM, Griffith GW, Gueidan C, Hawksworth DL, Hestmark G, Hosaka K, Humber RA, Hyde KD, Ironside JE, Koljalg U, Kurtzman CP, Larsson KH, Lichtwardt R, Longcore J, Miadlikowska J, Miller A, Moncalvo M, Mozley-Standridge S, Oberwinkler F, Parmasto E, Reeb V, Rogers JD, Rous C, Ryvarden L, Sampaio JP, Schu¨ssler A, Sugiyama J, Thorn RG, Tibell L, Untereiner WA, Walker C, Wang

151

Z, Weir A, Weiss M, White MM, Winka K, Yao YJ, Zhang N (2007) A high-level phylogenetic classification of the fungi. Mycol Res 111:509–547 Hiremath SS, Chowdhary A, Kowshik T, Randhawa HS, Sun S, Xu J (2008) Long-distance dispersal and recombination in environmental populations of Cryptococcus neoformans var. grubii from India. Microbiol 154:1513–1524 Hiscock SJ, Ku¨es U (1999) Cellular and molecular mechanisms of sexual incompatibility in plants and fungi. Int Rev Cytol 193:165–295 Hoffmann JA, Kendrick EL (1969) Genetic control of compatibility in Tilletia controversa. Phytopathol 59:79–83 Holton CS, Kendrick EL (1957) Fusion between secondary sporidia in culture as a valid index of sex compatibility in Tilletia caries. Phytopathol 47:688–89 Hood ME (2002) Dimorphic mating-type chromosomes in the fungus Microbotryum violaceum. Genetics 160:457–461 Hood ME (2005) Repetitive DNA in the automictic fungus Microbotryum violaceum. Genetica 124:1–10 Hood ME, Antonovics J (2004) Mating within the meiotic tetrad and the maintenance of genomic heterozygosity. Genetics 166:1751–1759 Hood ME, Antonovics J, Koskella B (2004) Shared forces of sex chromosome evolution in haploids and diploids. Genetics 168:141–146 Hsueh Y-P, Heitman J (2008) Orchestration of sexual reproduction and virulence by the fungal matingtype locus. Curr Opin Microbiol 11:617–524 Hsueh Y-P, Shen W-C (2005) A homolog of Ste6, the afactor transporter in Saccharomyces cerevisiae, is required for mating but not for monokaryotic fruiting in Cryptococcus neoformans. Eukaryot Cell 4:147–255 Hsueh Y-P, Idnurm A, Heitman J (2006) Recombination hotspots flank the Cryptococcus mating-type locus: Implications for the evolution of a fungal sex-chromosome. PLoS Genet 2:e184 Hsueh Y-P, Xue C, Heitman J (2007) G protein signaling governing cell fate decisions involves opposing Ga subunits in Cryptococcus neoformans. Mol Biol Cell 18:3237–3249 Hsueh Y-P, Fraser JA, Heitman J (2008) Transitions in sexuality: Recapitulation of an ancestral tri- and tetrapolar mating system in Cryptococcus neoformans. Eukaryot Cell 7:1847–1855 Hsueh Y-P, Xue C, Heitman J (2009) A constitutively active GPCR governs morphogenic transitions in Cryptococcus neoformans. EMBO J 28:1220–1233 Hsueh Y-P, Metin B, Findley K, Rodriguez-Carres, Heitman J (2011) The mating-type locus of Cryptococcus: evolution of gene clusters governing sex determination and sexual reproduction from phylogenomic perspective. In: Heitman J, Kozel TR, Kwon-Chung KJ, Perfect JR, Casadavall A (eds) Cryptococcus: From human pathogen to model yeast. ASM Press, Washington, DC, pp 139–149 Hull CM, Heitman J (2002) Genetics of Cryptococcus neoformans. Ann Rev Genet 36:557–615 Hull CM, Davidson RC, Heitman J (2002) Cell identity and sexual development in Cryptococcus neoformans are

152

Ursula Ku¨es et al.

controlled by the mating-type-specific homeodomain protein Sxi1a. Genes Dev 16:3046–3060 Hull CM, Cox GM, Heitman J (2004) The a-specific cell identity factor Sxi1a is not required for virulence of Cryptococcus neoformans. Infect Immun 72: 3643–3645 Hull CM, Boily MJ, Heitman J (2005) Sex-specific homeodomain proteins Sxi1a and Sxi2a coordinately regulate sexual development in Cryptococcus neoformans. Eukaryot Cell 4:526–535 Hurst LD (1996) Why are there only two sexes? Proc R Soc Lon Ser B Biol Sci 263:415–422 Huyer G, Kistler A, Nouvel FJ, George CM, Boyte ML, Michaelis S (2006) Saccharomyces cerevisiae a-factor mutants reveals residues criticical for processing, activity, and export. Eukaryot Cell. 5:1560–1570 Idnurm A, Heitman J (2005) Light controls growth and development via a conserved pathway in the fungal kingdom. PLoS Biol 3:e95 Idnurm A Bahn YS, Nielsen K, Lin X, Fraser JA, Heitman J (2005) Deciphering the model pathogenic fungus Cryptococcus neoformans. Nature Rev Microbiol 3:753–764 Idnurm A, James TY, Vilgalys R (2007) Sex in the rest: Mysterious mating in the Chytridiomycota and Zygomycota. In: Heitman J, Kronstad JW, Taylor JW, Casselton LA (eds) Sex in fungi. Molecular determination and evolutionary implications. ASM Press, Washington, DC, pp 407–418 Idnurm A, Walton FJ, Floyd A, Heitman J (2008) Identification of the sex genes in an early diverged fungus. Nature 451:193–196 Imai Y, Yamamoto M (1994) The fission yeast mating pheromone P-factor: its molecular structure, gene structure and physiological activities to induce gene expression and G1 arrest in the mating partner. Genes Dev 8:328–338 Isaya G, Sakati WR, Rollins RA, Shen GP, Hanson LC, Ullrich RC, Novotny CP (1995) Mammalian mitochondrial intermediate peptidase: Structure/function analysis of a new homolog from Schizophyllum commune and relationship to thimet oligopeptidases. Genomics 28:450–461 Ishibashi Y, Sakagami Y, Isogai A, Suziki A (1984) Structures of tremerogens A-9291-I and A-9291VIII. Peptidyl sex hormones of Tremella brasiliensis. Biochemistry 23:1399–1404 James TY, Ku¨es U, Rehner SA, Vilgalys R (2004a) Evolution of the gene encoding mitochondrial intermediate peptidase and its cosegregation with the A matingtype locus of mushroom fungi. Fungal Genet Biol 41:381–390 James TY, Liou SR, Vilgalys R (2004b) The genetic structure and diversity of the A and B mating-type genes from the tropical oyster mushroom, Pleurotus djamor. Fungal Genet Biol 41:813–825 James TY, Srivilai P, Ku¨es U, Vilgalys R (2006) Evolution of the bipolar mating system of the mushroom Coprinellus disseminatus from its tetrapolar ancestors involves loss of mating-type-specific pheromone receptor function. Genetics 172:1877–1891

James TY, Lee M, van Diepen LTA (2011) A single matingtype locus comprised of homeodomain genes promotes nuclear migration and heterokaryosis in the white-rot fungus Phanerochaete chrysosporium. Eukaryot Cell 10:249–261 Josefsson L-G (1999) Evidence for kinship between diverse G-protein coupled receptors. Gene 239:333–340 Judson OP, Nordmark BB (1996) Ancient asexual scandals. Trends Ecol Evol 1:11:41–46 Julian MC, Dullemans AM, van Silfthout CH, Keijer J (1997) Nuclear behaviour in homokaryotic and heterokaryotic fruiting of Thanatephorus cucumeris (Rhizoctonia solani) anastomosis group 1, subgroup IC. Mycologia 89:361–374 Kaffarnik F, Mu¨ller P, Leibundgut M, Kahmann R, Feldbru¨gge M (2003) PKA and MAPK phosporylation of Prf1 allows promoter discrimination in Ustilago maydis. EMBO J 22:5817–5826 Kahmann R, Ka¨mper J (2004) Ustilago maydis: how its biology relates to pathogenic development. New Phytol 164:31–42 Kahmann R, Schirawski J (2007) Mating in the smut fungi: From a to b to the downstream cascades. In: Heitman J, Kronstad JW, Taylor JW, Casselton LA (eds) Sex in fungi. Molecular determination and evolutionary implications. ASM Press, Washington, DC, pp 377–387 Kamiya Y, Sakarai A, Tamura S, Abe K, Tsuchiya E, Fukui S (1977) Isolation and chemical characterization of peptidyl factor controlling mating tube formation in Rhodosporidium toruloides. Agric Biol Chem 41:1099–1100 Kamiya Y, Sakurai A, Tamura S, Takahashi N, Abe K, Tsuchiya E, Fukui S (1978a) Amino-acid sequence of rhodotorucine-A, a peptidyl factor controlling mating tube formation in Rhodosporidium toruloides. Agric Biol Chem 42:209–211 Kamiya Y, Sakurai A, Tamura S, Takahashi N, Abe K, Tsuchiya E, Fukui S (1978b) Isolation of rhodotorucine A, a peptidyl factor inducing mating tube formation in Rhodosporidium toruloides. Agric Biol Chem 42:1239–1243 Kamiya Y, Sakurai A, Tamura S, Takahashi N, Abe K, Tsuchiya E, Fukui S, Kitada C, Fujino (1978c) Structure of rhodotorucine-A, a novel lipopeptide inducing mating tube formation in Rhodosporidium. Biochem Biophys Res Com 83:1077–1083 Kamiya Y, Sakurai A, Tamura S, Tsuchiya E, Abe K, Fujui S (1979) Structure of rhodotorucine-A, a peptidyl factor, inducing mating tube formation in Rhodosporidium toruloides. Agric Biol Chem 43:363–369 Ka¨mper J, Reichmann M, Romeis T, Bo¨lker M, Kahmann R (1995) Multiallelic recognition – Nonself-dependent dimerization of the bE and bW homeodomain proteins in Ustilago maydis. Cell 81:73–83 Ka¨mper J, Kahmann R, Bo¨lker M, Ma LJ, Brefort T, Saville BJ, Banuett F, Kronstad JW, Gold SE, Mu¨ller O, Perlin MH, Wo¨sten HAB, de Vries R, Ruiz-Herrera J, Reynaga-Pen˜a CG, Snetselaar K, McCann M, Pe´rez-Martı´n J, Feldbru¨gge M, Basse CW, Steinberg G, Ibeas JI, Holloman W, Guzman P, Farman M, Stajich JE, Sentandreu R, Gonza´lez-Prieto JM, Kennell JC, Molina L,

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality Schirawski J, Mendoza-Mendoza A, Greilinger D, Mu¨nch K, Ro¨ssel N, Scherer M, Vranesˇ M, Ladendorf O, Voncon V, Fuchs U, Sandrock B, Meng S, Ho EC, Cahill MJ, Boyce KJ, Klose J, Klosterman SJ, Deelstra HJ, Ortiz-Castellanos L, Li W, Sanchez-Alsonso P, Schreier PH, Hauser-Hahn I, Vaupel M, Koopmann E, Friedrich G, Voss H, Schlu¨ter T, Margolis J, Platt D, Schwimmer C, Gnirke A, Chen F, Vysotskaia V, Mannhaupt G, Gu¨ldener U, Mu¨nsterko¨tter M, Haase D, Oesterheld M, Mewes HW, Mauceli EW, DeCaprio D, Wade CM, Butler J, Young S, Jaffe DB, Calvo S, Nusbaum C, Galagan J, Birren BW (2006) Insights from the genome of the biotrophic fungal pathogen Ustilago maydis. Nature 444:97–101 Karos M, Chang YC, McClelland CM, Clarke DL, Fu J, Wickes BL, Kwon-Chung KJ (2000) Mapping of the Cryptococcus neoformans MATa locus: Presence of mating type-specific mitogen-activated protein cascade homologs. J Bacteriol 182:6222–6227 Kemp ROF (1974) Bifactorial incompatibility in the twospored basidiomycetes Coprinus sassii and Coprinus bilanatus. Trans Br Mycol Soc 62:547–555 Kidd SE, Guo H, Bartlett KH, Xu J, Kronstad JW (2005) Comparative gene genealogies indicate that two clonal lineages of Cryptococcus gattii in British Columbia resemble strains from other geographical areas. Eukaryot Cell 4:1629–1638 Kirk PM, Cannon PF, Stalpers JA (2008) Dictionary of the fungi, 10th edn. CABI, Wallingford, UK Kniep H (1928) Die Sexualita¨t der niederen Pflanzen. Fischer, Jena, Germany Kollmorgen JF, Trione EJ (1980) Mating-type interactions between sporidia of the wheat-bunt fungus Tilletia caries. Can J Bot 58:1994–2000 Koltin Y, Raper JR, Simchen G (1967) The genetic structure of the incompatibility factors of Schizophyllum commune. The B factor. Proc Natl Acad Sci USA 57:55–62 Koltin Y, Stamberg J, Lemke PA (1972) Genetic structure and evolution of the incompatibility factors in higher fungi. Bacteriol Rev 36:156–171 Koppitz M, Spellig T, Kahmann R, Kessler H (1996) Lipoconjugates: Structure–activity studies for pheromone analogues of Ustilago maydis with varied lipophilicity. Int J Peptide Protein Res 48:377–390 Kosted PJ, Gerhardt SA, Anderson CM, Stierle A, Sherwood JE (2000) Structural requirements for activity of the pheromones of Ustilago hordei. Fungal Genet Biol 29:107–117 Kothe E (1996) Tetrapolar fungal mating types: Sexes by the thousands. FEMS Microbiol Rev 18:65–87 Kothe E (1999) Mating types and pheromone recognition in the basidiomycete Schizophyllum commune. Fungal Genet Biol 27:146–152 Kothe E (2001) Mating-type genes for basidiomycete strain improvement in mushroom farming. Appl Microbiol Biotechnol 56:602–612 Kothe E, Raudaskoski M (2010) Basidiomycete mating type genes and pheromone signalling. Eukaryot Cell 9:847–859 Kothe E, Gola S, Wendland J (2003) Evolution of multispecific mating-type alleles for pheromone percep-

153

tion in the homobasidiomycete fungi. Curr Genet 42:268–275 Krejcˆ´ı R, Homolka L (1991) Genetic mapping in the lignindegrading basidiomycete Phanerochaete chrysosporium. Appl Env Microbiol 57:151–156 Kronstad JW, Leong SA (1990) The b mating type locus of Ustilago maydis contains variable and constant regions. Genes Dev 4:1384–1395 Kronstad JW, Staben C (1997) Mating type in filamentous fungi. Ann Rev Microbiol 31:245–276 Kruzel EK, Hull CM (2010) Establishing an unusual cell type: how to make a dikaryon. Curr Opin Microbiol 13:706–711 Ku¨es U (2000) Life history and developmental processes in the basidiomycete Coprinus cinereus. Microbiol Mol Biol Rev 64:316–353 Ku¨es U, Casselton LA (1992) Homeodomains and regulation of sexual development in basidiomycetes. Trends Genet 8:154–155 Ku¨es U, Casselton LA (1993) The origin of multiple mating types in mushrooms. J Cell Sci 104:227–230 Ku¨es U, Navarro-Gonza´lez M (2009) Communication of fungi on individual, species, kingdom, and above kingdom level. In: Anke T, Weber D (eds) The Mycota, Vol 15. Physiology and genetics, selected basic and applied aspects. Springer, Berlin, pp 79–106 Ku¨es U, Navarro-Gonza´lez M (2010) Mating-type orthologous genes in the primarily homothallic Moniliophthora perniciosa, the causal agent of Witches’ Broom Disease in cacao. J Basic Microbiol 50:442–451 Ku¨es U, Richardson WVJ, Tymon AM, Mutasa ES, Go¨ttgens B, Gaubatz S, Gregoriades A, Casselton LA (1992) The combination of dissimilar alleles of the Aa and Ab gene complexes, whose proteins contain homeo domain motifs, determines sexual development in the mushroom Coprinus cinereus. Genes Dev 6:568–577 Ku¨es U, Asante-Owusu RN, Mutasa ES, Tymon AM, Pardo EH, O´Shea SF, Go¨ttgens B, Casselton LA (1994a) Two classes of homeodomain proteins specify the multiple A mating types of the mushroom Coprinus cinereus. Plant Cell 6:1467–1475 Ku¨es U, Go¨ttgens B, Stratmann R, Richardson WSJ, ´ Shea SF, Casselton LA (1994b) A chimeric homeO odomain protein causes self compatibility and constitutive sexual development in the mushroom Coprinus cinereus. EMBO J 13:4054–4059 Ku¨es U, Tymon AM, Richardson WVJ, May G, Gieser PT, Casselton LA (1994c) A mating type factors of Coprinus cinereus have variable numbers of specificity genes encoding 2 classes of homeodomains. Mol Gen Genet 245:45–52 Ku¨es U, Granado JD, Hermann R, Boulianne RP, KerteszChaloupkova´ K, Aebi M (1998) The A mating type and blue light regulate all known differentiation processes in the basidiomycete Coprinus cinereus. Mol Gen Genet 260:81–91 Ku¨es U, James TY, Vilgalys R, Challen MP (2001a) The chromosomal region containing pab-1, mip, and the A mating type locus of the secondary homothallic homobasidiomycete Coprinus bilanatus. Curr Genet 39:16–24

154

Ursula Ku¨es et al.

Ku¨es U, Klaus MJ, Polak E, Aebi M (2001b) Multiple cotransformations in Coprinus cinereus. Fungal Genet Newslett 48:32–34 Ku¨es U, Polak E, Bottoli APF, Hollenstein M, Walser PJ, Boulianne RP, Hermann R, Aebi M (2002a) Vegetative development in Coprinus cinereus. In: Osiewacz HD (ed) Molecular biology of fungal development. Marcel Dekker, New York, pp 133–164 Ku¨es U, Walser PJ, Klaus MJ, Aebi M (2002b) Influence of activated A and B mating-type pathways on developmental processes in the basidiomycete Coprinus cinereus. Mol Gen Genom 268:262–271 Ku¨es U, Ku¨nzler M, Bottoli APF, Walser PJ, Granado JD, Liu Y, Bertossa RC, Ciardo D, Clergeot P-H, Loos S, Ruprich-Robert G, Aebi M (2004) Mushroom development in higher basidiomycetes. Implications for human and animal health. In: Kushwaha RKS (ed) Fungi in human and animal health. Scientific Publishers (India), Jodhpur, India, pp 431–470 Kurjan J, Herskowitz I (1982) Structure of a yeast pheromone gene (MFa): a putative a-factor precursor contains four tandem copies of mature a-factor. Cell 30:933–943 Kwon-Chung KJ (1975) A new genus, Filobasidiella, the perfect state of Cryptococcus neoformans. Mycologia 67:1197–1200 Kwon-Chung KJ (1976a) Morphogenesis of Filobasidiella neoformans, the sexual state of Cryptococcus neoformans. Mycologia 67:821–833 Kwon-Chung KJ (1976b) Morphogenesis of Filobasidiella, the sexual state of Cryptococcus neoformans B and C serotypes. Mycologia 68:821–833 Kwon-Chung KJ, Bennett JE (1978) Distribution of a and a mating types of Cryptococcus neoformans among natural and clinical isolates. Am J Epidemiol 108:337–340 Labare`re J, Noe¨l T (1992) Mating type switching in the tetrapolar basidiomycete Agrocybe aegerita. Genetics 131:307–319 Lahn BT, Page DC (1999) Four evolutionary strata on the human X chromosome. Science 286:964–967 Lawrence GJ (1980) Multiple mating-type specificities in the flax rust Melampsora lini. Science 209:501–503 Lee H, Chang YC, Kwon-Chung KJ (2005) TUP1 disruption reveals biological differences between MATa and MATa strains of Cryptococcus neoformans. Mol Microbiol 55:1222–1232 Lee H, Chang YC, Varma A, Kwon-Chung KJ (2009) Regulatory diversity of TUP1 in Cryptococcus neoformans. Eukaryot Cell 8:1901–1908 Lee N, Bakkeren G, Wong K, Sherwood JE, Kronstad JW (1999) The mating-type and pathogenicity locus of the fungus Ustilago hordei spans a 500-kb region. Proc Natl Acad Sci USA 96:15026–15031 Lee SC, Corradi N, Doan S, Dietrich F, Keeling PJ, Heitman J (2010a) Evolution of the sex-related locus and genomic features hsraed in microsporidia and fungi. PLoS One 5:e10539 Lee SC, Ni M, Li W, Shertz C, Heitman J (2010b) The evolution of sex: a perspective from the fungal kingdom. Microbiol Mol Biol Rev 74:298–340

Lemke PA (1969) A reevaluation of homothallism, heterothallism, and the species concept in Sistotrema brinkmannii. Mycologia 61:57–76 Lengeler KB, Davidson RC, D’Souza C, Harashima T, Shen WC, Wang P, Pan XW, Waugh M, Heitman J (2000a) Signal transduction cascades regulating fungal development and virulence. Microbiol Mol Biol Rev 64:746–785 Lengeler KB, Wang P, Cox GM, Perfect JR, Heitman J (2000b) Identification of the MATa mating-type locus of Cryptococcus neoformans reveals a serotype A MATa strain thought to have been extinct. Proc Natl Acad Sci USA 97:14455–14460 Lengeler KB, Fox DS, Fraser JA, Allen A, Forrester K, Dietrich FS, Heitman J (2002) Mating-type locus of Cryptococcus neoformans: a step in the evolution of sex chromosomes. Eukaryot Cell 1:704–718 Li AZ, Xu XF, Lin FX, Cheng SM, Lin FC (2007) Cloning and identification of partial DNA fragment for the B mating-type factor in Lentinula edodes using degenerate PCR. World J Microbiol Biotechnol 23:411–415 Li L, Shen G, Zhang ZG, Wang YL, Thompson JK, Wang P (2007) Canonical heteromeric G proteins regulating mating and virulence of Cryptococcus neoformans. Mol Biol Cell 18:4201–4209 Lin X (2009) Cryptococcus neoformans: Morphogenesis, infection, and evolution. Infect Genet Evol 9:401–416 Lin X, Heitman J (2006) The biology of the Cryptococcus neoformans species complex. Ann Rev Microbiol 60:69–105 Lin X, Heitman J (2007) Mechanisms of homothallism in fungi and transitions between heterothallism and homothallism. In: Heitman J, Kronstad JW, Taylor JW, Casselton LA (eds) Sex in fungi. Molecular determination and evolutionary implications. ASM Press, Washington, DC, pp 35–58 Lin X, Hull CM, Heitman J (2005) Sexual reproduction between partners of the same mating type in Cryptococcus neoformans. Nature 434:1017–1021 Lin X, Huang JC, Mitchell TG, Heitman J (2006) Virulence attributes and hyphal growth of C. neoformans are quantitative traits and the MATa allele enhances filamentation. PLoS Genet 2:e187 Lin X, Litvintseva AP, Nielsen K, Patel S, Floyd A, Mitchell TG, Heitman J (2007) aADa hybrids of Cryptococcus neoformans: Evidence of same sex mating in nature and hybrid fitness. PLoS Genet 3:1975–1990 Lin X, Patel S, Litvintseva AP, Floyd A, Mitchell TG, Heitman J (2009) Diploids in the Cryptococcus neoformans serotype A population homozygous for the a mating type originate via unisexual mating. PLoS Pathogens 5:e1000283 Lin X, Jackson JC, Feretzaki M, Xue C, Heitman J (2010) Transcription factors Mat2 and Znf2 operate cellular circuits orchestrating opposite- and same-sex mating in Cryptococcus neoformans. PLoS Genet 6:e1000953 Litvintseva AP, Marra RE, Nielsen K, Heitman J, Vilgalys RJ, Mitchell TG (2003) Evidence of sexual recombination among Cryptococcus neoformans serotype A isolates in sub-Saharan Africa. Eukaryot Cell 2:1162–1168

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality Litvintseva AP, Thakur R, Vilgalys R, Mitchell TG (2006) Multilocus sequence typing reveals three genetic subpopulations of Cryptococcus neoformans var. grubii (serotype A), including a unique population in Botswana. Genetics 172:2223–2238 Liu Y, Steenkamp ET, Brinkmann H, Forget L, Philippe H, Lang BF (2009) Phylogenomic analyses predict sistergroup relationship of nucleariids and fungi and paraphyly of zygomycetes with significant support. BMC Evol Biol 9:11 Lukens L, Yicun H, May G (1996) Correlation of genetic and physical maps at the A mating-type locus of Coprinus cinereus. Genetics 144:1471–1477 Luo XC, Cheng LG (1989) Sexuality and formation of monokaryotic fruitbody in Auricula auricula and Auricularia auricula. Mushroom J Trop 9:12–28 Mahlert M, Vogler C, Stelter K, Hause G, Basse CW (2009) The a2 mating-type-locus gene lga2 of Ustilago maydis interferes with mitochondrial dynamics and fusion, partially in dependence on a Dnm1-like fission component. J Cell Sci 122:2402–2412 Martinez D, Larrondo LF, Putnam N, Gelpke MDS, Huang K, Chapman J, Helfenbein KG, Ramaiya P, Detter JC, Larimer F, Coutinho PM, Henrissat B, Berka R, Cullen D, Rokhsar D (2004) Genome sequence of the lignocellulose degrading fungus Phanerochaete chrysosporium strain RP78. Nature Biotech 22: 695–700 Martinez D, Challacome J, Morgenstern I, Hibbett D, Schmoll M, Kubicek CP, Ferreira P, Ruiz-Duenas FJ, Martinez AT, Kersten P, Hammel KE, Vanden Wymelenberg A, Gaskell J, Lindquist E, Sabat G, BonDurant SS, Larrondo LF, Canessa P, Vicuna R, Yadav J, Doddapaneni H, Subramanian V, Pisabarro AG, Lavon JL, Oguiza JA, Master E, Henrissat B, Coutinho PM, Harris P, Magnuson JK, Baker SE, Bruno K, Kenealy W, Hoegger PJ, Ku¨es U, Ramaiya P, Lucash S, Salamov A, Shapiro H, Tu H, Chee CL, Mira M, Xie G, Teter S, Yaver D, James T, Mokrejs M, Pospisek M, Grigoriev IV, Brettin T, Rokhsar D, Berka R, Cullen D (2009) Genome, transcriptome, and secretome of the wood decay fungus Postia placenta supports unique mechanisms of lignocellulose conversion. Proc Natl Acad Sci USA 106:1954–1959 Martinez-Espinoza AD, Gerhardt SA, Sherwood JE (1993) Morphological and mutational analysis of mating in Ustilago hordei. Exp Mycol 17:200–214 May G, Matzke R (1995) Recombination and variation at the A mating-type of Coprinus cinereus. Mol Biol Evol 12:794–802 May G, Le Chevanton L, Pukkila PJ (1991) Molecular analysis of the Coprinus cinereus mating type-A factor demonstrates an unexpectedly complex structure. Genetics 128:529–538 May G, Shaw F, Badrane H, Vekemans X (1999) The signature of balancing selection: fungal mating compatibility gene evolution. Proc Natl Acad Sci USA 96: 9172–9177 McClelland CM, Fu J, Woodlee GL, Seymour TS, Wickes BL (2002) Isolation and characterisation of the Cryptococcus MATa pheromone gene. Genetics 160:935–947

155

McClelland CM, Chang YC, Varma A, Kwon-Chung KJ (2004) Uniqueness of the mating system in Cryptococcus neoformans. Trends Microbiol 12:208–212 McCusker JH (2006) Saccharomyces cerevisiae: an emerging and model pathogenic fungus. In: Heitman J, Filler SG, Edwards JE, Mitchell AP (eds) Molecular principles of fungal pathogenesis. ASM Press, Washington, DC, pp 245–259 McLaughlin DJ, Hibbett DS, Lutzoni F, Spatafora JW, Vilgalys R (2009) The search for the fungal tree of life. Trends Microbiol 17:488–497 Meinhardt F, Esser K (1990) Sex determination and sexual differentiation in filamentous fungi. Crit Rev Plant Sci 9:329–341 Mendoza-Mendoza A, Eskova A, Weise C, Czajkowski R, Kahmann R (2009) Hpa2 regulates the pheromone response transcription factor prf1 in Ustilago maydis. Mol Microbiol 72:683–698 Metin B, Findley K, Heitman J (2010) The mating type locus (MAT) and sexual reproduction of Cryptococcus heveanensis: insights into the evolution of sex and sex-determining chromosomal regions in fungi. PLoS Genet 6:e1000961 Metzenberg RL, Glass NL (1990) Mating type and mating strategies in Neurospora. BioEssays 12:53–59 Miyakawa T, Miyama R, Tabata M, Tsuchiya E, Fukui S (1985a) A study on the biosynthesis of tremerogen A-10, a polyisoprenyl peptide mating pheromone of Tremella mesenterica, using an inhibitor of mevalonate synthesis. Agric Biol Chem 49:1343–1347 Miyakawa T, Tabata M, Tsuchiya E, Fukui S (1985b) Biosynthesis and secretion of tremerogen A-10, a polyisoprenyl peptide mating pheromone of Tremella mesenterica. Eur J Biochem 147:489–493 Mondego JMC, Carazzolle MF, Costa GGL, Formighieri EF, Parizzi LP, Rincones J, Cotomacci C, Carraro DM, Cunha AF, Carrer H, Vidal RO, Estrela RC, Garcı´a O, Thomazella DPT, de Oliviera BV, Pires ABL, Rio MCS, Arau´jo MRR, de Morares MH, Castro LAB, Gramachi KP, Gonc¸alves MS, Moura Neto JP, Go´es Neto A, Barbosa LV, Guiltinan MJ, Bailey BA, Meinhardt LW, Cascardo LCM, Pereira GAG (2008) A genome survey of Moniliophthora perniciosa insights into Witches´ Broom Disease of cacao. BMC Genomics 9:548 Morrow CA, Fraser JA (2009) Sexual reproduction and dimorphism in the pathogenic basidiomycetes. FEMS Yeast Res 9:161–177 Moore TD, Edman JC (1993) The a-mating type locus of Cryptococcus neoformans contains a peptide pheromone gene. Mol Cell Biol 13:1962–1970 Mukherjee K, Brocchieri L, Bu¨rglin TR (2009) A comparative classification and evolutionary analysis of plant homeobox genes. Mol Biol Evol 26:2775–2794 Murata Y, Kamada T (2009) Identification of new mutant alleles of pcc1 in the homobasidiomycete Coprinopsis cinerea. Mycoscience 50:137–139 Murata Y, Fujii M, Zolan M, Kamada T (1998) Molecular analysis of pcc1, a gene that leads to A-regulated sexual morphogenesis in Coprinus cinereus. Genetics 149:1753–1761

156

Ursula Ku¨es et al.

Mutasa ES, Tymon AM, Go¨ttgens B, Mellon FM, Little PFR, Casselton LA (1990) Molecular organization of an A-mating type factor of the basidiomycete fungus Coprinus cinereus. Curr Genet 18:233–229 Naider F, Becker JM (2004) The a-factor mating pheromone of Saccharomyces cerevisiae: a model for studying the interaction of peptide hormones and G-coupled receptor. Peptides 25:1441–1463 Narisawa K, Yamaoka Y, Katsuya K (1995) Mating type of isolates derived from the spermogonial state of Puccinia coronata var. coronata. Mycoscience 35:131–135 Neiman M, Meirmans S, Meirmans PG (2009) What can asexual lineages tell us about the maintenance of sex? Ann NY Acad Sci 1168:185–200 Nichols CB, Fraser JA, Heitman J (2004) PAK kinases Ste20 and Pak1 govern cell polarity at different stages of mating on Cryptococcus neoformans. Mol Biol Cell 15:4476–4489 Niculita-Hirzel H, Labbe´ J, Kohler A, Le Tacon F, Martin F, Sanders IR, Ku¨es U (2008) Gene organization of the mating type regions in the ectomycorrhizal fungus Laccaria bicolor reveals distinct evolution between the two mating type loci. New Phytol 180:329–342 Nielsen K, Cox GM, Wang P, Toffaletti DL, Perfect JR, Heitman, J (2003) Sexual cycle of Cryptococcus neoformans var. grubii and virulence of congenic a and a isolates. Infect Immun 71:4831–4841 Nielsen K, De Obaldia AL, Heitman J (2007) Cryptococcus neoformans mates on pigeon guano: implications for the realized ecological niche and globalization. Eukaryot Cell 6:949–959 Nielsen O, Egel R (2007) The mat genes of Schizosaccharomyces pombe: Expression, homothallic switch, and silencing. In: Heitman J, Kronstad JW, Taylor JW, Casselton LA (eds) Sex in fungi. Molecular determination and evolutionary implications. ASM Press, Washington, DC, pp 143–158 Nobles MK (1943) A contribution towards a clarification of the Trametes serialis complex. Can J Res C 21: 211–234 North J (1990) Linkage map of Coprinus cinereus (Schaeff. ex Fr.) S.F.Gray. In: O´Brien SJ (ed) Genetic maps. Book 3. Lower eukaryotes. Fifth edition. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, pp 3.70–3.81 Ohm RA, de Jong JF, Lugones LG, Aerts A, Kothe E, Stajich JE, de Vries RP, Record E, Levasseur A, Baker SE, Bartholomew KA, Coutinho PM, Erdmann S, Fowler TJ, Gathman AC, Lombard V, Henrissat B, Knabe N, Ku¨es U, Lilly WW, Lindquist E, Lucas S, Magnusson JK, Oiumi F, Raudaskoski M, Salamov A, Schmutz J, Schwarze FWMR, vanKuyk PA, Horton JS, Grigoriev IV, Wo¨sten HAB (2010) Formation of mushrooms and lignocellulose degradation in the genome sequence of Schizophyllum commune. Nat Biotechnol 28:957–963 Ohno S (1967) Sex chromosomes and sex-linked genes. Springer, New York, NY Okabayashi K, Kano R, Watanabe T, Hasegawa A (2006) Serotypes and mating types of clinical isolates from feline cryptococcosis in Japan. J Vet Med Sci 68:91–94

Olesnicky NS, Brown AJ, Dowell SJ, Casselton LA (1999) A constitutively active G-protein-coupled receptor causes mating self-compatibility in the mushroom Coprinus. EMBO J 18:2756–2763 Olesnicky NS, Brown AJ, Honda Y, Dyos SL, Dowell SJ, Casselton LA (2000) Self-compatible B mutants in Coprinus with altered pheromone-receptor specificities. Genetics 156:1025–1033 Ongay-Larios L, Navarro-Olmes R, Kawasaki L, VelzquezZavala N, Sa´nchez-Paredes E, Torres-Quiroz F, Coello G, Coria R (2007) Kluyveromyces lactis sexual pheromones. Gene structures and cellular responses to afactor. FEMS Yeast Res 7:740–747 O’Shea SF, Chaure PT, Halsall JR, Olesnicky NS, Leibbrandt A, Connerton IF, Casselton LA (1998) A large pheromone and receptor gene complex determines multiple B mating type specificities in Coprinus cinereus. Genetics 148:1081–1090 Panwar SL, Legrand M, Dignard D, Whiteway M, Magee PT (2003) MFa1, the gene encoding the a mating pheromone of Candida albicans. Eukaryot Cell 2:1350–1360 Pardo EH, O´Shea SF, Casselton LA (1996) Multiple versions of the A mating type locus of Coprinus cinereus are generated by three paralogous pairs of multiallelic homeobox genes. Genetics 144:87–94 Park Y-D, Panepinto J, Sin S, Larsen P, Giles S, Williamson PR (2010) Mating pheromone in Cryptococcus is regulated by a transcriptional/degradative “futile” cycle. J Biol Chem 285:34746–34756 Pei MH, Bayon C, Ruiz C, Tubby I (2007) Population structure of poplar rust Melampsora larici-populina in the UK inferred from AFLP. Plant Pathol 56: 472–479 Piepenbring M, Stoll M, Oberwinkler F (2002) The generic position of Ustilago maydis, Ustilago scitaminea, and Ustilago esculenta (Ustilaginales). Mycol Prog 1:71–80 Phadke SS, Zufall RA (2009) Rapid diversification of mating systems in ciliates. Biol J Linn Soc 98:187–197 Po¨ggeler S (2007) MAT and its role in homothallic ascomycete Sordaria macrospora. In: Heitman J, Kronstad JW, Taylor JW, Casselton LA (eds) Sex in fungi. Molecular determination and evolutionary implications. ASM Press, Washington, DC, pp 171–188 Puhalla JE (1970) Genetic studies of the b incompatibility locus of Ustilago maydis. Genet Res 16:229–232 Qu P, Aratani A, Syoji T, Toda T, Kubota M, Hyakumachi M (2008a) Use of single-protoplast isolates in the study of the mating phenomena of Rhizoctonia solani (Thanatephorus cucumeris) AG-1 IC and IA. Mycoscience 49:132–137 Qu P, Yamashita K, Toda T, Priyatmojo A, Kubota M, Hyakumachi M (2008b) Heterokaryon formation in Thanatephorus cucumeris (Rhizoctonia solani) AG-1 IC. Mycol Res 112:1088–1100 Raeder UW, Thompson W, Broda P (1989) RFLP-based genetic map of Phanerochaete chrysosporium ME446: lignin peroxidase genes occur in clusters. Mol Microbiol 3:911–918 Raper CA (1990) Schizophyllum commune. In: O´Brien SJ (ed) Genetic maps. Book 3. Lower eukaryotes. Fifth

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality edition. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, pp 3.90–3.91 Raper CA (1998) Schizophyllum commune, a model for genetic studies of the Basidiomycetes. In: Sidhu GS (ed) Genetics of plant pathogenic fungi. Academic Press, London, UK, pp 511–522 Raper JR (1966) Genetics of sexuality of higher fungi. Ronald Press, New York, NY Raper JR, Flexer AS (1971) Mating systems and evolution of the basidiomycetes. In: Petersen RH (ed) Evolution of the higher basidiomycetes. University of Tennessee Press, Knoxville, Tennessee, pp 149–167 Raper JR, Krongelb (1958) Genetic and environmental aspects of fruiting in Schizophyllum commune Fr. Mycologia 50:707–740 Raper JR, Baxter MG, Middleton RB (1958) The genetic structure of the incompatibility loci in Schizophyllum. Proc Natl Acad Sci USA 44:889–900 Raper JR, Baxter MG, Ellingboe AH (1960) The genetic structure of the incompatibility factors of Schizophyllum commune – the A factor. Proc Natl Acad Sci USA 46: 833–842 Raper JR, Boyd DH, Raper CA (1965) Primary and secondary mutations at the incompatibility loci in Schizophyllum. Proc Natl Acad Sci USA 53:1324–1332 Raudaskoski M (1998) The relationship between B-mating-type genes and nuclear migration in Schizophyllum commune. Fungal Genet Biol 24:207–227 Regenfelder E, Spellig T, Hartmann A, Lauenstein S, Bo¨lker M, Kahmann R (1997) G proteins in Ustilago maydis: Transmission of multiple signals? EMBO J 16:1934–1942 Ren P, Roncaglia P, Springer DJ, Fan J, Chaturvedi V (2005) Genomic organization and expression of 23 new genes from MATa locus of Cryptococcus neoformans var. gattii. Biochim Biophys Res Commun 326:233–241 Ren P, Springer DJ, Behr MJ, Samsonoff WA, Chaturvedi S, Chaturvedi V (2006) Transcription factor STE12a has distinct roles in morphogenesis, virulence, and ecological fitness of the primary pathogenic yeast Cryptococcus gattii. Eukaryot Cell 5:1065–1080 Riquelme M, Challen MP, Casselton LA, Brown AJ (2005) The origin of multiple B mating specificities in Coprinus cinereus. Genetics 170:1105–1119 Robertson CI, Kende AM, Toenjes K, Novotny CP, Ullrich RC (2002) Evidence for interaction of Schizophyllum commune Y mating-type proteins in vivo. Genetics 160:1461–1467 Rodriguez-Carres M, Findley K, Sun S, Dietrich FS, Heitman J (2010) Morphological and genomic characterization of Filobasidiella depauperata: A homothallic sibling species of the pathogenic Cryptococcus species complex. PLoS ONE 5:e9620 Romeis T, Ka¨mper J, Kahmann R (1997) Single-chain fusions of two unrelated homeodomain proteins trigger pathogenicity in Ustilago maydis. Proc Natl Acad Sci USA 94:1230–1234 Romeis T, Brachmann A, Kahmann R, Ka¨mper J (2000) Identification of a target gene for the bE-bW homeodomain protein complex in Ustilago maydis. Mol Microbiol 37:54–66

157

Rusche LN, Rine J (2010) Switching the mechanism of mating type switching: a domesticated transposase supplants a domesticated homing endonuclease. Genes Dev 24:10–14 Rydholm C, Dyer PS, Lutzoni F (2007) DNA sequence characterization and molecular evolution of MAT1 and MAT2 mating-type loci of the self-compatible ascomycete mold Neosartorya fischeri. Eukaryot Cell 6:868–874 Sakagami Y, Isogai A, Suzuki S, Tamara S, Kitada C, Fujino M (1979) Structure of tremerogen A-10, a peptidyl hormone inducing conjugation tube formation in Tremella mesenterica. Agric Biol Chem 43:2643–2645 Sakagami Y, Yoshida M, Isogai A, Suzuki A (1981a) Peptidal sex hormones inducing conjugation tube formation in compatible mating-type cells of Tremella mesenterica. Science 212:1525–1527 Sakagami Y, Yoshida M, Isogai A, Suzuki A (1981b) Structure of tremerogen a-13, a peptidyl hormone of Tremella mesenterica. Agric Biol Chem 45:1045–1047 Saracli MA, Yildiran ST, Sener K, Gonlum A, Doganci L, Keller SM, Wickes BL (2006) Genotyping of Turkish environmental Cryptococcus neoformans var. neoformans isolates by pulsed field gel electrophoresis and mating type. Mycoses 49:124–129 Saul N, Krockenberger M, Carter D (2008) Evidence of recombination in mixed-mating-type and a-only populations of Cryptococcus gattii sourced from single Eucalyptus tree hollows. Eukaryot Cell 7:727–734 Schirawski J, Heinze B, Wagenknecht M, Kahmann R (2005) Mating type loci of Sporisorium reilianum: novel pattern with three a and multiple b specificities. Eukaryot Cell 4:1317–1327 Schlesinger R, Kahmann R, Ka¨mper J (1997) The homeodomain of the heterodimeric bE and bW proteins of Ustilago maydis are both critical for function. Mol Gen Genet 254:514–519 Schulz B, Banuett F, Dahl M, Schlesinger R, Scha¨fer W, Martin T, Herskowitz I, Kahmann R (1990) The b alleles of U. maydis, whose combinations program pathogenic development, code for polypeptides containing a homeodomain-related motif. Cell 60: 295–306 Shen G-P, Park DC, Ullrich RC, Novotny CP (1996) Characterization of a Schizophyllum gene with Ab6 mating-type activity. Curr Genet 29:136–142 Shen G-P, Chen Y, Song D, Peng Z, Novotny CP, Ullrich RC (2001) The Aa6 locus: its relation to mating-type regulation of sexual development in Schizophyllum commune. Curr Genet 39:340–345 Shen WC, Davidson RC, Cox GM, Heitman J (2002) Pheromones stimulate mating and differentiation via paracrine and autocrine signaling in Cryptococcus neoformans. Eukaryot Cell 1:366–377 Sherwood JE, Kosted PJ, Anderson CM, Gerhardt SA (1998) Production of a mating inhibitor of Ustilago hordei. Phytopathol 88:456–464 Silva J (1972) Alleles at the b incompatibility locus in Polish and North American populations of Ustilago maydis (CD) Corda. Physiol Plant Pathol 2:333–337

158

Ursula Ku¨es et al.

Snetselaar KM, Bo¨lker M, Kahmann R (1996) Ustilago maydis mating hyphae orient their growth toward pheromone sources. Fungal Genet Biol 20:299–312 Specht CA (1995) Isolation of the Ba and Bb matingtype loci of Schizophyllum commune. Curr Genet 28:374–379 Specht CA, Stankis MM, Giasson L, Novotny CP, Ullrich RC (1992) Functional analysis of the homeodomainrelated proteins of the Aa locus of Schizophyllum commune. Proc Natl Acad Sci 89:7174–7178 Specht CA, Stankis MM, Novotny CP, Ullrich RC (1994) Mapping the heterogeneous DNA region that determines the 9 Aa-mating-type specificities of Schizophyllum commune. Genetics 137:709–714 Spellig T, Bo¨lker M. Lottspeich F, Frank RW, Kahmann R (1994) Pheromones trigger filamentous growth in Ustilago maydis. EMBO J 13:1620–1627 Spiers AG, Hopcroft DH (1994) Comparative studies of the poplar rusts Melampsora medusae, M. larici-populina and their interspecific hybrid M. medusae-populina. Mycol Res 98:889–903 Spit A, Hyland RH, Mellor EJ, Casselton LA (1998) A role for heterodimerization in nuclear localization of a homeodomain protein. Proc Natl Acad Sci USA 95:6228–6233 Srivilai P (2006) Molecular analysis of genes acting in fruiting body development in basidiomycetes. PhD thesis, Georg-August-University Go¨ttingen, Go¨ttingen, Germany Stahl U, Esser K (1976) Genetics of fruit body production in higher basidiomycetes. 1. Monokaryotic fruiting and its correlation with dikaryotic fruiting in Polyporus ciliatus. Mol Gen Genet 148:183–197 Stajich JS, Wilke SK, Ahre´n D, Au CH, Birren BW, Borodovsky M, Burns C, Canba¨ck B, Casselton LA, Cheng CK, Deng J, Dietrich FS, Fargo DC, Farman ML, Gathman AC, Goldberg J, Guigo R, Hoegger PJ, Hooker JB, Huggins A, James TY, Kamada T, Kilaru S, Kodira C, Ku¨es U, Kupfer D, Kwan HS, Lomsadze A, Li W, Lilly WW, Ma L-J, Mackey AJ, Manning G, Martin F, Muraguchi H, Natvig DO, Palmeri H, Ramesh MA, Rehmeyer CJ, Roe BA, Shenoy N, Stanke M, Ter-Hovhannisyan V, Tunlid A, Velagapudi R, Vision TJ, Zeng Q, Zolan ME, Pukkila PJ (2010) Insights into evolution of multicellular fungi from the assembled chromosomes of the mushroom Coprinopsis cinerea (Coprinus cinereus). Proc Natl Acad Sci USA107:11655–11665 Stankis MM, Specht CA, Yang HL, Giasson L, Ullrich RC, Novotny CP (1992) The Aa mating locus of Schizophyllum commune encodes 2 dissimilar multiallelic homeodomain proteins. Proc Natl Acad Sci USA 89:7169–7173 Stanton BC, Hull CM (2007) Mating-type locus control of cell identity. In: Heitman J, Kronstad JW, Taylor JW, Casselton LA (eds) Sex in fungi. Molecular determination and evolutionary implications. ASM Press, Washington, DC, pp 59–74 Stanton BC, Giles SS, Kruzel EK, Warren CL, Ansari AZ, Hull CM (2009) Cognate site identifier analysis reveals novel binding properties of the sex inducer

homeodomain proteins of Cryptococcus neoformans. Mol Microbiol 72:1334–1347 Statzell-Tallman A, Fell JW (1998) Sporidiobolus Nyland. In: Kurtzman CP and Fell JW (eds) The yeasts, a taxonomic study, 4th edn. Elsevier, Amsterdam, The Netherlands, pp 693–699 Statzell-Tallman A, Belloch C, Fell JW (2008) Kwoniella mangroviensis gen. nov, sp. nov. (Tremellales, Basidiomycota), a teleomorphic yeast from mangrove habitats in the Florida Everglades and Bahamas. FEMS Yeast Res 8:103–113 Stoll M, Begerow D, Oberwinkler F (2005) Molecular phylogeny of Ustilago, Sporisorium, and related taxa based on combined analyses of rDNA sequences. Mycol Res 109:342–356 Swamy S, Uno I, Ishikawa T (1984) Morphogenetic effects of mutations at the A and B incompatibility factors in Coprinus cinereus. J Gen Microbiol 130:3219–3224 Takemaru T (1961) Genetical study on fungi. XI. The mating system in Lentinus edodes. Rep Tottori Mycol Inst 1:61–68 Tamura K, Dudley J, Nei M, Kumar S (2007) MEGA4: Molecular evolutionary genetic analysis (MEGA) software version 4.0. Mol Biol Evol 24:1596–1599 Tetsuka Y, Katsuya K (1984) Mating experiments of isolates derived from the spermogonial state of Gymnosporangium asiaticum. Proc Japan Acad 60, Ser B:149– 152 Thompson W, Broda P (1987) Mating behaviour in an isolate of Phanerochaete chrysosporium. Trans Br Mycol Soc 89:285–294 Toda T, Hyakumach M (2006) Heterokaryon formation in Thanetophorus cucumeris anastomosis group 2-2IV. Mycologia 98:726–736 Tokimoto K, Komatsu M, Takemaru T (1973) Incompatibility factors in the natural population of Lentinus edodes in Japan. Rep Tottori Mycol Inst 10:371–376 Trail F, Mills D (1990) Growth of haploid Tilletia strains in planta and genetic analysis of a cross of Tilletia caries X T. controversa. Phytopathol 80:367–370 Tscharke RL, Lazera M, Chang YC, Wickes BL, KwonChung KJ (2003) Haploid fruiting in Cryptococcus neoformans is not mating type a-specific. Fungal Genet Biol 39:230–237 Tsuchiya E, Fukui S (1987) Binding of Rhodotorucine A, a lipopeptidyl mating hormone, to A cells of Rhodosporidium toruloides for induction of sexual differentiation. Biochem Biophys Res Comm 85:473–479 Turgeon G, Debuchy R (2007) Cochliobolus and Podospora: Mechanism of sex determination and the evolution of reproductive life style. In: Heitman J, Kronstad JW, Taylor JW, Casselton LA (eds) Sex in fungi. Molecular determination and evolutionary implications. ASM Press, Washington, DC, pp 93–122 Tymon AM, Ku¨es U, Richardson WVJ, Casselton LA (1992) A fungal mating type protein that regulates sexual and asexual development contains a POUrelated domain. EMBO J 11:1805–1813 Ullrich RC, Raper JR (1975) Primary homothallism – Relation to heterothallism in the regulation of sexual morphogenesis in Sistotrema. Genetics 80:311–321

Mating Type in Basidiomycetes: Unipolar, Bipolar, and Tetrapolar Patterns of Sexuality Urban M, Kahmann R, Bo¨lker M (1996a) The biallelic a mating type locus of Ustilago maydis: Remnants of an additional pheromone gene indicate evolution from a multiallelic ancestor. Mol Gen Genet 250:414–420 Urban M, Kahmann R, Bo¨lker M (1996b) Identification of the pheromone response element in Ustilago maydis. Mol Gen Genet 251:31–37 Vaillancourt LJ, Raudaskoski M, Specht CA, Raper CA (1997) Multiple genes encoding pheromones and a pheromone receptor define the Bb1 mating-type specificity in Schizophyllum commune. Genetics 146:541–551 Vale´rio E, Gadanho M, Sampaio JP (2008a) Reappraisal of the Sporobolomyces roseus species complex and description of Sporidiobolus metaroseus sp. nov. Int J Syst Evol Microbiol 58:736–741 Vale´rio E, Gadanho M, Sampaio JP (2008b) Sporidiobolus johnsonii and Sporidiobolus salmenicolor revisited. Mycol Prog 7:125–131 van der Nest MA, Slippers B, Stenlid J, Wilken PM, Vasaitis R, Wingfield MJ, Wingfield BD (2008) Characterization of the system governing sexual and selfrecognition in the white rot homobasidiomycete Amylostereum areolatum. Curr Genet 53:323–336 van der Nest MA, Slippers B, Steenkamp ET, De Vos L, Van Zyl K, Stenlid J, Wingfield MJ, Wingfield BD (2009) Genetic linkage map for Amylostereum areolatum reveals an association betveen vegetative growth and sexual and self-recognition. Fungal Genet Biol 46:632–641 Viviani MA, Esposto MC, Cogliati M, Montagna MT, Wickes BL (2001) Isolation of a Cryptococcus neoformans serotype A MATa strain from the Italian environment. Med Mycol 39:383–386 Viviani MA, Cogliati M, Esposto MC, Lemmer K, Tintelnot K, Valiente MFC, Swinne D, Velegraki A, Velho R (2006) Molecular analysis of 311 Cryptococcus neoformans isolates from a 30-month ECMM survey of cryptococcosis in Europe. FEMS Yeast Res 6: 614–619 Votintserva AA, Filatov DA (2009) Evolutionary strata in a small mating-type-specific region of the smut fungus Microbotryum violaceum. Genetics 182:1391–1396 Wahl R, Zahiri A, Ka¨mper J (2010) The Ustilago b mating type locus controls hyphal proliferation and expression of secreted virulence factors in planta. Mol Microbiol 75:208–220 Wang P, Perfect JR, Heitman J (2000) G-protein b subunit Gpb1 is required for mating and haploid fruiting in Cryptococcus neoformans. Mol Cell Biol 20:352–362 Wang P, Nichols CB, Lengeler KB, Cardenas ME, Cox GM, Perfect PR, Heitman J (2002) Mating-type-specific and nonspecific PAK kinases play shared and divergent roles in Cryptococcus neoformans. Eukaryot Cell 1:257–272 Wendland J, Vaillancourt LJ, Hegner J, Lengeler KB, Laddison KJ, Specht CA, Raper CA, Kothe E (1995) The mating type locus Bb1 of Schizophyllum commune contains a pheromone receptor gene and putative pheromone genes. EMBO J 14:5271–5278 Whitehouse HLK (1949) Multiple allelomorph heterothallism in fungi. New Phytol 48:212–244

159

Whitehouse HLK (1952) A survey of heterothallism in the Ustilaginales. Trans Br Mycol Soc 340–355 Wickes BL, Mayorga ME, Edman U, Edman JC (1996) Dimorphism and haploid fruiting in Cryptococcus neoformans: association with the a-mating type. Proc Natl Acad Sci USA 93:7327–7331 Wickes BL, Edman U, Edman JC (1997) The Cryptococcus neoformans STE12a gene: a putative Saccharomyces cerevisiae STE12 homologue that is mating type specific. Mol Microbiol 26:951–960 Wong GJ, Wells K (1987) Comparative morphology, compatibility, and interfertility of Auricularia cornea, A. polytricha, and A. tenuis. Mycologia 79:847–858 Wright S, Finnegan D (2001) Genome evolution: sex and the transposable element. Curr Biol 11:R296–R299 Wu C, Huang Y, Yinbing B (2004) RAPD/BSA analysis on monokaryotic strains of different mating types of Auricularia auricula. J Huazhoung Agric Univ, DOI: CNKI:SUN:HZNY.0.2004-01-030 Xu J, Saunders CW, Hu P, Grant RA, Boekhout T, Kuramee 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 Natl Acad Sci USA 104:18730–18735 Xue CY, Hsueh Y-P, Heitman J (2008) Magnificent seven: roles of G protein-coupled receptors in extracellular fungi. FEMS Microbiol Rev 32:1010–1032 Yamazaki S, Katsuya K (1988) Mating type of pine gall rust fungus, Cronartium quercum. Proc Jpn Acad Ser B 64:197–200 Yan Z, Li XG, Xu JP (2002) Geographic distributions of mating type alleles of Cryptococcus neoformans in four areas of the United States. J Clin Microbiol 40:965–972 Yee AR, Kronstad JW (1998) Dual sets of chimeric alleles identify specificity sequences for the bE and bW mating and pathogenicity genes of Ustilago maydis. Mol Cell Biol 18:221–232 Yi R, Tachikawa T, Ishikawa M, Mukaiyama H, Bao D, Aimi T (2009) Genomic structure of the A matingtype locus in a bipolar basidiomycete, Pholiota nameko. Mycol Res 113:240–248 Yi R, Mukaiyama H, Tachikawa T, Shimomura N, Aimi T (2010) A-mating type gene expression can drive clamp cell formation in the bipolar mushroom Pholiota microspora (Pholiota nameko). Eukaryot Cell 9:1109–1119 Yockteng R, Marthey S, Chiapello HJ, Gendrault A, Hood ME, Rodolphe F, Devier B, Wincker P, Dossat C, Giraud T (2007) Expressed sequences tags of the anther smut fungus, Microbotryum violaceum, identfy mating and pathogenicity genes. BMC Genomics 8:272 Yu Z, Liang J, Cao Z, Guo Z, Dan J, Zhao G (2009) Nuclear behaviour in the life cycle of Melampsora laricipopulina Kleb. J Food Agric Env 7:791–794 Yue CL, Osier M, Novotny CP, Ullrich RC (1997) The specificity determinant of the Y mating-type proteins

160

Ursula Ku¨es et al.

of Schizophyllum commune is also essential for the Y-Z protein binding. Genetics 145:253–260 Yue CL, Cavallo LM, Alspaugh JA, Wang P, Cox GM, Perfect JR, Heitman J (1999) The STE12a homolog is required for haploid filamentation but largely dispensable for mating and virulence in Cryptococcus neoformans. Genetics 153:1601–1615 Zarnack K, Eichhorn H, Kahmann R, Feldbru¨gge M (2008) Pheromone-regulated target genes respond differen-

tially to MAPK phosphorylation of transcription factor Prf1. Mol Microbiol 69:1041–1053 Zakharov IA (2005) Intratetrad mating and its genetic and evolutionary consequences. Russian J Genet 41:402–411 Zhao X, Mehrabi R, Xu J (2007) Mitogen-activated protein kinase pathways and fungal pathogenesis. Eukaryot Cell 6:1701–1714

Evolution of Mutualistic Systems and Metabolism in Fungi

.

7

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

ARTHUR SCHU¨ßLER1, CHRISTOPHER WALKER2,3

CONTENTS I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. History of Glomeromycota Research . . . . . . . . . . . A. The Phase Before the Linkage of Glomeromycotan Fungi and AM was Known . . . . . . . . . . . . . . . . . . . . . . . . . B. The Linkage of Glomeromycotan Fungi with AM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. The ‘Modern’ Phase of Glomeromycota Research . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . III. Molecular Phylogeny of Glomeromycota . . . . . . A. The Impact of Understanding AMF Phylogenetic Relationships . . . . . . . . . . . . . . . . . . B. Phylogenies and a ‘Natural’ Systematic and Taxonomy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Multi-Gene Analysis and Analyses of Protein Coding Genes. . . . . . . . . . . . . . . . . . . . . IV. The Origin of Glomeromycota and the First Land Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. The Fossil Record . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Molecular Clock Estimates . . . . . . . . . . . . . . . . . . V. Glomeromycota and Land Plant Evolution . . . . A. The Geosiphon–Nostoc Endosymbiosis . . . . . B. Evolutionary Scenarios for Land Plant Evolution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Glomeromycota Endobacteria Co-Evolved with AMF (and Plants) for at Least 400 MY . . . . . . . . . . . . . . . . . . . . . . . . . . VI. Glomeromycota Asexual Evolution. . . . . . . . . . . . . A. The Oldest Known Eukaryotic Asexuals are Thought to be AMF. . . . . . . . . . . . . . . . . . . . . . B. Are AMF Hetero- or Homokaryotic?. . . . . . . . VII. Species Concepts, Systematics and Taxonomy. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Molecular Characterisation of Species . . . . . . B. Molecular Phylogeny – from Phylum Definition to Species Recognition in the Field . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Biogeography and Geology . . . . . . . . . . . . . . . . . . VIII. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1

I. Introduction 163 164 164 165 166 166 166 167 167 169 169 170 171 173 173 174 174 174 175 176 177 178 179 180 180

Genetics, Department of Biology, University of Munich (LMU), Grosshaderner Strasse 2–4, 82152 Martinsried, Germany; e-mail: [email protected] 2 Royal Botanic Garden Edinburgh, 20A Inverleith Row, Edinburgh EH3 5LR, UK 3 School of Earth Sciences and Environment, University of Western Australia, 35 Stirling Highway, Crawley, WA 6009, Australia

The understanding of phylogenetic relationships and evolution in the Fungi has considerably advanced in the past decade. Members of the phylum Glomeromycota, an evolutionary lineage considerably older than Asco- and Basidiomycota, form symbioses with photoautotrophs and are thought to be among the oldest living asexual eukaryotes (see Chapter 1 in this volume). Glomeromycotan fungi influence most terrestrial ecosystems by forming an intimate, mutualistic symbiosis, arbuscular mycorrhiza (AM), with roughly 80% of all land plants (Embryophyta), including many non-vascular ones. Because most land plants form AM, glomeromycotan fungi are key-drivers of terrestrial ecosystem’s primary productivity, significantly influencing global phosphorous and carbon cycles (Fitter 2005; Smith and Read 2008; see The Mycota vol. IV). The name of the symbiosis is derived from much branched hyphae, called arbuscules, which colonise root cortical cells and provide an interface with a huge surface area, facilitating bidirectional nutrient transfer between fungus and plant. In symbioses with rootless plants such as bryophytes, arbuscules are also formed, but in some plant associations only hyphal coils have been observed. We use the term ‘AM’ for all mutualistic associations of plants with ‘AM fungi’ (AMF), which are all members of the Glomeromycota. The AMF are inconspicuous because of their obligately symbiotic and underground lifestyle. They do not cause obvious harm to plants and thus their study does not attract so much attention (or funds) as pathogenic fungi (Benedetto and Bonfante 2004). Consequently still little is known of some of their important characteristics, for example, genome organisation, DNA ploidy levels and host preferences. Most plants would be unable to compete in their natural environment without their AMF partners. Their effects on plant Evolution of Fungi and Fungal-Like Organisms, The Mycota XIV S. Po¨ggeler and J. Wo¨stemeyer (Eds.) © Springer‐Verlag Berlin Heidelberg 2011

164

Arthur Schu¨ßler and Christopher Walker

nutrition have received most attention and different AMF species can considerably differ in nutrient acquisition ability (Munkvold et al. 2004; Smith et al. 2004). Development, composition and primary production of plant communities are strongly influenced by AMF diversity, at different phylogenetic levels (van der Heijden et al. 1998; Maherali and Klironomos 2007). These factors influence resource distribution in plant communities and host plant stress tolerance. AMF comprise a large part of soil microbial biomass (Jastrow and Miller 1997), influencing soil structure and stability (Rillig 2004) and restoration of microbial equilibrium in degraded soils. Improved plant resistance to biotic and abiotic stresses may partly be related to a better nutrition status or intraradical competition of AMF with pathogenic organisms (Maherali and Klironomos 2007), but also to specific yet unknown effects and mechanisms (Pozo and Azcon-Aguilar 2007). Such mechanisms are based on the long lasting co-evolution of the plants and AMF and are best understood in an evolutionary context. How, for example, can pathogen defence of plants be highly co-ordinated whilst still allowing the mutualistic AMF colonisation within a root? What are the underlying mechanisms that evolved in the AM, the likely foundation of the origin of land plant life as we know it, today? Understanding the evolution of Glomeromycota is challenging because they are thought to possess some extraordinary genomic features, for example, an unusually high genetic diversity within a species. Partly because of this, there are difficulties of formulating clear species concepts and defining boundaries of individuals and populations (Rosendahl 2008) and despite their fundamental place in the terrestrial ecosystem, even the detection of AMF species in the field is still difficult or impossible. Some fundamental aspects of AMF genome structure remain incompletely understood (e.g., the putatively heterokaryotic genome organisation, genome sizes in different lineages, genetic recombination frequency, general intraspecific genetic variability, the genetic structure of populations, and the definition of an individual, a genet or a clone). Taking the global impact into account, from sustainable food and biofuel production, through drought and salt resistance, to global CO2 sequestration, it is difficult to understand why the field of AM research (like mycology in general) is so much

under-represented worldwide, compared with other disciplines. Many basic aspects of AMF evolution are still unclear and can only superficially be treated here. We therefore focus, with some links to land plant evolution, on the evolution of glomeromycotan fungi from the phylum to the current concept of species. We begin with some historical aspects, move to molecular phylogeny, elaborating the recently revised ‘natural systematics’, and then discuss some fundamental evolutionary and coevolutionary aspects. We also give an overview of the current approaches characterising, describing and detecting AMF as foundations for future understanding their biogeography and related evolutionary aspects.

II. History of Glomeromycota Research The present understanding of AMF derives from study of limited duration compared with many other fungi. Until 2001, they were placed in an order (Glomerales) within the Zygomycetes and were sometimes considered para- or polyphyletic. Molecular studies then showed that they form a monophyletic clade separated from all known zygomycete lineages and merited their own phylum (Schu¨ßler et al. 2001). Historically, the study of AMF evolution, phylogeny and taxonomy can be separated into three phases. First there were purely morphological descriptions, before the link between the fungi and their plant hosts was understood. Second, once their mycorrhizal nature was established, species descriptions became more biologically meaningful and it was suggested already in 1975 that early land plant evolution may have been possible only by a mycotrophic lifestyle (Pirozynski and Malloch 1975). Third, the present phase followed incorporating molecular evidence into species descriptions, molecular ecology and phylogenetic and evolutionary analyses, including preliminary evidence of AMF genome organisation.

A. The Phase Before the Linkage of Glomeromycotan Fungi and AM was Known The first published recognition of a fungus in the group nowadays classified as Glomeromycota was in 1845 by Tulasne and Tulasne. They discovered

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

some small sporocarps in wooded environments. Recognising them as undescribed species, but without any knowledge of their biology, they named them Glomus macrocarpum and G. microcarpum. Later, they moved both species into Endogone, based on a perceived similarity of their chlamydospores with the zygospores characterising that genus (Tulasne and Tulasne 1851). A few other sporocarpic species were then described in several genera, though still nothing was known of their relationships with plants. Also around the turn of the nineteenth century, fungi were noted in roots that differed from any previously recognised organisms, one of which was suggested to be pathogenic, and named Rhizophagus (root eater) populinus (Dangeard 1900). Others were recognised as probably forming mycorrhizas, at the time a relatively new term coined by Frank (1885), and some very detailed drawings were produced by Gallaud (1905), reprinted by Smith and Read (2008). Bucholtz (1912) reviewed the genus Endogone and the first English language review of these fungi was carried out by Thaxter (1922), with all later Glomus species still maintained in the genus Endogone in the Endogoneae (¼ Endogonaceae).

B. The Linkage of Glomeromycotan Fungi with AM It was not until the second half of the twentieth century that some of these fungi were linked with the specific type of plant root colonisation (the AM) as described by, for example, Gallaud (1905). Although fruiting bodies identified as Endogone vesiculifera were traced by hyphae to AM (Peyronel 1937), Koch’s postulates were not satisfied for another 16 years when a fungus later named Glomus mosseae was proven to be the partner in a mycorrhizal symbiosis (Mosse 1953). From this time, many studies followed in which different organisms were cultured and shown to form AM, and Gerdemann and Trappe (1974) revised the known species. They separated them into the truly sexual zygomycetes (Endogone) and the remaining presumed asexual species. The genera Glomus and Sclerocystis were resurrected, and G. macrocarpum and G. microcarpum were re-established after >100 years, along with the description of new genera. Later new genera and species were all still placed within the family Endogonaceae,

165

until Pirozynski and Dalpe´ (1989) raised the family Glomeraceae (published as Glomaceae) to contain Glomus and Sclerocystis. Then, Morton and Benny (1990) showed that the species in the Glomeraceae were distant morphologically and developmentally from other zygomycetes and Endogone spp and placed all AMF, including the genera Acaulospora, Entrophospora, Sclerocystis, Scutellospora and Gigaspora in a separate order, the Glomerales (published as Glomales). This major revision was the first attempt to place AM fungi in a phylogenetic hierarchical structure. Families and subfamilies were erected from a cladistic analysis based partly on spore ontogeny, and with the assumed common production of arbuscules as a unifying basal characteristic. At that time, as now, many of the included species were known only from field collected spores, and have not yet been shown to be mycorrhiza formers. They are included as AMF by analogy with known symbiotic species. The taxonomy of AMF was mainly based on the asexual spore morphology and spore wall characters and later, spore ontogeny was sometimes included (Morton and Benny 1990; Franke and Morton 1994). This is further complicated because glomeromycotan spores probably represent different functional entities. In many species the structure of the spore wall is simple and not fundamentally different from the hyphal wall (glomoid chlamydospores), whereas in other genera (e.g., Acaulospora, Scutellospora) ‘spores’ have several ’walls’ or ‘wall layers’, develop complex germination shields (see below), and are most likely not homologous with glomoid spores. In the 1960s, morphologically different AMF spore types were named with non-Linnean concepts (e.g., Mosse and Bowen 1968). Linnean nomenclatural was applied to several AMF species, though still within Endogone, by Nicolson and Gerdemann (1968). After Thaxter (1922) retained the genera Sphaerocreas, Sclerocystis, and Glaziella in the Endogonaceae, Sphaerocreas was subsumed under Glomus by Gerdemann and Trappe (1974), With the splitting of Endogone, new genera were erected or old ones re-used, with the genera Gigaspora and Acaulospora being described as new, and Sclerocystis being resurrected to contain species with Glomus-like spores formed in sporocarps radially around a sterile central plexus. At that time, all the Gigaspora and Acaulospora species were described from pot cultures, and so were known to form AM. Sclerocystis rubiformis and Sc. coremioides also had been shown to form mycorrhizas. The genus Sclerocystis remained somewhat controversial and was combined entirely with Glomus by Redecker et al. (2000b), but

166

Arthur Schu¨ßler and Christopher Walker

some authors retained the genus. Two other genera, Modicella and Glaziella, were mentioned in Gerdemann and Trappe (1974) as members of the Endogonaceae, neither of which are considered to be mycorrhizal fungi. Glaziella has since been placed in the Ascomycota, and Modicella is in the Mortierellales.

C. The ‘Modern’ Phase of Glomeromycota Research The ‘molecular’ phase of studying phylogenetic relationships went through investigations with monoclonal antibodies, isozyme analyses, fatty acid composition studies, and eventually to the use of DNA. The first indication that obligate symbiotic fungi in the Glomeromycota are not necessarily always mycorrhizal in the strict sense came towards the end of the twentieth century, when the ‘Geosiphon-symbiosis’ of a fungus with a cyanobacterium (Nostoc punctiforme) was recognised to be formed by a glomeromycotan fungus (Schu¨ßler et al. 1994). This study led to a complete reorganisation of the systematics of the AMF (Schu¨ßler et al. 2001). Since that time it is acknowledged that sequence information is essential for understanding AMF phylogeny and for separating them into their taxa, as it has become evident that morphology alone often is inadequate for this purpose. For example, glomoid spores of Paraglomus species look like those of some Glomus spp, although hundreds of millions of years separate these lineages. Individual species of Ambispora may form two different spore morphs, one similar to Acaulospora (acaulosporoid) and the other to Glomus (glomoid) spores. It is likely, therefore, that glomoid and acaulosporoid spores in different higher taxa are not homologous (Morton and Redecker 2001). In the meantime, it is clear that several families in the Glomeromycota contain species producing more than one spore morphotype.

III. Molecular Phylogeny of Glomeromycota Until the mid-1990s, the phylogeny of AMF, classified as Glomerales within the zygomycetes, was still enigmatic and the monophyletic nature of the AMF was questioned (Morton 2000), despite evidence to the contrary from molecular studies

(Gehrig et al. 1996). Morphologically, the classification of Zygomycota always was uncertain because a zygosporic sexual stage is unknown for most organisms assigned to it (Benjamin 1979) and it became clear that families in the Zygomycota could not be sustained in a reliable phylogenetic classification (Benny et al. 2001; O’Donnell et al. 2001). The SSU rDNA phylogeny clearly revealed the monophyletic origin of AMF, and so the new phylum Glomeromycota was described to host AMF (Schu¨ßler et al. 2001). Other genes also resolved the Glomeromycota as monophyletic and a multi gene analysis (James et al. 2006) confirmed them as a very old group of terrestrial fungi.

A. The Impact of Understanding AMF Phylogenetic Relationships Phylogenetic relationships provide a foundation for understanding many aspects of biology. One example is Geosiphon pyriformis, a fungus that forms the only known fungal endosymbiosis with cyanobacteria. The revelation by molecular phylogenetics that Geosiphon is a member of the Glomeromycota made it a valuable experimental model system. From it, Schu¨ßler et al. (2006) characterised a fungal monosaccharide transporter, GpMST1, representing the type of sugar transporters in the AMF that perhaps are responsible for a globally significant carbon sink. Knowing its proper phylogenetic position also informs consideration of theories about land plant evolution (see below). As another example, there is a need to understand functional diversity and traits of different AMF taxa, but phylogenetic studies have shown, for example, that the genus Glomus was non-monophyletic, containing clades that diverged several hundreds of million years (MY) ago. It is impossible to predict functional aspects at, say, a generic level, if the comparisons are made among species so wrongly grouped. Although some such mis-placed organisms had been moved, for example, to Paraglomus (Morton and Redecker 2001), others remained and even if all the misplaced species were transferred, Glomus would still have represented several families and genera in a clade at the ordinal level. Taxon-related functional diversity in AMF is gradually being discovered, and a molecular phylogeny based ‘natural

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

systematics’ of the group was formally published recently (Schu¨ßler and Walker 2010), facilitating the recognition of such functional diversity. There are other important aspects that can be addressed through understanding the phylogeny of AMF. The biogeography of these symbionts is little understood at present. How and when did the AMF lineages evolve? How can it be that extant AMF species occur as symbionts with land plants worldwide? Do apparently identical morphospecies have very different genetic characteristics? Isolates of certain species occurring all over the globe seem genetically similar, but is this due to human dispersion as suggested by Rosendahl et al. (2009)? Certainly species found in more natural ecosystems also occur on different continents. Can it be true that panglobally distributed extant AMF morphospecies radiated before Pangaea split? How can this be related to land plant evolution? Were land plants indeed the first symbiotic partners of these fungi, or were AMF precursors already on the ‘symbiotic track’ before land plants evolved? We cannot yet answer many of these questions, but we have some indications from molecular data and the fossil record for the evolution of the Glomeromycota, and their co-evolution with land plants (and perhaps other organisms).

B. Phylogenies and a ‘Natural’ Systematic and Taxonomy In general, modern fungal systematics is based largely on phylogenetic data (e.g., Hibbett et al. 2007). The first work on rRNA gene phylogeny of AMF was that of Simon et al. (1993) followed by the data on Geosiphon pyriformis (Gehrig et al. 1996) and the deep phylogenetic roots of AMF were shown. At that time SSU rDNA sequences of only 14 AMF were known. Naming of species and higher taxa since then has often included molecular evidence (Kramadibrata et al. 2000; Morton and Redecker 2001). However, some recently erected taxa were not based on sound evidence and complicated the group’s systematics. A large set of such new taxa eventually was rejected (see for example, Morton and Msiska 2010). Currently, 11 families of comparable phylogenetic rank are described. The complete or near complete SSU rRNA gene

167

sequence data still have the best taxonomic coverage and resolve the Glomeromycota well from the level of order to genus, and it was used as the foundation of the current systematics of AMF, summarised in Fig. 7.1 (Schu¨ßler and Walker 2010). These newly established taxa will be used in this review. Mainly based on the SSU rRNA gene phylogenies a new familial (Schwarzott et al. 2001) and ordinal ranking was established (Schu¨ßler et al. 2001; Walker and Schu¨ßler 2004) to resolve some conflicts of molecular phylogenetic relationships with the previous systematics. The incorporation of rDNA sequence data as characters in the formal description of new taxa was initiated (Morton and Redecker 2001; Schu¨ßler et al. 2001), and with some exceptions the new classification was well supported by nuclear rDNA sequence analyses. However, over-interpretation of molecular data also resulted in the erection of poorly supported taxa (Oehl et al. 2008). This study, splitting one genus from an evolutionary young family (the Gigasporaceae) into several often monogeneric families, was heavily debated and currently only one of the newly proposed taxa, the genus Racocetra, is accepted (Morton and Msiska 2010). However, at least one of the other genera proposed by Oehl et al. (2008) will most likely be resurrected (Kru¨ger et al. 2011). The renaming of many species in the Diversisporaceae that were artificially placed in Glomus was undertaken recently (Schu¨ßler and Walker 2010; Schu¨ßler et al. 2011) and the two monophyletic main Glomus-clades (Glomus Group A and B) in the Glomerales were each given the rank of family (Glomeraceae and Claroideoglomeraceae). A new species recently described on morphological grounds as Ambispora brasiliensis (Goto et al. 2008) belongs to Acaulospora (Kru¨ger et al. 2011). Evolutionarily, members of these distinct genera most likely separated >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

168 ORDER

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

169

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

170

Arthur Schu¨ßler and Christopher Walker

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).

171

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

600 800

500 600

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

400

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

172

Arthur Schu¨ßler and Christopher Walker

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.

173

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

174

Arthur Schu¨ßler and Christopher Walker

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,

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

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

175

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

176

Arthur Schu¨ßler and Christopher Walker

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

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

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

177

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

178

Arthur Schu¨ßler and Christopher Walker

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

179

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.

180

Arthur Schu¨ßler and Christopher Walker

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.

References Agerer R, Amirati J, Blanz P, Courtecuisse R, Desjardin DE, Gams W, Hallenberg N, Halling R, Hawksworth DL, Horak E, Korf RP, Mueller GM, Oberwinkler F, Rambold G, Summerbell RC, Triebel D, Watling R (2000) Open letter to the scientific community of mycologists. Can J Bot 78:981–983

Angelard C, Colard A, Niculita-Hirzel H, Croll D, Sanders IR (2010) Segregation in a mycorrhizal fungus alters rice growth and symbiosis-specific gene transcription. Curr Biol 20:1216–1221 Benedetto A, Bonfante P (2004) 2004 snapshots of AM fungi: still an endless tale. Mycol Res 108:338–340 Benjamin RK (1979) Zygomycetes and their spores. In: Kendrick B (ed) The whole fungus, vol 2. National Museum of Natural Sciences and National Museum of Canada, Ottawa, pp 573–622 Benny GL, Humber RA, Morton JB (2001) Zygomycota: Zygomycetes. In: McLaughlin DJ, McLaughlin EG, Lemke PA (eds) The Mycota, vol 7A. Systematics and evolution. Springer, Berlin Heidelberg New York, pp 113–146 Berbee ML, Taylor JW (2001) Fungal molecular evolution: gene trees and geologic time. In: McLaughlin DJ, McLaughlin EG, Lemke PA (eds) The Mycota, vol 7B. Systematics and evolution. Springer, Berlin Heidelberg New York, pp 229–245 Bidartondo MI, et al. (2008) Preserving Accuracy in Genbank, Science 21:1616 Bo¨rstler B, Thie´ry O, Sy´korov Z, Berner A, Redecker D (2010) Diversity of mitochondrial large subunit rDNA haplotypes of Glomus intraradices in two agricultural field experiments and two semi-natural grasslands. Mol Ecol 19:1497–1511 Bucholtz F (1912) Beitra¨ge zur Kenntnis der Gattung Endogone Link. Beih Bot Zbl 29:147–225 Cai CY, Ouyang S, Wang Y, Fang ZJ, Rong JY, Geng LY, Li XX (1996) An Early Silurian vascular plant. Nature 379:592 Castagnone-Sereno P (2006) Genetic variability and adaptive evolution in parthenogenetic root-knot nematodes. Heredity 96:282–289 Check-Hayden E (2008) Evolution: Scandal! Sex-starved and still surviving. Nature 452:678–680 Corradi N, Kuhn G, Sanders IR (2004a) Monophyly of b-tubulin and H+ATPase gene variants in Glomus intraradices: consequences for molecular evolutionary studies of AM fungal genes. Fungal Genet Biol 41:262–273 Corradi N, Hijri M, Fumagalli L, Sanders IR (2004b) Arbuscular mycorrhizal fungi (Glomeromycota) harbour ancient fungal tubulin genes that resemble those of the chytrids (Chytridiomycota). Fungal Genet Biol 41:1037–1045 Corradi N, Sanders IR (2006) Evolution of the P-type II ATPase gene family in the fungi and presence of structural genomic changes among isolates of Glomus intraradices. BMC Evol Biol 6:21 Couch BC, Fudal I, Lebrun MH, Tharreau D, Valent B, Kim PV, Notte´ghem JL, Kohn LM (2005) Origins of hostspecific populations of the blast pathogen, Magnaporthe oryzae, in crop domestication with subsequent expansion of pandemic clones on rice and weeds of rice. Genetics 170:613–630 Coyne JA, Orr HA (1998) The evolutionary genetics of speciation. Phil Trans R Soc B:287–305 Croll D, Wille L, Gamper HA, Mathimaran N, Lammers PJ, Corradi N, Sanders IR (2008) Genetic diversity and host plant preferences revealed by simple sequence

Evolution of the ‘Plant-Symbiotic’ Fungal Phylum, Glomeromycota repeat and mitochondrial markers in a population of the arbuscular mycorrhizal fungus Glomus intraradices. New Phytol 178:672–687 Dangeard PA (1900) Le Rhizophagus populinus. Botaniste 7:285–287 de Souza FA, Leeflang P, Kowalchuk GA, van Veen JA, Smit E (2004) PCR-denaturing gradient gel electrophoresis profiling of the inter- and intraspecies 18S rRNA gene sequence heterogeneity is an accurate and sensitive method to assess species diversity of arbuscular mycorrhizal fungi of the genus Gigaspora. Appl Environ Microbiol 70:1413–1424 Dotzler N, Krings M, Hass H, Walker C, Taylor TN, Agerer R (2009) Acaulosporoid glomeromycotan spores with germination shields from 400-millionyear-old Rhynie chert. Mycol Progr 8:9–18 Dotzler N, Krings M, Taylor TN, Agerer R (2006) Germination shields in Scutellospora (Glomeromycota: Diversisporales, Gigasporaceae) from the 400 million-year-old Rhynie chert. Mycol Progr 5:178–184 Douhan GW, Petersen C, Bledsoe CS, Rizzo DM (2005) Contrasting root associated fungi of three common oak-woodland plant species based on molecular identification: host specificity or non-specific amplification? Mycorrhiza 15:365–372 Ekelund F, Rønn R (2008) If you don’t need change, maybe you don’t need sex. Nature 453:587. Ferrol N, Barea JM, Azcon-Aguilar C (2000) The plasma membrane H+ATPase gene family in the arbuscular mycorrhizal fungus Glomus mosseae. Curr Genet 37: 112–118 Fitter AH (2005) Darkness visible: reflections on underground ecology. J Ecol 93:231–243 Fontaneto DE, Herniou EA, Boschetti C, Caprioli M, Melone G, Ricci C, Barraclough TG (2007) Independently evolving species in asexual bdelloid rotifers. PLoS Biol 5:e87 ¨ ber die auf Wurzelsymbiose beruhende Frank AB (1885) U Erna¨hrung gewisser Ba¨ume durch unterirdische Pilze. Ber Dtsch Bot Ges 3:128–145 Franke M, Morton JB (1994) Ontogenetic comparisons of arbuscular mycorrhizal fungi Scutellospora heterogama and Scutellospora pellucida: revision of taxonomic character concepts, species descriptions, and phylogenetic hypotheses. Can J Bot 72: 122–134 Gallaud I (1905) Etudes sur les mycorrhizes endotrophes. Rev Gen Bot 17:5–48, 66–83, 123–135, 223–239, 313–325, 425–433, 479–500 Gamper H, Walker C, Schu¨ßler A (2009) Diversispora celata sp. nov: molecular ecology and phylotaxonomy of an inconspicuous arbuscular mycorrhizal fungus. New Phytol 182:495–506 Gehrig H, Schu¨ßler A, Kluge M (1996) Geosiphon pyriformis, a fungus forming endocytobiosis with Nostoc (Cyanobacteria), is an ancestral member of the Glomerales: evidence by SSU rRNA analysis. J Mol Evol 43:71–81 Gerdemann JW (1968) Vesicular–arbuscular mycorrhiza and plant growth. Annu Rev Phytopathol 6:397–418 Gerdemann JW, Trappe JM (1974) Endogonaceae in the Pacific Northwest. Mycol Mem 5:1–76

181

Giovannetti M, Sbrana C, Strani P, Agnolucci M, Rinaudo V, Avio L (2003) Genetic diversity of isolates of Glomus mosseae from different geographic areas detected by vegetative compatibility testing and biochemical and molecular analysis. Appl Environ Microbiol 69:616–624 Goto BT, Maia LC, Oehl F (2008) Ambispora brasiliensis a new ornamented species in the arbuscular mycorrhiza-forming Glomeromycetes. Mycotaxon 105:11–18 Hamilton WD (2001) Narrow roads of gene land, vol 2. Evolution of sex. Oxford University Press, Oxford Heckman DS, Geiser DM, Eidell BR, Stauffer RL, Kardos NL, Hedges SB (2001) Molecular evidence for the early colonization of land by fungi and plants. Science 293:1129–1133 Hedges SB, Kumar S (2003) Genomic clocks and evolutionary timescales. Trends Genet 19:200–206 Helgason T, Merryweather JW, Denison J, Wilson P, Young JPW, Fitter A (2002) Selectivity and functional diversity in arbuscular mycorrhizas of co-occurring fungi and plants from a temperate deciduous woodland. J Ecol 90:371–384 Helgason T, Watson IJ, Young JPW (2003) Phylogeny of the Glomerales and Diversisporales (Fungi: Glomeromycota) from actin and elongation factor 1-alpha sequences. FEMS Microbiol Lett 229:127–132 Hibbett DS, Binder M, Bischoff JF, Blackwell M, Cannon PF, Eriksson OE, Huhndorf S, James T, Kirk PM, Lu¨cking R, Lumbsch HT, Lutzoni F, Matheny PB, McLaughlin DJ, Powell MJ, Redhead S, Schoch CL, Spatafora JW, Stalpers JA, Vilgalys R, Aime MC, Aptroot A, Bauer R, Begerow D, Benny GL, Castlebury LA, Crous PW, Dai YC, Gams W, Geiser DM, Griffith GW, Gueidan C, Hawksworth DL, Hestmark G, Hosaka K, Humber RA, Hyde KD, Ironside JE, Ko˜ljalg U, Kurtzman CP, Larsson KH, Lichtwardt R, Longcore J, Mia˛dlikowska J, Miller A, Moncalvo JM, MozleyStandridge S, Oberwinkler F, Parmasto E, Reeb V, Rogers JD, Roux C, Ryvarden L, Sampaio JP, Schu¨ßler A, Sugiyama J, Thorn RG, Tibell L, Untereiner WA, Walker C, Wang Z, Weir A, Weiss M, White MM, Winka K, Yao YJ, Zhang N (2007) A higher-level phylogenetic classification of the Fungi. Mycol Res 111:509–547 Hijri M, Sanders IR (2005) Low gene copy number shows that arbuscular mycorrhizal fungi inherit genetically different nuclei. Nature 433:160–163 Husband R, Herre EA, Young JP (2002) Temporal variation in the arbuscular mycorrhizal communities colonising seedlings in a tropical forest. FEMS Microbiol Ecol 42:131–136 Ijdo M, Schtickzelle N, Cranenbrouck S, Declerck S (2010) Do arbuscular mycorrhizal fungi with contrasting life-history strategies differ in their responses to repeated defoliation? FEMS Microbiol Ecol 72:114–122 James TY, Kauff F, Schoch CL, Matheny PB, Hofstetter V, Cox C, Celio G, Gueidan C, Fraker E, Mia˛dlikowska J, Lumbsch, HT, Rauhut A, Reeb V, Arnold EA, Amtoft A, Stajich JE, Hosaka K, Sung GH, Johnson D, O’Rourke B, Crockett M, Binder M, Curtis JM, Slot JC, Wang Z, Wilson AW, Schu¨ßler A, Longcore JE,

182

Arthur Schu¨ßler and Christopher Walker

O’Donnell K, Mozley-Standridge S, Porter D, Letcher PM, Powell MJ, Taylor JW, White MM, Griffith GW, Davies DR, Humber RA, Morton J, Sugiyama J, Rossman AY, Rogers, JD, Pfister DH, Hewitt D, Hansen K, Hambleton S, Shoemaker RA, Kohlmeyer J, Volkmann-Kohlmeyer B, Spotts RA, Serdani M, Crous PW, Hughes KW, Matsuura K, Langer E, Langer G, Untereiner WA, Lu¨cking R, Bu¨del B, Geiser DM, Aptroot A, Diederich P, Schmitt I, Schultz M, Yahr R, Hibbett DS, Lutzoni F, McLaughlin D, Spatafora J, Vilgalys R (2006) Reconstructing the early evolution of the fungi using a six gene phylogeny. Nature 443: 818–822 Jansa J, Mozafar A, Banke S, McDonald BA, Frossard E (2002) Intra- and intersporal diversity of ITS rDNA sequences in Glomus intraradices assessed by cloning and sequencing, and by SSCP analysis. Mycol Res 106: 670–681 Jastrow JD, Miller RM (1997) Soil aggregate stabilization and carbon sequestration: feedbacks through organomineral associations. In: Lal R, Kimble J, Follet R, Stewart B (eds) Soil processes and the carbon cycle. CRC, Boca Raton Johnson D, Vandenkoornhuyse PJ, Leake J, Gilbert L, Booth RE, Grime JP, Young JPW, Read D (2003) Plant communities affect arbuscular mycorrhizal fungal diversity and community composition in grassland microcosms. New Phytol 161:503–515 Keeling PJ (2003) Congruent evidence from a-tubulin and b-tubulin gene phylogenies for a zygomycete origin of microsporidia. Fungal Genet Biol 38:298–309 Kjøller R, Rosendahl S (2000) Detection of arbuscular mycorrhizal fungi (Glomales) in roots by nested PCR and SSCP (single stranded conformation polymorphism). Plant Soil 226:189–196 Kondrashov AS (1988) Deleterious mutations and the evolution of sexual reproduction. Nature 336:435–440 Kottke I, Haug I, Setaro S, Surez JP, Weiß M, Preußing M, Nebel M, Oberwinkler F (2008) Guilds of mycorrhizal fungi and their relation to trees, ericads, orchids and liverworts in a neotropical mountain rain forest. Basic Appl Ecol 9:13–23 Kramadibrata K, Walker C, Schwarzott D, Schu¨ßler A (2000) A new species of Scutellospora with a coiled germination shield. Ann Bot 86:21–27 Kru¨ger M, Kru¨ger C, Walker C, Stockinger H, Schu¨ßler A (2011) A phylogenetic framework for the natural systematics of arbuscular mycorrhizal fungi - from phylum to species-level resolution and environmental deep sequencing. New Phytol: in revision Kru¨ger M, Stockinger H, Kru¨ger C, Schu¨ßler A (2009) DNA-based species level detection of Glomeromycota: one PCR primer set for all arbuscular mycorrhizal fungi. New Phytol 183:212–223 Kru¨ger M, Walker C, Schu¨ßler A (2011) Acaulospora brasiliensis comb. nov. and Acaulospora alpina (Glomeromycota) from upland Scotland: morphology, molecular phylogeny and DNA based detection in roots. Mycorrhiza. doi: 10.1007/s00572-011-0361-7 Kuga Y, Saito K, Nayuki K, Peterson RL, Saito M (2008) Ultrastructure of rapidly frozen and freezesubstituted germ tubes of an arbuscular mycorrhizal

fungus and localization of polyphosphate. New Phytol 178:189–200 Lanfranco L, Delpero M, Bonfante P (1999) Intrasporal variability of ribosomal sequences in the endomycorrhizal fungus Gigaspora margarita. Mol Ecol 8:37–45 Lee J, Young JP (2009) The mitochondrial genome sequence of the arbuscular mycorrhizal fungus Glomus intraradices isolate 494 and implications for the phylogenetic placement of Glomus. New Phytol 183: 200–211 Lekberg Y, Koide RT, Rohr JR, Aldrich-Wolfe L, Morton JB (2007) Role of niche restrictions and dispersal in the composition of arbuscular mycorrhizal fungal communities. J Ecol 95:95–105 LoBuglio KF, Pitt JI, Taylor JW (1993) Phylogenetic analysis of two ribosomal DNA regions indicates multiple independent losses of a sexual Talaromyces state among asexual Penicillium species in subgenus Biverticillium. Mycologia 85:592–604 Lumini E, Bianciotto V, Jargeat P, Novero M, Salvioli A, Faccio A, Be´card G, Bonfante P (2007) Presymbiotic growth and sporal morphology are affected in the arbuscular mycorrhizal fungus Gigaspora margarita cured of its endobacteria. Cell Microbiol 9: 1716–1729 Maherali H, Klironomos JN (2007) Influence of phylogeny on fungal community assembly and ecosystem functioning. Science 316:1746–1748 Mathimaran N, Falquet L, Ineichen K, Picard C, Redecker D, Boller T, Wiemken A (2008) Microsatellites for disentangling underground networks: strain-specific identification of Glomus intraradices, an arbuscular mycorrhizal fungus. Fungal Genet Biol 45:812–817 Mayden RL (1997) A hierarchy of species concepts: The denouement in the saga of the species problem. In: Claridge MF, Dawah HA, Wilson MR (eds) Species: the units of biodiversity. Chapman and Hall, New York, pp 381–424 Mercereau-Puijalon O, Barale JC, Bischoff E (2002) Three multigene families in Plasmodium parasites: facts and questions. Int J Parasitol 32:1323–1344 Morton JB (2000) Evolution of endophytism in arbuscular mycorrhizal fungi of Glomales. In: Bacon CW, White JH (eds) Microbial endophytes. Dekker, New York, pp 121–140 Morton JB, Benny GL (1990) Revised classification of arbuscular mycorrhizal fungi (Zygomycetes): a new order, Glomales, two new suborders, Glomineae and Gigasporineae, and two new families, Acaulosporacea and Gigasporaceae, with an emendation of Glomaceae. Mycotaxon 37:471–491 Morton JB, Redecker D (2001) Two new families of Glomales, Archaeosporaceae and Paraglomaceae, with two new genera, Archaeospora and Paraglomus, based on concordant molecular and morphological characters. Mycologia 93:181–195 Morton JB, Msiska Z (2010) Phylogenies from genetic and morphological characters do not support a revision of Gigasporaceae (Glomeromycota) into four families and five genera. Mycorrhiza 20:483–496 Mosse B (1953) Fructifications associated with mycorrhizal strawberry roots. Nature 171:974

Evolution of the ‘Plant-Symbiotic’ Fungal Phylum, Glomeromycota Mosse B, Bowen GD (1968) A key to the recognition of some Endogone spore types. Trans Br Mycol Soc 51: 469–483 Munkvold L, Kjøller R, Vestberg M, Rosendahl S, Jakobsen I (2004) High functional diversity within species of arbuscular mycorrhizal fungi. New Phytol 164:357–364 Naumann M, Schu¨ßler A, Bonfante P (2010) The obligate endobacteria of arbuscular mycorrhizal fungi are ancient heritable components related to the Mollicutes. ISME J 4:862–871 Nicolson TH, Gerdemann JW (1968) Mycorrhizal Endogone species. Mycologia 60:313–325 Nielsen KB, Kjøller R, Olsson PA, Schweiger PF, Andersen FØ, Rosendahl S (2004) Colonization intensity and molecular diversity of arbuscular mycorrhizal fungi in the aquatic plants Littorella uniflora and Lobelia dortmanna in Southern Sweden. Mycol Res 108:616–625 Normark BB, Judson OP, Moran NA (2003) Genomic signatures of ancient asexual lineages. Biol J Linn Soc 79: 69–84 O’Donnell K, Lutzoni F, Ward TJ, Benny GL (2001) Evolutionary relationships among mucoralean fungi (Zygomycota): evidence for family polyphyly on a large scale. Mycologia 93:286–297 ¨ pik M, Moora M, Zobel M, Saks U ¨ , Wheatley R, Wright F, O Daniell T (2008) High diversity of arbuscular mycorrhizal fungi in a boreal herb-rich coniferous forest. New Phytol 179:867–876 Parniske M (2008) Arbuscular mycorrhiza: the mother of plant root endosymbioses. Nature Rev Microbiol 6: 763–776 Pawlowska TE, Taylor JW (2004) Organization of genetic variation in individuals of arbuscular mycorrhizal fungi. Nature 427:733–737 Peyronel B (1937) Le “Endogyne” quail produttrici di micorrize endotrofische nell fanerogame alpestri. Nuovo Gior Bot Ital N S 44:584–586 Phipps CJ, Taylor TN (1996) Mixed arbuscular-mycorrhizae from the Triassic of Antarctica. Mycologia 88: 707–714 Pirozynski KA, Dalpe´ Y (1989) Geological history of the Glomaceae with particular reference to mycorrhizal symbiosis. Symbiosis 7:1–36 Pirozynski KA, Malloch DW (1975) The origin of land plants: a matter of mycotrophism. BioSystems 6: 153–164 Poxleitner MK, Carpenter ML, Mancuso JJ, Wang CJR, Dawson SC, Cande WZ (2008) Evidence for karyogamy and exchange of genetic material in the binucleate intestinal parasite Giardia intestinalis. Science 319:1530–1533 Pozo MJ, Azcon-Aguilar C (2007) Unraveling mycorrhizainduced resistance. Curr Opin Plant Biol 10:393–398 Redecker D (2002) Molecular identification and phylogeny of arbuscular mycorrhizal fungi. Plant Soil 244:67–73 Redecker D, Hijri M, Dulieu H, Sanders IR (1999) Phylogenetic analysis of a dataset of fungal 5.8S rDNA sequences shows that highly divergent copies of Internal Transcribed Spacers reported from Scutellospora castanea are of Ascomycete origin. Fungal Genet Biol 28:238–244

183

Redecker D, Kodner R, Graham LE (2000a) Glomalean fungi from the Ordovician. Science 289:1920–1921 Redecker D, Morton JB, Bruns TD (2000b) Molecular phylogeny of the arbuscular mycorrhizal fungi Glomus sinuosum and Sclerocystis coremioides. Mycologia 92: 282–285 Redecker D, Raab P, Oehl F, Camacho FJ, Courtecuisse R (2007) A novel clade of sporocarp-forming species of glomeromycotan fungi in the Diversisporales lineage. Mycol Progress 6:35–44 Redon R, Ishikawa S, Fitch KR, Feuk L, Perry GH, Andrews TD, Fiegler H, Shapero MH, Carson AR, Chen W, Cho EK, Dallaire S, Freeman JL, Gonzalez JR, Gratacos M, Huang J, Kalaitzopoulos D, Komura D, MacDonald JR, Marshall CR, Mei R, Montgomery L, Nishimura K, Okamura K, Shen F, Somerville MJ, Tchinda J, Valsesia A, Woodwark C, Yang F, Zhang J, Zerjal T, Zhang J, Armengol L, Conrad DF, Estivill X, TylerSmith C, Carter NP, Aburatani H, Lee C, Jones KW, Scherer SW, Hurles ME (2006) Global variation in copy number in the human genome. Nature 444: 444–454 Remy W, Taylor TN, Hass H, Kerp H (1994) Four-hundredmillion-year-old vesicular arbuscular mycorrhizae. Proc Natl Acad Sci USA 91:11841–11843 Requena N, Breuninger M, Franken P, Oco´n A (2003) Symbiotic status, phosphate, and sucrose regulate the expression of two plasma membrane H+ATPase genes from the mycorrhizal fungus Glomus mosseae. Plant Physiol 132:1540–1549 Rillig MC (2004) Arbuscular mycorrhizae and terrestrial ecosystem processes. Ecol Lett 7:740–754 Rosendahl S (2008) Communities, populations and individuals of arbuscular mycorrhizal fungi. New Phytol 178:253–266 Rosendahl S, McGee P, Morton JB. (2009) Lack of global population genetic differentiation in the arbuscular mycorrhizal fungus Glomus mosseae suggests a recent range expansion which may have coincided with the spread of agriculture. Mol Ecol 18:4316–4329 Rubinstein CV, Gerrienne P, De La Puente GS, Astini RA, Steemans P (2010) Early Middle Ordovician evidence for land plants in Argentina (eastern Gondwana). New Phytol 188:365–369 Scheublin TR, Ridgway KP, Young JPW, van der Heijden MGA (2004) Nonlegumes, legumes, and root nodules harbor different arbuscular mycorrhizal fungal communities. Appl Environ Microbiol 70:6240–6246 Schu¨ßler A (1999) Glomales SSU rRNA gene diversity. New Phytol 144:205–207 Schu¨ßler A (2000) Glomus claroideum forms an arbuscular mycorrhiza-like symbiosis with the hornwort Anthoceros punctatus. Mycorrhiza 10:15–21 Schu¨ßler A (2002) Molecular phylogeny, taxonomy, and evolution of Geosiphon pyriformis and arbuscular mycorrhizal fungi. Plant Soil 244:75–83 Schu¨ßler A, Kluge M (2001) Geosiphon pyriforme, an endocytosymbiosis between fungus and cyanobacteria, and its meaning as a model system for arbuscular mycorrhizal research. In: Hock B (ed) The Mycota, vol 9. Fungal associations. Springer, Berlin Heidelberg New York, pp 151–161

184

Arthur Schu¨ßler and Christopher Walker

Schu¨ßler A, Wolf E (2005) Geosiphon pyriformis – a glomeromycotan soil fungus forming endosymbiosis with cyanobacteria. In: Declerck S, Strullu DG, Fortin JA (eds) In vitro culture of mycorrhizas, Soil biology, vol 4. Springer, Berlin Heidelberg New York, pp 271–289 Schu¨ßler A, Mollenhauer D, Schnepf E, Kluge M (1994) Geosiphon pyriforme, an endosymbiotic association of fungus and cyanobacteria: the spore structure resembles that of arbuscular mycorrhizal (AM) fungi. Botanica Acta 107:36–45 Schu¨ßler A, Schwarzott D, Walker C (2001) A new fungal phylum, the Glomeromycota: phylogeny and evolution. Mycol Res 105:1413–1421 Schu¨ßler A, Martin H, Cohen D, Fitz M, Wipf D (2006) Characterization of a carbohydrate transporter from symbiotic glomeromycotan fungi. Nature 444:933–936 Schu¨ßler A, Martin H, Cohen D, Wipf D (2008) The Geosiphon–Nostoc symbiosis as a tool to characterize symbiotic nutrient transporters in the arbuscular mycorrhiza symbiosis. Biol Mol Plant Microbe Interact 6:1–6 Schu¨ßler A, Walker C (2010) Arbuscular Mycorrhizal Fungi: placing an experimental model fungus in its natural systematic relationship - culture BEG47 is Diversispora epigaea, not Glomus versiforme. PLoS ONE: in revision Schu¨ßler A, Walker C (2010) The Glomeromycota: a species list with new families and genera. Arthur Schu¨ßler & Christopher Walker, Gloucester. Published in December 2010 in libraries at The Royal Botanic Garden Edinburgh, The Royal Botanic Garden Kew, Botanische Staatssammlung Munich, and Oregon State University. Electronic version freely available online at www.amf-phylogeny.com Schwarzott D, Walker C, Schu¨ßler A (2001) Glomus, the largest genus of the arbuscular mycorrhizal fungi (Glomales), is non-monophyletic. Mol Phylogenet Evol 21:190–197 Sieverding E, Oehl F (2006) Revision of Entrophospora and description of Kuklospora and Intraspora, two new genera in the arbuscular mycorrhizal Glomeromycetes. J Appl Bot Food Qual 80:69–81 Simon L, Bousquet J, Le!vesque RC, Lalonde M (1993) Origin and diversification of endomycorrhizal fungi and coincidence with vascular land plants. Nature 363:67–69 Smith SA, Beaulieu JM, Donoghue MJ (2010) An uncorrelated relaxed-clock analysis suggests an earlier origin for flowering plants. Proc Nat Acad Sci 107:5897–5902 Smith SE, Read DJ (2008) Mycorrhizal symbiosis, 3rd edn. Academic Press, London Smith SE, Smith FA, Jakobsen I (2004) Functional diversity in arbuscular mycorrhizal (AM) symbioses: the contribution of the mycorrhizal P uptake pathway is not correlated with mycorrhizal responses in growth or total P uptake. New Phytol 162:511–524 Stewart WN, Rothwell GW (1993) Paleobotany and the evolution of plants. Cambridge Univ Press, Cambridge Stockinger H, Walker C, Schu¨ßler A (2009) Glomus intraradices DAOM197198’, a model fungus in arbuscular

mycorrhiza research, is not Glomus intraradices. New Phytol 183:1176–1187 Stockinger H, Kru¨ger M, Schu¨ßler A (2010) DNA barcoding of arbuscular mycorrhizal fungi. New Phytol 187: 461–474 Strullu-Derrien C, Strullu DG (2007) Mycorrhization of fossil and living plants. Crit Rev Paleoevol 6:483–494 Stubblefield SP, Taylor TN, Trappe JM (1987) Fossil mycorrhizae: a case for symbiosis. Science 237:59–60 Stukenbrock EH, Rosendahl S (2005) Clonal diversity and population genetic structure of arbuscular mycorrhizal fungi (Glomus spp) studied by multilocus genotyping of single spores. Mol Ecol 14:743–752 Taga M, Tsuchiya D, Murata M (2003) Dynamic changes of rDNA condensation state during mitosis in filamentous fungi revealed by fluorescence in situ hybridisation. Mycol Res 107:1012–1020 Tanabe Y, Saikawa M, Watanabe MM, Sugiyama J (2003) Molecular phylogeny of Zygomycota based on EF-1a and RPB1 sequences: limitations and utility of alternative markers to rDNA. Mol Phylogen Evol 30: 438–449 Taylor JW, Berbee ML (2006) Dating divergences in the fungal tree of life: review and new analyses. Mycologia 98:838–849 Taylor JW, Jacobson DJ, Kroken S, Kasuga T, Geiser DM (2000) Phylogenetic species recognition and species concepts in fungi. Fungal Genet Biol 31:21–32 Thaxter R (1922) A revision of the Endogoneae. Proc Am Acad Arts Sci 57:291–351 Trouvelot S, van Tuinen D, Hijri M, Gianinazzi-Pearson V (1999) Visualization of ribosomal DNA loci in spore interphasic nuclei of glomalean fungi by fluorescence in situ hybridization. Mycorrhiza 8:203–206 Tulasne LR, Tulasne C (1845) Fungi nonnulli hypogaei, novi v. minus cogniti act. Giorn Bot Ital 2:35–63 Tulasne LR, Tulasne C (1851) Fungi hypogaei, 1st edn. Paris van der Heijden MGA, Klironomos JN, Ursic M, Moutoglis P, Streitwolf-Engel R, Boller T, Wiemken A, Sanders IR (1998) Mycorrhizal fungal diversity determines plant biodiversity, ecosystem variability and productivity. Nature 396:69–72 van Tuinen D, Jacquot E, Zhao B, Gollotte A, GianinazziPearson V (1998) Characterization of root colonization profiles by a microcosm community of arbuscular mycorrhizal fungi using 25S rDNA-targeted nested PCR. Mol Ecol 7:879–887 Vandenkoornhuyse P, Husband R, Daniell TJ, Watson IJ, Duck JM, Fitter AH, Young JPW (2002) Arbuscular mycorrhizal community composition associated with two plant species in a grassland ecosystem. Mol Ecol 11:1555–1564 Vandenkoornhuyse P, Ridgway KP, Watson IJ, Fitter AH, Young JPW (2003) Coexisting grass species have distinctive arbuscular mycorrhizal communities. Mol Ecol 12:3085–3095 Wang B, Yeun LH, Xue JY, Liu Y, Ane´ JM, Qiu YL (2010) Presence of three mycorrhizal genes in the common ancestors of land plants suggests a key role of mycorrhizas in the colonization of land plants. New Phytol 186:514–525

Evolution of the ‘Plant-Symbiotic’ Fungal Phylum, Glomeromycota Walker C, Sanders FE (1986) Taxonomic concepts in the Endogonaceae: III. The separation of Scutellospora gen. nov. from Gigaspora Gerd. and Trappe. Mycotaxon 27:169–182 Walker C, Schu¨ßler A (2004) Nomenclatural clarifications and new taxa in the Glomeromycota. Mycol Res 108: 981–982 Walker C, Vestberg M, Demircik F, Stockinger H, Saito M, Sawaki H, Nishmura I, Schu¨ßler A (2007) Molecular phylogeny and new taxa in the Archaeosporales (Glomeromycota): Ambispora fennica gen. sp nov., Ambisporaceae fam. nov., and emendation of Archaeospora and Archaeosporaceae. Mycol Res 111:137–153

185

Wellman CH, Osterloff PL, Mohluddin U (2003) Fragments of the earliest land plants. Nature 425: 282–285 Yadav, V, Kumar, M, Deep, DK, Kumar, H, Sharma, R, Tripathi, T, Tuteja, N, Saxena, AK, Johri, AK (2010). A phosphate transporter from the root endophytic fungus Piriformospora indica plays a role in the phosphate transport to the host plant. J Biol Chem 285: 26532–26544 Zhang, Q, Blaylock, LA, Harrison, MJ (2010) Two Medicago truncatula half-ABC transporters are essential for arbuscule development in arbuscular mycorrhizal symbiosis. Plant Cell 22:1483–1497

.

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

188

Imke Schmitt

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

Fruiting Body Evolution in the Ascomycota: a Molecular Perspective

189

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

190

Imke Schmitt

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

Fruiting Body Evolution in the Ascomycota: a Molecular Perspective

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

191

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

192

Imke Schmitt

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

Fruiting Body Evolution in the Ascomycota: a Molecular Perspective

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.

193

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

194

Imke Schmitt

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

Fruiting Body Evolution in the Ascomycota: a Molecular Perspective

195

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

196

Imke Schmitt

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

Fruiting Body Evolution in the Ascomycota: a Molecular Perspective

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,

197

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)

198

Imke Schmitt

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).

Fruiting Body Evolution in the Ascomycota: a Molecular Perspective

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-

199

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).

References Baral HO (1992) Vital versus herbarium taxonomy: morphological differences between living and dead cells of ascomycetes, and their taxonomic implications. Mycotaxon 44:333–390 Belleme`re A (1967) Contributions a` l’e´tude du de´veloppement de l’pothe´cie chez les Discomyce`tes inopercule´s. Bull Soc Mycol France 83:393–640, 753–931 Belleme`re A (1977) Apical apparatus of some Discomycetes-asci – ultrastructural study. Rev Mycol 41: 233–264 Belleme`re A, Letrouit-Galinou MA (1987) Differentiation of lichen asci including dehiscence and sporogenesis: an ultrastructural survey. Bibl Lichenol 25:137–161 Berbee ML (1996) Loculoascomycete origins and evolution of filamentous ascomycete morphology based on 18S rRNA gene sequence data. Mol Biol Evol 13:462–470 Berbee ML, Carmean DA, Winka K (2000) Ribosomal DNA and resolution of branching order among the ascomycota: How many nucleotides are enough? Mol Phylogenet Evol 17:337–344 Berbee ML, Taylor JW (1992a) Convergence in ascospore discharge mechanism among pyrenomycete fungi based on 18S ribosomal RNA gene sequence. Mol Phylogenet Evol 1:59–71 Berbee ML, Taylor JW (1992b) Detecting morphological convergence in true fungi, using-18s Ribosomal-Rna Gene Sequence Data. Biosystems 28:117–125 Berbee ML, Taylor JW (1992c) Two ascomycete classes based on fruiting-body characters and ribosomal DNA-sequence. Mol Biol Evol 9:278–284 Berbee ML, Taylor JW (1995) From 18S ribosomal sequence data to evolution of morphology among the Fungi. Can J Bot 73:S677–S683

200

Imke Schmitt

Binder M, Hibbett DS (2006) Molecular systematics and biological diversification of Boletales. Mycologia 98: 971–981 Bistis GN, Perkins DD, Read ND (2003) Different cell types in Neurospora crassa. Fungal Genet Biol 50:17–19 Blackwell M (1994) Minute mycological mysteries – the influence of arthropods on the lives of fungi. Mycologia 86:1–17 Bruns TD, Fogel R, White TJ, Palmer JD (1989) Accelerated evolution of a false-truffle from a mushroom ancestor. Nature 339:140–142 Busch S, Braus GH (2007) How to build a fungal fruit body: from uniform cells to specialized tissue. Mol Microbiol 64:873–876 Buschbom J, Mueller G (2004) Resolving evolutionary relationships in the lichen-forming genus Porpidia and related allies (Porpidiaceae, Ascomycota). Mol Phylogenet Evol 32:66–82 Cain RF (1972) Evolution of the fungi. Mycologia 64:1–14 Chadefaud M (1973) Les asques et la syste´matique des Ascomyce`tes. Bull Soc Mycol Fr 89:127–170 Chadefaud M, Letrouit-Galinou MA, Favre MC (1963) Sur l’e´volution des asques du type archaeasce chez les Discomyce`tes de l’ordre des Lecanorales. CR Acad Sci 257:4003–4005 Chadefaud M, Letrouit-Galinou MA, Janex-Favre MC (1968) Sur l’origine phylogenetique et l’e´volution des Ascomyce`tes des lichens. Bull Soc Bot France 115:79–111 Cunningham CW, Omland KE, Oakley TH (1998) Reconstructing ancestral character states: a critical reappraisal. Trends Ecol Evol 13:361–366 del Prado R, Schmitt I, Kautz S, Palice Z, Lu¨cking R, Lumbsch HT (2006) Molecular data place Trypetheliaceae in Dothideomycetes. Mycol Res 110:511–520 Doppelbaur AW (1960) Studien zur Anatomie und Entwicklungsgeschichte einiger endolithischen pyrenocarpen Flechten. Planta 53:246–292 Do¨ring H, Clerc P, Grube M, Wedin M (2000) Mycobiontspecific PCR primers for the amplification of nuclear ITS and LSU rDNA from lichenized ascomycetes. Lichenologist 32:200–204 Do¨ring H, Henssen A, Wedin M (1999) Ascoma development in Neophyllis melacarpa (Lecanorales, Ascomycota), with notes on the systematic position of the genus. Austral J Bot 47:783–794 Do¨ring H, Lumbsch HT (1998) Ascoma ontogeny: is this character set of any use in the systematics of lichenized ascomycetes? Lichenologist 30:489–500 Do¨ring H, Wedin M (2000) Homology assessment of the boundary tissue in fruiting bodies of the lichen family Sphaerophoraceae (Lecanorales, Ascomycota). Plant Biol 2:361–367 Ekman S, Andersen HL, Wedin M (2008) The limitations of ancestral state reconstruction and the evolution of the ascus in the Lecanorales (lichenized Ascomycota). Syst Biol 57:141–156 Engh I, Nowrousian M, Ku¨ck U (2007a) Regulation of melanin biosynthesis via the dihydroxynaphthalene pathway is dependent on sexual development in the ascomycete Sordaria macrospora. FEMS Microbiol Lett 275:62–70

Engh I, Wuertz C, Witzel-Schlomp K, Zhang HY, Hoff B, Nowrousian M, Rottensteiner H, Kuck U (2007b) The WW domain protein PRO40 is required for fungal fertility and associates with Woronin bodies. Eukaryot Cell 6:831–843 Eriksson OE, Hawksworth DL (1993) Outline of the ascomycetes – 1993. Syst Ascomycetum 12:51–257 Esser, K, Straub, J (1958) Genetische Untersuchungen an Sordaria macrospora Auersw., Kompensation und Induktion bei genbedingten Entwicklungsdefekten. Zeitschr Vererbungsl 89:729–746 Fries EM (1821) Systema mycologicum. A series of volumes, published 1821–1832. Berlinge, Lund Gargas A, DePriest PT (1996) A nomenclature for fungal PCR primers with examples from intron-containing SSU rDNA. Mycologia 88:745–748 Gargas A, Taylor JW (1995) Phylogeny of discomycetes and early radiations of the apothecial Ascomycotina inferred from SSU rDNA sequence data. Exp Mycol 19:7–15 Geiser DM, Gueidan C, Miadlikowska J, Lutzoni F, Kauff F, Hofstetter V, Fraker E, Schoch CL, Tibell L, Untereiner WA, Aptroot A (2006) Eurotiomycetes: Eurotiomycetidae and Chaetothyriomycetidae. Mycologia 98:1053–1064 Greif MD, Gibas CFC, Tsuneda A, Currah RS (2007) Ascoma development and phylogeny of an apothecioid dothideomycete, Catinella olivacea. Am J Bot 94: 1890–1899 Greif MD, Tsuneda A, Currah RS (2004) The peridial development and dehiscence mechanism of Cryptendoxyla hypophloia, a cleistothecial ascomycete isolated from the bodies of arthropods. Int J Plant Sci 165:957–964 Grube M (1998) Classification and phylogeny in the Arthoniales (lichenized ascomycetes). Bryologist 101:377–391 Grube M, Baloch E, Lumbsch HT (2004) The phylogeny of Porinaceae (Ostropomycetidae) suggests a neotenic origin of perithecia in Lecanoromycetes. Mycol Res 108:1111–1118 Gueidan C, Roux C, Lutzoni F (2007) Using a multigene phylogenetic analysis to assess generic delineation and character evolution in Verrucariaceae (Verrucariales, Ascomycota). Mycol Res 111:1145–1168 Hafellner J (1984) Studien in Richtung einer natu¨rlicheren Gliederung der Sammelfamilien Lecanoraceae und Lecideaceae. Beih Nova Hedwigia 79:241–371 Hansen K, Læssøe T, Pfister DH (2001) Phylogenetics of the Pezizaceae, with an emphasis on Peziza. Mycologia 93:958–990 Hansen K, Perry BA, Pfister DH (2005) Phylogenetic origins of two cleistothecial fungi, Orbicula parietina and Lasiobolidium orbiculoides, within the operculate discomycetes. Mycologia 97:1023–1033 Hansen K, Pfister DH (2006) Systematics of the Pezizomycetes – the operculate discomycetes. Mycologia 98: 1029–1040 Hawksworth DL (1991) The fungal dimension of biodiversity – magnitude, significance, and conservation. Mycol Res 95:641–655

Fruiting Body Evolution in the Ascomycota: a Molecular Perspective Hawksworth DL (2001) The magnitude of fungal diversity: the 1.5 million species estimate revisited. Mycol Res 105:1422–1432 Henssen A (1963) Eine Revision der Flechtenfamilien Lichinaceae und Ephebaceae. Symb Bot Ups 18:1–123 Henssen A (1969a) Die Entstehung des Thallusrandes bei den Pannariaceen (Lichenes) mit einer generellen Diskussion uber die Entwicklung lecanoriner und biatoriner Flechtenapothecien. Ber Dtsch Bot Gesell 82:235–248 Henssen A (1969b) Eine Studie u¨ber die Gattung Arctomia. Svensk Bot Tideskr 63:126–138 Henssen A (1970) Die Apothecienentwicklung bei Umbilicaria Hoffm. emend Frey. Vortr Gesamtgeb Bot NF 4:103–126 Henssen A (1976) Studies in the developmental morphology of lichenized Ascomycetes. In: Brown DH, Hawksworth DL, Bailey RH (eds) Lichenology: progress and problems. Academic Press, London, pp 107–138 Henssen A (1994) Contributions to the morphology ans species delimitation in Heppia sensu stricto (lichenized Ascomycotina). Acta Bot Fenn 150:57–73 Henssen A, Jahns HM (1974) Lichenes. Thieme, Stuttgart Henssen A, Keuck G, Renner B, Vobis G (1981) The lecanoralean centrum. In: Reynolds DR (ed) Ascomycete systematics: the Luttrellian concept. Springer, Berlin Heidelberg New York, pp 138–234 Henssen A, Lu¨cking R (2002) Morphology, anatomy, and ontogeny in the Asterothyriaceae (Ascomycota: Ostropales), a misunderstood group of lichenized fungi. Ann Bot Fenn 39:273–299 Henssen A, Thor G (1994) Developmental morphology of the ‘Zwischengruppe’ between Ascohymeniales and Ascoloculares. In: Hawksworth D (ed) Ascomycete systematics problems and perspectives in the nineties. NATO Advanced Science Institutes Series. Plenum, New York, pp 43–56 Hibbett DS (2004) Trends in morphological evolution in homobasidiomycetes inferred using maximum likelihood: A comparison of binary and multistate approaches. Syst Biol 53:889–903 Hibbett DS (2007) After the gold rush, or before the flood? Evolutionary morphology of mushroom-forming fungi (Agaricomycetes) in the early 21st century. Mycol Res 111:1001–1018 Hibbett DS, Binder M (2002) Evolution of complex fruiting-body morphologies in homobasidiomycetes. Proc Roy Soc London Ser B 269:1963–1969 Hibbett DS, Binder M, Bischoff JF, Blackwell M, Cannon PF, Eriksson OE, Huhndorf S, James T, Kirk PM, Lu¨cking R, Lumbsch HT, Lutzoni F, Matheny PB, McLaughlin DJ, Powell MJ, Redhead S, Schoch CL, Spatafora JW, Stalpers JA, Vilgalys R, Aime MC, Aptroot A, Bauer R, Begerow D, Benny GL, Castlebury LA, Crous PW, Dai Y-C, Gams W, Geiser DM, Griffith GW, Gueidan C, Hawksworth DL, Hestmark G, Hosaka K, Humber RA, Hyde KD, Ironside JE, Koljalg U, Kurtzman CP, Larsson K-H, Lichtwardt R, Longcore J, Miadlikowska J, Miller A, Moncalvo J-M, Mozley-Standridge S, Oberwinkler F, Parmasto E, Reeb V, Rogers JD, Roux C, Ryvarden L, Sampaio JP,

201

Schuler A, Sugiyama J, Thorn RG, Tibell L, Untereiner WA, Walker C, Wang Z, Weir A, Weiss M, White MM, Winka K, Yao Y-J, Zhang N (2007) A higher-level phylogenetic classification of the Fungi. Mycol Res 111:509–547 Honegger R (1982) The ascus apex in lichenized fungi. 3. The Pertusaria-type. Lichenologist 14:205–217 Honegger R (1991) Functional aspects of the lichen symbiosis. Annu Rev Plant Physiol Plant Mol Biol 42: 553–578 Huelsenbeck JP, Bollback JP (2001) Empirical and hierarchical Bayesian estimation of ancestral states. Syst Biol 50:351–366 Huelsenbeck JP, Rannala B, Masly JP (2000) Accommodating phylogenetic uncertainty in evolutionary studies. Science 288:2349–2350 Ingold CT (1971) Fungal spores: their liberation and dispersal. Clarendon, Oxford James TY, Kauff F, Schoch CL, Matheny PB, Hofstetter V, Cox CJ, Celio G, Gueidan C, Fraker E, Miadlikowska J, Lumbsch HT, Rauhut A, Reeb V, Arnold AE, Amtoft A, Stajich JE, Hosaka K, Sung GH, Johnson D, O’Rourke B, Crockett M, Binder M, Curtis JM, Slot JC, Wang Z, Wilson AW, Schussler A, Longcore JE, O’Donnell K, Mozley-Standridge S, Porter D, Letcher PM, Powell MJ, Taylor JW, White MM, Griffith GW, Davies DR, Humber RA, Morton JB, Sugiyama J, Rossman AY, Rogers JD, Pfister DH, Hewitt D, Hansen K, Hambleton S, Shoemaker RA, Kohlmeyer J, Volkmann-Kohlmeyer B, Spotts RA, Serdani M, Crous PW, Hughes KW, Matsuura K, Langer E, Langer G, Untereiner WA, Lucking R, Budel B, Geiser DM, Aptroot A, Diederich P, Schmitt I, Schultz M, Yahr R, Hibbett DS, Lutzoni F, McLaughlin DJ, Spatafora JW, Vilgalys R (2006) Reconstructing the early evolution of Fungi using a six-gene phylogeny. Nature 443:818–822 Janex-Favre M (1970) Sur le de´veloppement et la structure des ascocarpes et la position syste´matique du mycobionte de Lichen pyre´nocarpe Arthopyrenia conoidea (Fr.) Zahlbr. Rev Bryol Lichenol 37:163–182 Kirk PM, Cannon PF, David JC, Stalpers JA (2001) Ainsworth and Bisby’s dictionary of the fungi. CAB International, Egham Læssøe T, Hansen K (2007) Truffle trouble: what happened to the Tuberales? Mycol Res 111:1075–1099 Landvik S (1996) Neolecta, a fruit-body-producing genus of the basal ascomycetes, as shown by SSU and LSU rDNA sequences. Mycol Res 100:199–202 Landvik S, Eriksson OE, Berbee ML (2001) Neolecta – a fungal dinosaur? Evidence from beta-tubulin amino acid sequences. Mycologia 93:1151–1163 Landvik S, Schumacher TK, Eriksson OE, Moss ST (2003) Morphology and ultrastructure of Neolecta species. Mycol Res 107:1021–1031 Liu YJ, Hall BD (2004) Body plan evolution of ascomycetes, as inferred from an RNA polymerase II, phylogeny. Proc Natl Acad Sci USA 101:4507–4512 Liu YJJ, Whelen S, Benjamin DH (1999) Phylogenetic relationships among ascomycetes: evidence from an RNA polymerase II subunit. Mol Biol Evol 16:1799–1808

202

Imke Schmitt

Lumbsch HT (1999) The ascoma development in Mycoporum elabens (Mycoporaceae, Dothideales). Plant Biol 1:321–326 Lumbsch HT (2000) Phylogeny of filamentous ascomycetes. Naturwissenschaften 87:335–342 Lumbsch HT, del Prado R, Kantvilas G (2005a) Gregorella, a new genus to accommodate Moelleropsis humida and a molecular phylogeny of Arctomiaceae. Lichenologist 37:291–302 Lumbsch HT, Huhndorf SM (2007) Whatever happened to the pyrenomycetes and loculoascomycetes? Mycol Res 111:1064–1074 Lumbsch HT, Lindemuth R, Schmitt I (2000) Evolution of filamentous ascomycetes inferred from LSU rDNA sequence data. Plant Biol 2:525–529 Lumbsch HT, Mangold A, Lu¨cking R, Garcı´a MA, Martı´n MP (2004) Phylogenetic position of the genera Nadvornikia and Pyrgillus (Ascomycota) based on molecular data. Symb Bot Ups 34:9–17 Lumbsch HT, Schmitt I, Barker D, Pagel M (2006) Evolution of micromorphological and chemical characters in the lichen-forming fungal family Pertusariaceae. Biol J Linn Soc 89:615–626 Lumbsch HT, Schmitt I, Do¨ring H, Wedin M (2001a) ITS sequence data suggest variability of ascus types and support ontogenetic characters as phylogenetic discriminators in the Agyriales (Ascomycota). Mycol Res 105:265–274 Lumbsch HT, Schmitt I, Lindemuth R, Miller A, Mangold A, Fernandez F, Huhndorf S (2005b) Performance of four ribosomal DNA regions to infer higher-level phylogenetic relationships of inoperculate euascomycetes (Leotiomyceta). Mol Phylogenet Evol 34:512–524 Lumbsch HT, Schmitt I, Mangold A, Wedin M (2007) Ascus types are phylogenetically misleading in Trapeliaceae and Agyriaceae (Ostropomycetidae, Ascomycota). Mycol Res 111:1133–1141 Lumbsch HT, Schmitt I, Messuti MI (2001b) Utility of nuclear SSU and LSU rDNA data sets to discover the ordinal placement of the Coccotremataceae (Ascomycota). Org Divers Evol 1:99–112 Lumbsch HT, Wirtz N, Lindemuth R, Schmitt I (2002) Higher level phylogenetic relationships of euascomycetes (Pezizomycotina) inferred from a combined analysis of nuclear and mitochondrial sequence data. Mycol Prog 1:57–70 Luttrell ES (1951) Taxonomy of pyrenomycetes. Univ Miss Stud 24:1–120 Luttrell ES (1955) The ascostromatic ascomycetes. Mycologia 47:511–532 Lutzoni F, Kauff F, Cox CJ, McLaughlin D, Celio G, Dentinger B, Padamsee M, Hibbett D, James TY, Baloch E, Grube M, Reeb V, Hofstetter V, Schoch C, Arnold AE, Miadlikowska J, Spatafora J, Johnson D, Hambleton S, Crockett M, Shoemaker R, Sung GH, Lu¨cking R, Lumbsch HT, O’Donnell K, Binder M, Diederich P, Ertz D, Gueidan C, Hansen K, Harris RC, Hosaka K, Lim YW, Matheny B, Nishida H, Pfister D, Rogers J, Rossman A, Schmitt I, Sipman H, Stone J, Sugiyama J, Yahr R, Vilgalys R (2004) Assembling the fungal tree of life: progress, classification, and evolution of subcellular traits. Am J Bot 91:1446–1480

Lutzoni F, Pagel M, Reeb V (2001) Major fungal lineages are derived from lichen symbiotic ancestors. Nature 411:937–940 Maddison DR, Maddison WP (2000) MacClade. Sinauer Associates, Sunderland Maddison WP (1995) Calculating the probability distributions of ancestral states reconstructed by parsimony on phylogenetic trees. Syst Bio 44:474–481 Malloch D (1981) The plectomycete centrum. In: Reynolds DR (ed) Ascomycete systematics: the Luttrellian concept. Springer, Berlin Heidelberg New York, pp 73–91 Miadlikowska J, Lutzoni F (2004) Phylogenetic classification of peltigeralean fungi (Peltigerales, Ascomycota) based on ribosomal RNA small and large subunits. Am J Bot 91:449–464 Muggia L, Hafellner J, Wirtz N, Hawksworth DL, Grube M (2008) The sterile microfilamentous lichenized fungi Cystocoleus ebeneus and Racodium rupestre are relatives of plant pathogens and clinically important dothidealean fungi. Mycol Res 112:50–56 Nannfeldt JA (1932) Studien u¨ber die Morphologie und Systematik der nicht-lichenisierten inoperculaten Discomyceten. Nov Act Reg Soc Upsal 8:1–368 Nolting N, Po¨ggeler S (2006) A MADS box protein interacts with a mating-type protein and is required for fruiting body development in the homothallic ascomycete Sordaria macrospora. Eukayot Cell 5:1043–1056 Nowrousian M, Ku¨ck U (2006) Comparative gene expression analysis of fruiting body development in two filamentous fungi. FEMS Microbiol Lett 257:328–335 Nowrousian M, Ringelberg C, Dunlap JC, Loros JL, Ku¨ck U (2005) Cross-species microarray hybridization to identify developmentally regulated genes in the filamentous fungus Sordaria macrospora. Mol Gen Genom 273:137–149 O’Donnell K, Cigelnik E, Weber NS, Trappe JM (1997) Phylogenetic relationships among ascomycetous truffles and the true and false morels inferred from 18S and 28S ribosomal DNA sequence analysis. Mycologia 89:48–65 O’Donnell K, Ward TJ, Geiser DM, Kistler HC, Aokid T (2004) Genealogical concordance between the mating type locus and seven other nuclear genes supports formal recognition of nine phylogenetically distinct species within the Fusarium graminearum clade. Fung Genet Biol 41:600–623 Pagel M (1997) Inferring evolutionary processes from phylogenies. Zool Scr 26:331–348 Pagel M (1999) The maximum likelihood approach to reconstructing ancestral character states of discrete characters on phylogenies. Syst Bio 48:612–622 Pagel M, Meade A (2006) Bayesian analysis of correlated evolution of discrete characters by reversible-jump Markov chain Monte Carlo. Am Nat 167:808–825 Pagel M, Meade A, Barker D (2004) Bayesian estimation of ancestral character states on phylogenies. Syst Bio 53:673–684 Parguey-Leduc A, Janex-Favre MC (1981) The ascocarps of ascohymenial pyrenomycetes. In: Reynolds DR (ed) Ascomycete systematics: the Luttrellian concept. Springer, Berlin Heidelberg New York, pp 102–123

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

203

Spatafora JW (1995) Ascomal evolution of filamentous ascomycetes – evidence from molecular-data. Can J Bot 73:S811–S815 Spatafora JW, Sung GH, Johnson D, Hesse C, O’Rourke B, Serdani M, Spotts R, Lutzoni F, Hofstetter V, Miadlikowska J, Reeb V, Gueidan C, Fraker E, Lumbsch T, Lucking R, Schmitt I, Hosaka K, Aptroot A, Roux C, Miller AN, Geiser DM, Hafellner J, Hestmark G, Arnold AE, Budel B, Rauhut A, Hewitt D, Untereiner WA, Cole MS, Scheidegger C, Schultz M, Sipman H, Schoch CL (2006) A five-gene phylogeny of Pezizomycotina. Mycologia 98:1018–1028 Spatafora JW, Sung GH, Sung JM, Hywel-Jones NL, White JF (2007) Phylogenetic evidence for an animal pathogen origin of ergot and the grass endophytes. Mol Ecol 16:1701–1711 Spatafora JW, Volkmann-Kohlmeyer B, Kohlmeyer J (1998) Independent terrestrial origins of the Halosphaeriales (marine Ascomycota). Am J Bot 85: 1569–1580 Suh SO, Blackwell M (1999) Molecular phylogeny of the cleistothecial fungi placed in Cephalothecaceae and Pseudeurotiaceae. Mycologia 91:836–848 Tehler A (1995) Morphological data, molecular-data, and total evidence in phylogenetic analysis. Can J Bot 73:S667–S676 Tehler A, Irestedt M (2007) Parallel evolution of lichen growth forms in the family Roccellaceae (Arthoniales, Ascomycota). Cladistics 23:432–454 Thell A, Mattsson J-E, Ka¨rnefelt I (1995) Lecanoralean ascus types in the lichenized families Alectoriaceae and Parmeliaceae. Cryptogam Bot 5:120–127 Thiers HD (1984) The secotioid syndrome. Mycologia 76: 1–8 Tibell L, Wedin M (2000) Mycocaliciales, a new order for nonlichenized calicioid fungi. Mycologia 92:577–581 Trail F (2007) Fungal cannons: explosive spore discharge in the Ascomycota. FEMS Microbiol Lett 276:12–18 Vainio EA (1890) E´tude sur la classification naturelle et la morphologie des lichens du Bre´sil. I. Acta Soc Faun Flora Fenn 7:1–256 van Brummelen J (1995) A world-monograph of the genus Pseudombrophila (Pezizales, Ascomycotina). IHW, Munich von Arx JA (1973) Ostiolate and nonostiolate Pyrenomycetes. Proc K Ned Akad Wet 76:289–296 von Ho¨hnel F (1907) Fragmente zur Mykologie III Mitt, Nr 92–155. Sitz Kaiserl Akad Wiss Wien, Math Nat Kl, Abt 1 116:615–647 Wang Z, Binder M, Schoch CL, Johnston PR, Spatafora JW, Hibbett DS (2006) Evolution of helotialean fungi (Leotiomycetes, Pezizomycotina): a nuclear rDNA phylogeny. Mol Phylogenet Evol 41:295–312 Wedin M, Do¨ring H, Gilenstam G (2004) Saprotrophy and lichenization as options for the same fungal species on different substrata: environmental plasticity and fungal lifestyles in the Stictis–Conotrema complex. New Phytol 164:459–465 Wedin M, Tibell L (1997) Phylogeny and evolution of Caliciaceae, Mycocaliciaceae, and Sphinctrinaceae

204

Imke Schmitt

(Ascomycota), with notes on the evolution of the prototunicate ascus. Can J Bot 75:1236–1242 Wedin M, Wiklund E, Crewe A, Do¨ring H, Ekman S, Nyberg A, Schmitt I, Lumbsch HT (2005) Phylogenetic relationships of Lecanoromycetes (Ascomycota) as revealed by analyses of mtSSU and nLSU rDNA sequence data. Mycol Res 109:159–172 Weir A, Blackwell M (2001) Molecular data support the Laboulbeniales as a separate class of Ascomycota, Laboulbeniomycetes. Mycol Res 105:1182–1190

Wik L, Karlsson M, Johannesson H (2008) The evolutionary trajectory of the mating-type (mat) genes in Neurospora relates to reproductive behavior of taxa. BMC Evol Biol 8:109 Winka K (2000) Phylogenetic relationships within the Ascomycota based on 18S rDNA sequences. PhD thesis, Umea˚ University, Umea˚ Zoller S, Scheidegger C, Sperisen C (1999) PCR primers for the amplification of mitochondrial small subunit ribosomal DNA of lichen-forming ascomycetes. Lichenologist 31:511–516

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

206

James K. Hane et al.

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

Genomic and Comparative Analysis of the Class Dothideomycetes

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

207

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.

208

James K. Hane et al.

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

NCBI [Trace Archive]

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

NCBI [Trace Archive]

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

NCBI [Trace Archive]

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

NCBI [Trace Archive]

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

James K. Hane et al.

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

James K. Hane et al.

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)

Genomic and Comparative Analysis of the Class Dothideomycetes

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

James K. Hane et al.

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

Genomic and Comparative Analysis of the Class Dothideomycetes

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

James K. Hane et al.

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.

Genomic and Comparative Analysis of the Class Dothideomycetes

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

222

James K. Hane et al.

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).

Genomic and Comparative Analysis of the Class Dothideomycetes

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

223

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

224

James K. Hane et al.

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

Genomic and Comparative Analysis of the Class Dothideomycetes

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-

225

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

226

James K. Hane et al.

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.

References Andrew M, Peever TL and Pryor BM (2009) An expanded multilocus phylogeny does not resolve morphological species within the small-spored Altemaria species complex. Mycologia 101:95–109 Aptroot A, Lu¨cking R, Sipman H, Umana L and Chaves J-L (2008) Pyrenocarpous lichens with bitunicate asci. A first assessment of the lichen biodiversity inventory in Costa Rica. Bibl Lichenol 97:1–162 Bao Z and Eddy SR (2002) Automated de novo identification of repeat sequence families in sequenced genomes. Genome Res 12(8):1269–1276 Bearchell SJ, Fraaije BA, Shaw MW and Fitt BD (2005) Wheat archive links long-term fungal pathogen population dynamics to air pollution. Proc Natl Acad Sci 102:5438–5442 Bhathal JS, Loughman R and Speijers J (2003) Yield reduction in wheat in relation to leaf disease from yellow (tan) spot and Septoria nodorum blotch. Eur J Plant Pathol 109:435–443 Boehm EW, Mugambi GK, Miller AN, Huhndorf SM, Marincowitz S, Spatafora JW and Schoch CL (2009) A molecular phylogenetic reappraisal of the Hysteriaceae, Mytilinidiaceae and Gloniaceae (Pleosporomyce-

tidae, Dothideomycetes) with keys to world species. Stud Mycol 64:49–83 Bringans S, Hane JK, Casey T, Tan KC, Lipscombe R, Solomon PS and Oliver RP (2009) Deep proteogenomics; high throughput gene validation by multidimensional liquid chromatography and mass spectrometry of proteins from the fungal wheat pathogen Stagonospora nodorum. BMC Bioinformatics 10: 301 Brun H, Levivier S, Somda I, Ruer D, Renard M and Chevre AM (2000) A field method for evaluating the potential durability of new resistance sources: application to the Leptosphaeria maculans–Brassica napus pathosystem. Phytopathology 90(9):961–966 Cannon SB, Sterck L, Rombauts S, Sato S, Cheung F, Gouzy J, Wang X, Mudge J, Vasdewani J, Schiex T, Spannagl M, Monaghan E, Nicholson C, Humphray SJ, Schoof H, Mayer KF, Rogers J, Quetier F, Oldroyd GE, Debelle F, Cook DR, Retzel EF, Roe BA, Town CD, Tabata S, Van de Peer Y and Young ND (2006) Legume genome evolution viewed through the Medicago truncatula and Lotus japonicus genomes. Proc Natl Acad Sci USA 103(40):14959–14964 Cooley RN and Caten CE (1991) Variation in electrophoretic karyotype between strains of Septoria nodorum. Mol Gen Genet 228(1/2):17–23 Curtis MJ and Wolpert TJ (2004) The victorin-induced mitochondrial permeability transition precedes cell shrinkage and biochemical markers of cell death, and shrinkage occurs without loss of membrane integrity. Plant J 38(2):244–259 Dear PH and Cook PR (1993) Happy mapping: linkage mapping using a physical analogue of meiosis. Nucleic Acids Res 21(1):13–20 Edgar RC and Myers EW (2005) PILER: identification and classification of genomic repeats. Bioinformatics 21 [Suppl 1]:i152–il58 Espagne E, Lespinet O, Malagnac F, Da Silva C, Jaillon O, Porcel BM, Couloux A, Aury JM, Segurens B, Poulain J, Anthouard V, Grossetete S, Khalili H, Coppin E, Dequard-Chablat M, Picard M, Contamine V, Arnaise S, Bourdais A, Berteaux-Lecellier V, Gautheret D, de Vries RP, Battaglia E, Coutinho PM, Danchin EG, Henrissat B, Khoury RE, Sainsard-Chanet A, Boivin A, Pinan-Lucarre B, Sellem CH, Debuchy R, Wincker P, Weissenbach J and Silar P (2008) The genome sequence of the model ascomycete fungus Podospora anserina. Genome Biol 9(5):R77 Faris JD, Anderson JA, Francl LJ and Jordahl JG (1996) Chromosomal location of a gene conditioning insensitivity in wheat to a necrosis-inducing culture filtrate from Pyrenophora tritici-repentis. Phytopathology 86: 459–463 Fitt BDL, Brun H, Barbetti MJ and Rimmer SR (2006) World-wide importance of Phoma stem canker (Leptosphaeria maculans and L. biglobosa) on oilseed rape (Brassica napus). Eur J Plant Pathol 114:3–15 Friesen TL, Stukenbrock EH, Liu Z, Meinhardt S, Ling H, Faris JD, Rasmussen JB, Solomon PS, McDonald BA and Oliver RP (2006) Emergence of a new disease as a result of interspecific virulence gene transfer. Nat Genet 38(8):953–956

Genomic and Comparative Analysis of the Class Dothideomycetes Fudal I, Ross S, Brun H, Besnard AL, Ermel M, Kuhn ML, Balesdent MH and Rouxel T (2009) Repeat-induced point mutation (RIP) as an alternative mechanism of evolution toward virulence in Leptosphaeria maculans. Mol Plant Microbe Interact 22(8):932–941 Galagan JE, Calvo SE, Borkovich KA, Selker EU, Read ND, Jaffe D, FitzHugh W, Ma LJ, Smirnov S, Purcell S, Rehman B, Elkins T, Engels R, Wang S, Nielsen CB, Butler J, Endrizzi M, Qui D, Ianakiev P, Bell-Pedersen D, Nelson MA, Werner-Washburne M, Selitrennikoff CP, Kinsey JA, Braun EL, Zelter A, Schulte U, Kothe GO, Jedd G, Mewes W, Staben C, Marcotte E, Greenberg D, Roy A, Foley K, Naylor J, Stange-Thomann N, Barrett R, Gnerre S, Kamal M, Kamvysselis M, Mauceli E, Bielke C, Rudd S, Frishman D, Krystofova S, Rasmussen C, Metzenberg RL, Perkins DD, Kroken S, Cogoni C, Macino G, Catcheside D, Li W, Pratt RJ, Osmani SA, DeSouza CP, Glass L, Orbach MJ, Berglund JA, Voelker R, Yarden O, Plamann M, Seiler S, Dunlap J, Radford A, Aramayo R, Natvig DO, Alex LA, Mannhaupt G, Ebbole DJ, Freitag M, Paulsen I, Sachs MS, Lander ES, Nusbaum C and Birren B (2003) The genome sequence of the filamentous fungus Neurospora crassa. Nature 422(6934):859–868 Galagan JE, Calvo SE, Cuomo C, Ma LJ, Wortman JR, Batzoglou S, Lee SI, Basturkmen M, Spevak CC, Clutterbuck J, Kapitonov V, Jurka J, Scazzocchio C, Farman M, Butler J, Purcell S, Harris S, Braus GH, Draht O, Busch S, D’Enfert C, Bouchier C, Goldman GH, Bell-Pedersen D, Griffiths-Jones S, Doonan JH, Yu J, Vienken K, Pain A, Freitag M, Selker EU, Archer DB, Penalva MA, Oakley BR, Momany M, Tanaka T, Kumagai T, Asai K, Machida M, Nierman WC, Denning DW, Caddick M, Hynes M, Paoletti M, Fischer R, Miller B, Dyer P, Sachs MS, Osmani SA and Birren BW (2005) Sequencing of Aspergillus nidulans and comparative analysis with A. fumigatus and A. oryzae. Nature 438(7071):1105–1115 Gardiner DM, Cozijnsen AJ, Wilson LM, Pedras MS and Howlett BJ (2004) The sirodesmin biosynthetic gene cluster of the plant pathogenic fungus Leptosphaeria maculans. Mol Microbiol 53(5):1307–1318 Gardiner DM and Howlett BJ (2005) Bioinformatic and expression analysis of the putative gliotoxin biosynthetic gene cluster of Aspergillus fumigatus. FEMS Microbiol Lett 248(2):241–248 Gladieux P, Zhang XG, Afoufa-Bastien D, Valdebenito Sanhueza RM, Sbaghi M and Le Cam B (2008) On the origin and spread of the Scab disease of apple: out of central Asia. PLoS One 3(1):e1455 Goodwin SB, Ben M’Barek S, Dhillon B, Wittenberg A, Crane CF, Van der Lee TAJ, Grimwood J, Aerts A, Antoniw J, Bailey A, Bluhm B, Bowler J, Bristow J, Brokstein P, Canto-Canche B, Churchill A, CondeFerra`ez L, Cools H, Coutinho PM, Csukai M, Dehal P, Donzelli B, Foster AJ, Hammond-Kosack K, Hane J, Henrissat B, Kilian A, Koopmann E, Kourmpetis Y, Kuo A, Kuzniar A, Lindquist E, Maliepaard C, Martins N, Mehrabi R, Oliver R, Platt D, Ponomarenko A, Rudd J, Salamov A, Schwarz J, Shapiro H, Stergiopoulos I, Torriani S, Tu H, de Vries R, Wiebenga A,

227

Zwiers L-H, Grigoriev IV and Kema GHJ (2011) Finished genome of Mycosphaerella graminicola reveals stealth pathogenesis and dispensome structure. PLoS Genet (in press) Groenewald M, Groenewald JZ and Crous PW (2005) Distinct species exist within the Cercospora apii morphotype. Phytopathology 95:951–959 Hane JK (2011) Bioinformatic genome analysis of the necrotrophic wheat-pathogenic fungus Phaeosphaeria nodorum. PhD thesis, Murdoch University, Murdoch Hane JK, Lowe RGT, Solomon PS, Tan KC, Schoch CL, Spatafora JW, Crous PW, Kodira C, Birren BW, Galagan JE, Torriani SFF, McDonald BA and Oliver RP (2007) Dothideomycete–plant interactions illuminated by genome sequencing and EST analysis of the wheat pathogen Stagonospora nodorum. Plant Cell 19(11):3347–3368 Hane JK and Oliver RP (2008) RIPCAL: a tool for alignmentbased analysis of repeat-induced point mutations in fungal genomic sequences. BMC Bioinformatics 9:478 Hane JK and Oliver RP (2010) In silico reversal of repeatinduced point mutation (RIP) identifies the origins of repeat families and uncovers obscured duplicated genes. BMC Genomics 2010, 11:655 Hane JK, Rouxel T, Howlett BJ, Kema GHJ, Goodwin SB, Oliver RP (2011) A novel mode of chromosomal evolution called Mesosynteny that is peculiar to filamentous Ascomycete fungi. Genome Biology (in press). Hardwick NV, Jones DR and Slough JE (2001) Factors affecting diseases of winter wheat in England and Wales, 1989–98. Plant Pathol 50:453–462 Hashim I (1998) Disease survey. IRRDB, Kuala Lumpur Hooker AL, Smith DR, Lim SM and Musson MD (1970) Physiological races of Helminthosporium maydis and disease resistance. Plant Dis Rep 54:1109–1110 Howlett BJ, Idnurm A and Pedras MSC (2001) Leptosphaeria maculans, the causal agent of blackleg disease of Brassicas. Fungal Genet Biol 33:1–14 Idnurm A and Howlett BJ (2003) Analysis of loss of pathogenicity mutants reveals that repeat-induced point mutations can occur in the Dothideomycete Leptosphaeria maculans. Fungal Genet Biol 39(1):31–37 IpCho SV S (2010) Pathogenicity of Stagonospora nodorum. PhD thesis, Murdoch University, Murdoch Irelan JT, Hagemann AT and Selker EU (1994) High frequency repeat-induced point mutation (RIP) is not associated with efficient recombination in Neurospora. Genetics 138(4):1093–1103 Kale SD, Gu B, Capelluto DGS, Dou D, Cronin A, Arredondo FD, Feldman E, Fudal I, Rouxel T, Lawrence CB, Shan W and Tyler BM (2010) External phosphatidylinositol-3-phosphate mediates host cell entry by eukaryotic pathogen effectors. Cell 142(2): 284–295 Khaldi N, Collemare J, Lebrun M-H and Wolfe KH (2008) Evidence for horizontal transfer of a secondary metabolite gene cluster between fungi. Genome Biol 9:R18 Kohn M, Kehrer-Sawatzki H, Vogel W, Graves JA and Hameister H (2004) Wide genome comparisons reveal the origins of the human X chromosome. Trends Genet 20(12):598–603

228

James K. Hane et al.

Kroken S, Glass NL, Taylor JW, Yoder OC and Turgeon BG (2003) Phylogenomic analysis of type I polyketide synthase genes in pathogenic and saprobic ascomycetes. Proc Natl Acad Sci USA 100(26): 15670–15675 Kurtz S, Phillippy A, Delcher AL, Smoot M, Shumway M, Antonescu C and Salzberg SL (2004) Versatile and open software for comparing large genomes. Genome Biol 5(2):R12 Levings CS, 3rd and Siedow JN (1992) Molecular basis of disease susceptibility in the Texas cytoplasm of maize. Plant Mol Biol 19(1):135–147 Li CX and Cowling WA (2003) Identification of a single dominant allele for resistance to blackleg in Brassica napus “Surpass 400”. Plant Breed 122:485–488 Liu Z, Faris JD, Oliver RP, Tan KC, Solomon PS, McDonald MC, McDonald BA, Nunez A, Lu S, Rasmussen JB and Friesen TL (2009) SnTox3 acts in effector triggered susceptibility to induce disease on wheat carrying the Snn3 gene. PLoS Pathog 5(9):e1000581 Luttrell ES (1955) The ascostromatic Ascomycetes. Mycologia 47:511–532 Machida M, Asai K, Sano M, Tanaka T, Kumagai T, Terai G, Kusumoto K, Arima T, Akita O, Kashiwagi Y, Abe K, Gomi K, Horiuchi H, Kitamoto K, Kobayashi T, Takeuchi M, Denning DW, Galagan JE, Nierman WC, Yu J, Archer DB, Bennett JW, Bhatnagar D, Cleveland TE, Fedorova ND, Gotoh O, Horikawa H, Hosoyama A, Ichinomiya M, Igarashi R, Iwashita K, Juvvadi PR, Kato M, Kato Y, Kin T, Kokubun A, Maeda H, Maeyama N, Maruyama J, Nagasaki H, Nakajima T, Oda K, Okada K, Paulsen I, Sakamoto K, Sawano T, Takahashi M, Takase K, Terabayashi Y, Wortman JR, Yamada O, Yamagata Y, Anazawa H, Hata Y, Koide Y, Komori T, Koyama Y, Minetoki T, Suharnan S, Tanaka A, Isono K, Kuhara S, Ogasawara N and Kikuchi H (2005) Genome sequencing and analysis of Aspergillus oryzae. Nature 438(7071): 1157–1161 Malkus A, Song Q, Cregan P, Arseniuk E and Ueng PP (2009) Genetic linkage map of Phaeosphaeria nodorum, the causal agent of stagonospora nodorum blotch disease of wheat. Eur J Plant Pathol:1–10 Marcet-Houben M and Gabaldon T (2010) Acquisition of prokaryotic genes by fungal genomes. Trends Genet 26(1):5–8 McLysaght A, Enright AJ, Skrabanek L and Wolfe KH (2000) Estimation of synteny conservation and genome compaction between pufferfish (Fugu) and human. Yeast 17(1):22–36 Murray GM and Brennan JP (2009) Estimating disease losses to the Australian wheat industry. Aust Plant Pathol 38:558–570 Nowrousian M, Stajich JE, Chu M, Engh I, Espagne E, Halliday K, Kamerewerd J, Kempken F, Knab B, Kuo HC, Osiewacz HD, Poggeler S, Read ND, Seiler S, Smith KM, Zickler D, Kuck U and Freitag M (2010) De novo assembly of a 40 Mb eukaryotic genome from short sequence reads: Sordaria macrospora, a model organism for fungal morphogenesis. PLoS Genet 6(4): e1000891

Oliver R (2009) Plant breeding for disease resistance in the age of effectors. Phytoparasitica 37:1–5 Oliver RP (1992) A model system for the study of plant– fungal interactions: Tomato leaf mold caused by Cladosporium fulvum.In: Verma DPS (ed) Molecular signals in plant–microbe communications. CRC, Boca Raton, pp 97–106 Oliver RP and Solomon PS (2010) New developments in pathogenicity and virulence of necrotrophs. Curr Opin Plant Biol 13:1–5 Palmer C-L and Skinner W (2002) Mycosphaerella graminicola: latent infection, crop devastation and genomics. Mol Plant Pathol 3(2):63–70 Pel HJ, de Winde JH, Archer DB, Dyer PS, Hofmann G, Schaap PJ, Turner G, de Vries RP, Albang R, Albermann K, Andersen MR, Bendtsen JD, Benen JA, van den Berg M, Breestraat S, Caddick MX, Contreras R, Cornell M, Coutinho PM, Danchin EG, Debets AJ, Dekker P, van Dijck PW, van Dijk A, Dijkhuizen L, Driessen AJ, d’Enfert C, Geysens S, Goosen C, Groot GS, de Groot PW, Guillemette T, Henrissat B, Herweijer M, van den Hombergh JP, van den Hondel CA, van der Heijden RT, van der Kaaij RM, Klis FM, Kools HJ, Kubicek CP, van Kuyk PA, Lauber J, Lu X, van der Maarel MJ, Meulenberg R, Menke H, Mortimer MA, Nielsen J, Oliver SG, Olsthoorn M, Pal K, van Peij NN, Ram AF, Rinas U, Roubos JA, Sagt CM, Schmoll M, Sun J, Ussery D, Varga J, Vervecken W, van de Vondervoort PJ, Wedler H, Wosten HA, Zeng AP, van Ooyen AJ, Visser J and Stam H (2007) Genome sequencing and analysis of the versatile cell factory Aspergillus niger CBS 513.88. Nat Biotechnol 25(2):221–231 Pennacchio LA (2003) Insights from human/mouse genome comparisons. Mamm Genome 14(7):429–436 Phan HT, Ellwood SR, Hane JK, Ford R, Materne M and Oliver RP (2007) Extensive macrosynteny between Medicago truncatula and Lens culinaris ssp. culinaris. Theor Appl Genet 114(3):549–558 Ploetz RC (2001) Black sigatoka of banana. The Plant Health Instructor, New York Price AL, Jones NC and Pevzner PA (2005) De novo identification of repeat families in large genomes. Bioinformatics 21 [Suppl 1]:i351–i358 Quesneville H, Nouaud D and Anxolabehere D (2005) Recurrent recruitment of the THAP DNA-binding domain and molecular domestication of the P-transposable element. Mol Biol Evol 22(3):741–746 Rouxel T, Grandaubert J, Hane J, Hoede C, van de Wouw A, Couloux A, Dominguez V, Anthouard V, Bally P, Bourras S, Cozijnsen A, Ciuffetti L, Dimaghani A, Duret L, Fudal I, Goodwin S, Gout L, Glaser N, Kema G, Lapalu N, Lawrence C, May K, Meyer M, Ollivier B, Poulain J, Turgeon G, Tyler BM, Vincent D, Weissenbach J, Amselem J, Balesdent M-H, Howlett BJ, Oliver R, Quesneville H and Wincker P (2011) The patchwork genome of Leptosphaeria maculans: effector diversification driven by location within RIPaffected isochores. Nat Commun 2:202 Rouxel T, Penaud A, Pinochet X, Brun H, Gout L, Delourne R, Schmit J and Bealesdent MH (2003) A 10-year survey of populations of Leptosphaeria maculans in

Genomic and Comparative Analysis of the Class Dothideomycetes France indicates a rapid adaptation towards to Rlm1 resistance gene of oilseed rape. Eur J Plant Pathol 109:871–881 Schmitt I and Lumbsch HT (2009) Ancient horizontal gene transfer from bacteria enhances biosynthetic capabilities of fungi. PLoS One 4(2):e4437 Schoch CL, Crous PW, Groenewald JZ, Boehm EW, Burgess TI, de Gruyter J, de Hoog GS, Dixon LJ, Grube M, Gueidan C, Harada Y, Hatakeyama S, Hirayama K, Hosoya T, Huhndorf SM, Hyde KD, Jones EB, Kohlmeyer J, Kruys A, Li YM, Lucking R, Lumbsch HT, Marvanova L, Mbatchou JS, McVay AH, Miller AN, Mugambi GK, Muggia L, Nelsen MP, Nelson P, Owensby CA, Phillips AJ, Phongpaichit S, Pointing SB, Pujade-Renaud V, Raja HA, Plata ER, Robbertse B, Ruibal C, Sakayaroj J, Sano T, Selbmann L, Shearer CA, Shirouzu T, Slippers B, Suetrong S, Tanaka K, Volkmann-Kohlmeyer B, Wingfield MJ, Wood AR, Woudenberg JH, Yonezawa H, Zhang Y and Spatafora JW (2009) A class-wide phylogenetic assessment of Dothideomycetes. Stud Mycol 64:1–15 Schoch CL, Shoemaker RA, Seifert KA, Hambleton S, Spatafora JW and Crous PW (2006) A multigene phylogeny of the Dothideomycetes using four nuclear loci. Mycologia 98:1043–1054 Schwartz DC, Li X, Hernandez LI, Ramnarain SP, Huff EJ and Wang YK (1993) Ordered restriction maps of Saccharomyces cerevisiae chromosomes constructed by optical mapping. Science 262(5130):110–114 Selker EU, Cambareri EB, Jensen BC and Haack KR (1987) Rearrangement of duplicated DNA in specialized cells of Neurospora. Cell 51(5):741–752 Shultz JL, Ray JD and Lightfoot DA (2007) A sequence based synteny map between soybean and Arabidopsis thaliana. BMC Genomics 8:8 Silva WPK, Wijesundera RIC, Karunanayake EH, Jayasinghe CK and Priyanka UMS (2000) New hosts of Corynespora cassiicola in Sri Lanka. Plant Dis 84:202 Smith DR and White DG (1988) Diseases of corn. In: Sprague GF and Dudley JW (eds) Corn and corn improvement. Agronomy series 18. ASA/CSSA/SSS, Madison, pp 701–766 Solomon PS, Lowe RG T, Tan KC, Waters ODC and Oliver RP (2006) Stagonospora nodorum: cause of stagonospora nodorum blotch of wheat. Mol Plant Pathol 7:147–156 Stukenbrock EH and McDonald BA (2007) Geographical variation and positive diversifying selection in the

229

host-specific toxin SnToxA. Mol Plant Pathol 8(3): 321–332 Thomma BPHJ (2003) Alternaria spp.: from general saprophyte to specific parasite. Mol Plant Pathol 4:225–235 Torriani SF, Goodwin SB, Kema GH, Pangilinan JL and McDonald BA (2008) Intraspecific comparison and annotation of two complete mitochondrial genome sequences from the plant pathogenic fungus Mycosphaerella graminicola. Fungal Genet Biol 45(5): 628–637 Turgeon BG and Baker SE (2007) Genetic and genomic dissection of the Cochliobolus heterostrophus Tox1 locus controlling biosynthesis of the polyketide virulence factor T-toxin. Adv Genet 57:219–261 Tzeng T-H, Lyngholm L, Ford C and Bronson C (1992) A restriction fragment length polymorphism map and electrophoretic karyotype of the fungal maize pathogen Cochliobolus heterostrophus. Genetics 130 (1):81–96 Ullstrup AJ (1972) The impacts of the southern corn leaf blight epidemics of 1970–71. Annu Rev Phytopathol 10:37–50 Van de Wouw AP, Cozijnsen AJ, Hane JK, Brunner PC, McDonald BA, Oliver RP and Howlett BJ (2010) Evolution of linked avirulence effectors in Leptosphaeria maculans is affected by genomic environment and exposure to resistance genes in host plants. PLoS Pathogens 6(11):e1001180 Watters MK, Randall TA, Margolin BS, Selker EU and Stadler DR (1999) Action of repeat-induced point mutation on both strands of a duplex and on tandem duplications of various sizes in Neurospora. Genetics 153(2):705–714 Wei J, Lui K, Chen J, Luo P and Lee-Stadelmann OY (1988) Pathological and physiological identification of race C of Bipolaris maydis in China. Phytopathology 78: 550–554 Wulff BBH, Chakrabarti A and Jones DA (2009) Recognitional Specificity and Evolution in the Tomato– Cladosporium fulvum Pathosystem. Mol Plant Microbe Interact 22(10):1191–1202 Zhang Y, Schoch CL, Fournier J, Crous PW, de Gruyter J, Woudenberg JH, Hirayama K, Tanaka K, Pointing SB, Spatafora JW and Hyde KD (2009) Multi-locus phylogeny of Pleosporales: a taxonomic, ecological and evolutionary re-evaluation. Stud Mycol 64:85–1022

.

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

232

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.

Evolution of Genes for Secondary Metabolism in Fungi

233

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)

234

Ines Teichert and Minou Nowrousian

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

Evolution of Genes for Secondary Metabolism in Fungi

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

235

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

236

Ines Teichert and Minou Nowrousian

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

Evolution of Genes for Secondary Metabolism in Fungi

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

237

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

238

Ines Teichert and Minou Nowrousian

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

Evolution of Genes for Secondary Metabolism in Fungi

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.

239

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

240

Ines Teichert and Minou Nowrousian

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

Evolution of Genes for Secondary Metabolism in Fungi

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,

241

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.

242

Ines Teichert and Minou Nowrousian

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

Evolution of Genes for Secondary Metabolism in Fungi

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

243

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

244

Ines Teichert and Minou Nowrousian

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.

Evolution of Genes for Secondary Metabolism in Fungi

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.

References Akamatsu HO, Chilvers MI, Stewart JE, Peever TL (2010) Identification and function of a polyketide synthase gene responsible for 1,8-dihydroxynaphthalenemelanin pigment biosynthesis in Ascochyta rabiei. Curr Genet (in press) Alviano CS, Farbiarz SR, De Souza W, Angluster J, Travassos LR (1991) Characterization of Fonsecaea pedrosoi melanin. J Gen Microbiol 137:837–844 Austin MB, Saito T, Bowman ME, Haydock S, Kato A, Moore BS, Kay RR, Noel JP (2006) Biosynthesis of Dictyostelium discoideum differentiation-inducing factor by a hybrid type I fatty acid-type III polyketide synthase. Nat Chem Biol 2:494–502 Baker SE, Kroken S, Inderbitzin P, Asvarak T, Li BY, Shi L, Yoder OC, Turgeon BG (2006) Two polyketide synthase-encoding genes are required for biosynthesis of the polyketide virulence factor, T-toxin, by Cochliobolus heterostrophus. Mol Plant Microbe Interact 19:139–149 Baldrian P (2006) Fungal laccases – occurrence and properties. FEMS Microbiol Rev 30:215–242 ¨ , Krappmann S, Ni M, Bok JW, Helmstaedt K, Bayram O Valerius O, Braus-Stromeyer S, Kwon N-J, Keller NP, Yu J-H, Braus GH (2008) VelB/VeA/LaeA complex coordinates light signal with fungal development and secondary metabolism. Science 320:1504–1506 Bell AA, Puhalla JE, Tolmsoff WJ, Stipanovic RD (1976) Use of mutants to establish (þ)-scytalone as

245

an intermediate in melanin biosynthesis by Verticillium dahliae. Can J Microbiol 22:787–799 Bell AA, Wheeler MH (1986) Biosynthesis and functions of fungal melanins. Annu Rev Phytopathol 24:411–451 Bennett JW, Klich M (2003) Mycotoxins. Clin Microbiol Rev 16:497–516 Bo¨hnert HU, Fudal I, Dioh W, Tharreau D, Notteghem JL, Lebrun MH (2004) A putative polyketide synthase/ peptide synthetase from Magnaporthe grisea signals pathogen attack to resistant rice. Plant Cell 16: 2499–2513 Bok JW, Keller NP (2004) LaeA, a regulator of secondary metabolism in Aspergillus spp. Eukaryot Cell 3:527–535 Bok JW, Hoffmeister D, Maggio-Hall LA, Murillo R, Glasner JD, Keller NP (2006a) Genomic mining for Aspergillus natural products. Chem Biol 13:31–37 Bok JW, Noordermeer D, Kale SP, Keller NP (2006b) Secondary metabolic gene cluster silencing in Aspergillus nidulans. Mol Microbiol 61:1636–1645 Bok JW, Chiang Y-M, Szewczyk E, Reyes-Dominguez Y, Davidson AD, Sanchez JF, Lo H-C, Watanabe K, Strauss J, Oakley BR, Wang CCC, Keller NP (2009) Chromatin-level regulation of biosynthetic gene clusters. Nat Chem Biol 5:462–464 Bo¨lker M, Basse CW, Schirawski J (2008) Ustilago maydis secondary metabolism – From genomics to biochemistry. Fungal Genet Biol 45:S88–S93 Bo¨mke C, Rojas MC, Gong F, Hedden P, Tudzynski B (2008) Isolation and characterization of the gibberellin biosynthetic gene cluster in Sphaceloma manihoticola. Appl Environ Microbiol 74:5325–5339 Bo¨mke C, Tudzynski B (2009) Diversity, regulation, and evolution of the gibberellin biosynthetic pathway in fungi compared to plants and bacteria. Phytochemistry 70:1876–1893 Borkovich KA, Alex LA, Yarden O, Freitag M, Turner GE, Read ND, Seiler S, Bell-Pedersen D, Paietta J, Plesofsky N, Plamann M, Goodrich-Tanrikulu M, Schulte U, Mannhaupt G, Nargang FE, Radford A, Selitrennikoff C, Galagan JE, Dunlap JC, Loros JJ, Catcheside D, Inoue H, Aramayo R, Polymenis M, Selker EU, Sachs MS, Marzluf GA, Paulsen I, Davis R, Ebbole DJ, Zelter A, Kalkman ER, O’Rourke R, Bowring F, Yeadon J, Ishii C, Suzuki K, Sakai W, Pratt R (2004) Lessons from the genome sequence of Neurospora crassa: tracing the path from genomic blueprint to multicellular organism. Microbiol Mol Biol Rev 68:1–108 Brakhage AA, Langfelder K, Wanner G, Schmidt A, Jahn B (1999) Pigment biosynthesis and virulence. Contrib Microbiol 2:205–215 Brakhage AA, Tho¨n M, Spro¨te P, Scharf DH, Al-Abdallah Q, Wolke SM, Hortschansky P (2009) Aspects on evolution of fungal beta-lactam biosynthesis gene clusters and recruitment of trans-acting factors. Phytochemistry 70:1801–1811 Brakhage AA, Schroeckh V (2010) Fungal secondary metabolites – Strategies to activate silent gene clusters. Fungal Genet Biol In Press:doi:10.1016/j. fgb.2010.04.004 Brown DW, Dyer RB, McCormick SP, Kendra DF, Plattner RD (2004) Functional demarcation of the Fusarium

246

Ines Teichert and Minou Nowrousian

core trichothecene gene cluster. Fungal Genet Biol 41: 454–462 Buades C, Moya A (1996) Phylogenetic analysis of the isopenicillin-N-synthetase horizontal gene transfer. J Mol Evol 45:537–542 Bushley KE, Ripoll DR, Turgeon BG (2008) Module evolution and substrate specificity of fungal nonribosomal peptide synthetases involved in siderophore biosynthesis. BMC Evol Biol 8:328 Bushley KE, Turgeon BG (2010) Phylogenomics reveals subfamilies of fungal nonribosomal peptide synthetases and their evolutionary relationships. BMC Evol Biol 10:26 Butler MJ, Lazarovits G, Higgins VJ, Lachance MA (1989) Identification of a Black Yeast Isolated from Oak Bark as Belonging to the Genus Phaeococcomyces sp. Analysis of Melanin Produced by the Yeast. Can J Microbiol 35:728–734 Butler MJ, Day AW (1998) Fungal melanins: a review. Can J Microbiol 44:1115–1136 Butler MJ, Day AW, Henson JM, Money NP (2001) Pathogenic properties of fungal melanins. Mycologia 93:1–8 Butler MJ, Gardiner RB, Day AW (2009) Melanin synthesis by Sclerotinia sclerotiorum. Mycologia 101:296–304 Caesar-Tonthat T, Van Ommen KF, Geesey GG, Henson JM (1995) Melanin production by a filamentous soil fungus in response to copper and localization of copper sulfide by sulfide-silver staining. Appl Env Microbiol 61:1968–1975 Calvo AM, Wilson RA, Bok JW, Keller NP (2002) Relationship between secondary metabolism and fungal development. Microbiol Mol Biol Rev 66:447–459 Casadevall A, Rosas AL, Nosanchuk JD (2000) Melanin and virulence in Cryptococcus neoformans. Current Opinion in Microbiology 3:354–358 Chai LY, Netea MG, Sugui J, Vonk AG, van de Sande WW, Warris A, Kwon-Chung KJ, Jan Kullberg B (2009) Aspergillus fumigatus conidial melanin modulates host cytokine response. Immunobiology (in press) Choquer M, Dekkers KL, Chen H-Q, Cao L, Ueng PP, Daub ME, Chung K-R (2005) The CTB1 gene encoding a fungal polyketide synthase is required for cercosporin biosynthesis and fungal virulence of Cercospora nicotianae. Mol Plant Microbe Interact 18:468–476 Chumley FG, Valent B (1990) Genetic analysis of melanindeficient, nonpathogenic mutants of Magnaporthe grisea. Mol Plant Microbe Interact 3:135–143 Claus H, Decker H (2006) Bacterial tyrosinases. Syst Appl Microbiol 29:3–14 Coleman JJ, Rounsley SD, Rodriguez-Carres M, Kuo A, Wasmann CC, Grimwood J, Schmutz J, Taga M, White GJ, Zhou S, Schwartz DC, Freitag M, Ma L-j, Danchin EGJ, Henrissat B, Coutinho PM, Nelson DR, Straney D, Napoli CA, Barker BM, Gribskov M, Rep ˜ ¡r In, Rensing C, Kennell JC, M, Kroken S, MolnA Zamora J, Farman ML, Selker EU, Salamov A, Shapiro H, Pangilinan J, Lindquist E, Lamers C, Grigoriev IV, Geiser DM, Covert SF, Temporini E, VanEtten HD (2009) The genome of Nectria haematococca: Contribution of supernumerary chromosomes to gene expansion. PLoS Genet 5:e1000618

Collemare J, Pianfetti M, Houlle A-E, Morin D, Camborde L, Gagey M-J, Barbisan C, Fudal I, Lebrun M-H, Bo¨hnert HU (2008) Magnaporthe grisea avirulence gene ACE1 belongs to an infection-specific gene cluster involved in secondary metabolism. New Phytol 179:196–208 Coppin E, Silar P (2007) Identification of PaPKS1, a polyketide synthase involved in melanin formation and its use as a genetic tool in Podospora anserina. Mycol Res 111:901–908 Cox RJ (2007) Polyketides, proteins and genes in fungi: programmed nano-machines begin to reveal their secrets. Org Biomol Chem 5:2010–2026 Cunha MM, Franzen AJ, Alviano DS, Zanardi E, Alviano CS, De Souza W, Rozental S (2005) Inhibition of melanin synthesis pathway by tricyclazole increases susceptibility of Fonsecaea pedrosoi against mouse macrophages. Microsc Res Tech 68:377–384 Cuomo CA, Gu¨ldener U, Xu JR, Trail F, Turgeon BG, Di Pietro A, Walton JD, Ma LJ, Baker SE, Rep M, Adam G, Antoniw J, Baldwin T, Calvo S, Chang YL, Decaprio D, Gale LR, Gnerre S, Goswami RS, HammondKosack K, Harris LJ, Hilburn K, Kennell JC, Kroken S, Magnuson JK, Mannhaupt G, Mauceli E, Mewes HW, Mitterbauer R, Muehlbauer GJ, Mu¨nsterko¨tter M, Nelson D, O’donnell K, Ouellet T, Qi W, Quesneville H, Roncero MI, Seong KY, Tetko IV, M. U, Waalwijk C, Ward TJ, Yao J, Birren BW, Kistler HC (2007) The Fusarium graminearum genome reveals a link between localized polymorphism and pathogen specialization. Science 317:1400–1402 da Silva MB, Marques AF, Nosanchuk JD, Casadevall A, Travassos LR, Taborda CP (2006) Melanin in the dimorphic fungal pathogen Paracoccidioides brasiliensis: effects on phagocytosis, intracellular resistance and drug susceptibility. Microbes Infect 8: 197–205 Dadachova E, Bryan RA, Huang X, Moadel T, Schweitzer AD, Aisen P, Nosanchuk JD, Casadevall A (2007) Ionizing radiation changes the electronic properties of melanin and enhances the growth of melanized fungi. PLoS One 2:e457 Dadachova E, Casadevall A (2008) Ionizing radiation: how fungi cope, adapt, and exploit with the help of melanin. Curr Opin Microbiol 11:525–531 de Jong JC, McCormack BJ, Smirnoff N, Talbot NJ (1997) Glycerol generates turgor in rice blast. Nature 389: 244–244 Dean RA, Talbot NJ, Ebbole DJ, Farman ML, Mitchell TK, Orbach MJ, Thon M, Kulkarni R, Xu J-R, Pan H, Read ND, Lee Y-H, Carbone I, Brown D, Oh YY, Donofrio N, Jeong JS, Soanes DM, Djonovic S, Kolomiets E, Rehmeyer C, Li W, Harding M, Kim S, Lebrun M-H, Bohnert H, Coughlan S, Butler J, Calvo S, Ma L-J, Nicol R, Purcell S, Nusbaum C, Galagan JE, Birren BW (2005) The genome sequence of the rice blast fungus Magnaporthe grisea. Nature 434:980–986 Dighton J, Tugay T, Zhdanova N (2008) Fungi and ionizing radiation from radionuclides. FEMS Microbiol Lett 281:109–120 Ding Y, Williams RM, Sherman DH (2008) Molecular analysis of a 4-dimethylallyltryptophan synthase

Evolution of Genes for Secondary Metabolism in Fungi from Malbranchea aurantiaca. J Biol Chem 283: 16068–16076 Dittmer NT, Kanost MR (2010) Insect multicopper oxidases: diversity, properties, and physiological roles. Insect Biochem Mol Biol 40:179–188 Eichinger L, Pachebat JA, Glockner G, Rajandream MA, Sucgang R, Berriman M, Song J, Olsen R, Szafranski K, Xu Q, Tunggal B, Kummerfeld S, Madera M, Konfortov BA, Rivero F, Bankier AT, Lehmann R, Hamlin N, Davies R, Gaudet P, Fey P, Pilcher K, Chen G, Saunders D, Sodergren E, Davis P, Kerhornou A, Nie X, Hall N, Anjard C, Hemphill L, Bason N, Farbrother P, Desany B, Just E, Morio T, Rost R, Churcher C, Cooper J, Haydock S, van Driessche N, Cronin A, Goodhead I, Muzny D, Mourier T, Pain A, Lu M, Harper D, Lindsay R, Hauser H, James K, Quiles M, Madan Babu M, Saito T, Buchrieser C, Wardroper A, Felder M, Thangavelu M, Johnson D, Knights A, Loulseged H, Mungall K, Oliver K, Price C, Quail MA, Urushihara H, Hernandez J, Rabbinowitsch E, Steffen D, Sanders M, Ma J, Kohara Y, Sharp S, Simmonds M, Spiegler S, Tivey A, Sugano S, White B, Walker D, Woodward J, Winckler T, Tanaka Y, Shaulsky G, Schleicher M, Weinstock G, Rosenthal A, Cox EC, Chisholm RL, Gibbs R, Loomis WF, Platzer M, Kay RR, Williams J, Dear PH, Noegel AA, Barrell B, Kuspa A (2005) The genome of the social amoeba Dictyostelium discoideum. Nature 435: 43–57 Eisendle M, Oberegger H, Zadra I, Haas H (2003) The siderophore system is essential for viability of Aspergillus nidulans: functional analysis of two genes encoding l-ornithine N 5-monooxygenase (sidA) and a non-ribosomal peptide synthetase (sidC). Mol Microbiol 49:359–375 Eisendle M, Schrettl M, Kragl C, Muller D, Illmer P, Haas H (2006) The intracellular siderophore ferricrocin is involved in iron storage, oxidative-stress resistance, germination, and sexual development in Aspergillus nidulans. Eukaryot Cell 5:1596–1603 Elliott CE, Gardiner DM, Thomas G, Cozijnsen A, Van De Wouw A, Howlett BJ (2007) Production of the toxin sirodesmin PL by Leptosphaeria maculans during infection of Brassica napus. Mol Plant Pathol 8: 791–802 Engh I, Nowrousian M, Ku¨ck U (2007) Regulation of melain biosynthesis via the dihydroxynaphtalene pathway is dependent on sexual development in the ascomycete Sordaria macrospora. FEMS Microbiol Lett 275:62–70 Espagne E, Lespinet O, Malagnac F, Da Silva C, Jaillon O, Porcel B, Couloux A, Aury J-M, Segurens B, Poulain J, Anthouard V, Grossetete S, Khalili H, Coppin E, Dequard-Chablat M, Picard M, Contamine V, Arnaise S, Bourdais A, Berteaux-Lecellier V, Gautheret D, de Vries R, Battaglia E, Coutinho P, Danchin E, Henrissat B, Khoury R, Sainsard-Chanet A, Boivin A, Pinan-Lucarre B, Sellem C, Debuchy R, Wincker P, Weissenbach J, Silar P (2008) The genome sequence of the model ascomycete fungus Podospora anserina. Genome Biol 9:R77

247

Esser K (1966) Die Phenoloxidasen des Ascomyceten Podospora anserina. III. Quantitative und qualitative Enzymunterschiede nach Mutation an nicht gekoppelten Loci. Zeitschrift fu¨r Vererbungslehre 97: 327–344 Esser K (1968) Phenol oxidases and morphogenesis in Podospora anserina. Genetics 60:281–288 Fedorova ND, Khaldi N, Joardar VS, Maiti R, Amedeo P, Anderson MJ, Crabtree J, Silva JC, Badger JH, Albarraq A, Angiuoli S, Bussey H, Bowyer P, Cotty PJ, Dyer PS, Egan A, Galens K, Fraser-Liggett CM, Haas BJ, Inman JM, Kent R, Lemieux S, Malavazi I, Orvis J, Roemer T, Ronning CM, Sundaram JP, Sutton G, Turner G, Venter JC, White OR, Whitty BR, Youngman P, Wolfe KH, Goldman GH, Wortman JR, Jiang B, Denning DW, Nierman WC (2008) Genomic islands in the pathogenic filamentous fungus Aspergillus fumigatus. PLOS Genet 4:e1000046 Fernandes M, Keller NP, Adams TH (1998) Sequencespecific binding by Aspergillus nidulans AflR, a C6 zinc cluster protein regulating mycotoxin biosynthesis. Mol Microbiol 28:1355–1365 Flores-Sanchez IJ, Verpoorte R (2009) Plant polyketide synthases: a fascinating group of enzymes. Plant Physiol Biochem 47:167–174 Fox EM, Howlett BJ (2008) Secondary metabolism: regulation and role in fungal biology. Curr Opin Microbiol 11:481–487 Franzen AJ, Cunha MM, Miranda K, Hentschel J, Plattner H, da Silva MB, Salgado CG, de Souza W, Rozental S (2008) Ultrastructural characterization of melanosomes of the human pathogenic fungus Fonsecaea pedrosoi. J Struct Biol 162:75–84 Frases S, Chaskes S, Dadachova E, Casadevall A (2006) Induction by Klebsiella aerogenes of a melanin-like pigment in Cryptococcus neoformans. Appl Environ Microbiol 72:1542–1550 Frases S, Salazar A, Dadachova E, Casadevall A (2007) Cryptococcus neoformans can utilize the bacterial melanin precursor homogentisic acid for fungal melanogenesis. Appl Environ Microbiol 73:615–621 Friesen TL, Stukenbrock EH, Liu Z, Meinhardt S, Ling H, Faris JD, Rasmussen JC, Solomon PS, McDonald BA, Oliver RP (2006) Emergence of a new disease as a result of interspecific virulence gene transfer. Nat Genet 38:953–956 Fujihara N, Sakaguchi A, Tanaka S, Fujii S, Tsuji G, Shiraishi T, O’Connell R, Kubo Y (2010) Peroxisome biogenesis factor PEX13 is required for appressorium-mediated plant infection by the anthracnose fungus Colletotrichum orbiculare. Mol Plant Microbe Interact 23:436–445 Fujii I, Yasuoka Y, Tsai HF, Chang YC, Kwon-Chung KJ, Ebizuka Y (2004) Hydrolytic polyketide shortening by Ayg1p, a novel enzyme involved in fungal melanin biosynthesis. J Biol Chem 279:44613–44620 Fulton TR, Ibrahim N, Losada MC, Grzegorski D, Tkacz JS (1999) A melanin polyketide synthase (PKS) gene from Nodulisporium sp. that shows homology to the pks1 gene of Colletotrichum lagenarium. Mol Gen Genet 262:714–720

248

Ines Teichert and Minou Nowrousian

Funa N, Awakawa T, Horinouchi S (2007) Pentaketide resorcylic acid synthesis by type III polyketide synthase from Neurospora crassa. J Biol Chem 282: 14476–14481 Gachomo EW, Seufferheld MJ, Kotchoni SO (2010) Melanization of appressoria is critical for the pathogenicity of Diplocarpon rosae. Mol Biol Rep DOI 10.1007/s11033-010-0007-4: Gaffoor I, Brown DW, Plattner R, Proctor RH, Qi W, Trail F (2005) Functional analysis of the polyketide synthase genes in the filamentous fungus Gibberella zeae (anamorph Fusarium graminearum). Eukaryot Cell 4:1926–1933 Galagan JE, Calvo SE, Borkovich KA, Selker EU, Read ND, Jaffe D, FitzHugh W, Ma LJ, Smirnov S, Purcell S, Rehman B, Elkins T, Engels R, Wang S, Nielsen CB, Butler J, Endrizzi M, Qui D, Ianakiev P, Bell-Pedersen D, Nelson MA, Werner-Washburne M, Selitrennikoff CP, Kinsey JA, Braun EL, Zelter A, Schulte U, Kothe GO, Jedd G, Mewes W, Staben C, Marcotte E, Greenberg D, Roy A, Foley K, Naylor J, Stange-Thomann N, Barrett R, Gnerre S, Kamal M, Kamvysselis M, Mauceli E, Bielke C, Rudd S, Frishman D, Krystofova S, Rasmussen C, Metzenberg RL, Perkins DD, Kroken S, Cogoni C, Macino G, Catcheside D, Li W, Pratt RJ, Osmani SA, DeSouza CP, Glass L, Orbach MJ, Berglund JA, Voelker R, Yarden O, Plamann M, Seiler S, Dunlap JC, Radford A, Aramayo R, Natvig DO, Alex LA, Mannhaupt G, Ebbole DJ, Freitag M, Paulsen I, Sachs MS, Lander ES, Nusbaum C, Birren B (2003) The genome sequence of the filamentous fungus Neurospora crassa. Nature 422:859–868 Galagan JE, Calvo SE, Cuomo C, Ma L-J, Wortman JR, Batzoglou S, Lee S-I, Basturkmen M, Spevak CC, Clutterbuck J, Kapitonov V, Jurka J, Scazzocchio C, Farman M, Butler J, Purcell S, Harris S, Braus GH, Draht O, Busch S, D’Enfert C, Bouchier C, Goldman GH, Bell-Pedersen D, Griffiths-Jones S, Doonan JH, Yu J, Vienken K, Pain A, Freitag M, Selker EU, Archer DB, Penalva MA, Oakley BR, Momany M, Tanaka T, Kumagai T, Asai K, Machida M, Nierman WC, Denning DW, Caddick M, Hynes M, Paoletti M, Fischer R, Miller B, Dyer P, Sachs MS, Osmani SA, Birren BW (2005) Sequencing of Aspergillus nidulans and comparative analysis with A. fumigatus and A. oryzae. Nature 438:1105–1115 Gallagher RT, White EP, Mortimer PH (1981) Ryegrass staggers: isolation of potent neurotoxins lolitrem A and lolitrem B from staggers-producing pastures. N Z Vet J 29:189–190 Garcia-Vallve´ S, Romeu A, Palau J (2000) Horizontal gene transfer of glycosyl hydrolases of the rumen fungi. Mol Biol Evol 17:352–361 Gardiner DM, Cozijnsen AJ, Wilson LM, Pedras MSC, Howlett BJ (2004) The sirodesmin biosynthetic gene cluster of the plant pathogenic fungus Leptosphaeria maculans. Mol Microbiol 53:1307–1318 Gomez BL, Nosanchuk JD, Diez S, Youngchim S, Aisen P, Cano LE, Restrepo A, Casadevall A, Hamilton AJ (2001) Detection of melanin-like pigments in the dimorphic

fungal pathogen Paracoccidioides brasiliensis in vitro and during infection. Infect Immun 69:5760–5767 Gomez BL, Nosanchuk JD (2003) Melanin and fungi. Curr Opin Infect Dis 16:91–96 Gostincar C, Grube M, de Hoog S, Zalar P, Gunde-Cimerman N (2010) Extremotolerance in fungi: evolution on the edge. FEMS Microbiol Ecol 71:2–11 Haarmann T, Machado C, Lu¨bbe Y, Correira T, Schardl CL, Panaccione DG, Tudzynski P (2005) The ergot alkaloid gene cluster in Claviceps purpurea: extension of the cluster sequence and intra species evolution. Phytochemistry 66:1312–1320 Haarmann T, Lorenz N, Tudzynski P (2008) Use of a nonhomologous end joining deficient strain ([Delta] ku70) of the ergot fungus Claviceps purpurea for identification of a nonribosomal peptide synthetase gene involved in ergotamine biosynthesis. Fungal Genet Biol 45:35–44 Haarmann T, Rolke Y, Giesbert S, Tudzynski P (2009) Ergot: from witchcraft to biotechnology. Mol Plant Pathol 10:563–577 Haas H, Eisendle M, Turgeon BG (2008) Siderophores in fungal physiology and virulence. Annu Rev Phytopathol 46:149–187 Halaouli S, Asther M, Sigoillot JC, Hamdi M, Lomascolo A (2006) Fungal tyrosinases: new prospects in molecular characteristics, bioengineering and biotechnological applications. J Appl Microbiol 100:219–232 Hane JK, Lowe RGT, Solomon PS, Tan K-C, Schoch CL, Spatafora JW, Crous PW, Kodira C, Birren BW, Galagan JE, Torriani SFF, McDonald BA, Oliver RP (2007) Dothideomycete plant interactions illuminated by genome sequencing and EST analysis of the wheat pathogen Stagonospora nodorum. Plant Cell 19: 3347–3368 Hedden P, Kamiya Y (1997) Gibberellin biosynthesis: enzymes, genes and their regulation. Annu Rev Plant Physiol Plant Mol Biol 48:431–460 Henson JM, Butler MJ, Day AW (1999) The dark side of the mycelium: Melanins of Phytopathogenic Fungi. Annu Rev Phytopathol 37:447–471 Herrmann M, Zocher R, Haese A (1996) Effect of disruption of the enniatin synthetase gene on the virulence of Fusarium avenaceum. Mol Plant Microbe Interact 9:226–232 Hirsch HM (1954) Environmental factors influencing the differentiation of protoperithecia and their relation to tyrosinase and melanin formation in Neurospora crassa. Physiologia Plantarum 7:72–97 Hoffmeister D, Keller NP (2007) Natural products of filamentous fungi: enzymes, genes, and their regulation. Nat Prod Rep 24:393–416 Hohn TM, Beremand PD (1989) Isolation and nucleotide sequence of a sesquiterpene cyclase gene from the trichothecene-producing fungus Fusarium sporotrichioides. Gene 79:131–138 Hohn TM, McCormick SP, Desjardins AE (1993) Evidence for a gene cluster involving trichothecene-pathway biosynthetic genes in Fusarium sporotrichioides. Curr Genet 24:291–295

Evolution of Genes for Secondary Metabolism in Fungi Horowitz NH, Charlang G, Horn G, Williams NP (1976) Isolation and identification of the conidial germination factor of Neurospora crassa. J Bacteriol 127: 135–140 Howard RJ, Ferrari MA (1989) Role of melanin in appressorium formation. Exp Mycol 13:403–418 Howard RJ, Ferrari MA, Roach DH, Money NP (1991) Penetration of hard substrates by a fungus employing enormous turgor pressures. Proc Nat Acad Sci USA 88:11281–11284 Howard RJ, Valent B (1996) Breaking and entering: host penetration by the fungal rice blast pathogen Magnaporthe grisea. Annu Rev Microbiol 50:491–512 Inderbitzin P, Asvarak T, Turgeon BG (2010) Six new genes required for production of T-toxin, a polyketide determinant of high virulence of Cochliobolus heterostrophus to maize. Mol Plant Microbe Interact 23:458–472 Jacobson ES (2000) Pathogenic roles for fungal melanins. Clin Microbiol Rev 13:708–717 Jaenicke E, Decker H (2004) Functional changes in the family of type 3 copper proteins during evolution. Chembiochem 5:163–169 Jahn B, Koch A, Schmidt A, Wanner G, Gehringer H, Bhakdi S, Brakhage AA (1997) Isolation and characterization of a pigmentless–conidium mutant of Aspergillus fumigatus with altered conidial surface and reduced virulence. Infect Immun 65:5110–5117 Jahn B, Langfelder K, Schneider U, Schindel C, Brakhage AA (2002) PKSP-dependent reduction of phagolysosome fusion and intracellular kill of Aspergillus fumigatus conidia by human monocyte-derived macrophages. Cell Microbiol 4:793–803 Johnson RD, Johnson L, Itoh Y, Kodama M, Otani H, Kohmoto K (2000) Cloning and characterization of a cyclic peptide synthetase gene from Alternaria alternata apple pathotype whose product is involved in AM-toxin synthesis and pathogenicity. Mol Plant Microbe Interact 13:742–753 Ka¨mper J, Kahmann R, Bo¨lker M, Ma LJ, Brefort T, Saville BJ, Banuett F, Kronstad JW, Gold SE, Mu¨ller O, Perlin MH, Wo¨sten HA, de V, R., Ruiz-Herrera J, ReynagaPena CG, Snetselaar K, McCann M, Pe´rez-Martin J, Feldbru¨gge M, Basse CW, Steinberg G, Ibeas JI, Holloman W, Guzman P, Farman M, Stajich JE, Sentandreu R, Gonzlez-Prieto JM, Kennell JC, Molina L, Schirawski J, Mendoza-Mendoza A, Greilinger D, Mu¨nch K, Ro¨ssel N, Scherer M, Vranes M, Ladendorf O, Vincon V, Fuchs U, Sandrock B, Meng S, Ho EC, Cahill MJ, Boyce K, Klose J, Klosterman SJ, Deelstra HJ, Ortiz-Castellanos L, Li W, Sanchez-Alonso P, Schreier PH, Ha¨user-Hahn I, Vaupel M, Koopmann E, Friedrich G, Voss H, Schlu¨ter T, Margolis J, Platt D, Swimmer C, Gnirke A, Chen F, Vysotskaia V, Mannhaupt G, Gu¨ldener U, Mu¨nsterko¨tter M, Haase D, Oesterheld M, Mewes HW, Mauceli EW, DeCaprio D, Wade CM, Butler J, Young S, Jaffe DB, Calvo S, Nusbaum C, Galagan J, Birren BW (2006) Insights from the genome of the biotrophic fungal plant pathogen Ustilago maydis. Nature 444:97–101

249

Karkowska-Kuleta J, Rapala-Kozik M, Kozik A (2009) Fungi pathogenic to humans: molecular bases of virulence of Candida albicans, Cryptococcus neoformans and Aspergillus fumigatus. Acta Biochim Pol 56: 211–224 Kawaide H, Imai R, Sassa T, Kamiya Y (1997) Ent-kaurene synthase from the fungus Phaeosphaeria sp. L487. cDNA isolation, characterization, and bacterial expression of a bifunctional diterpene cyclase in fungal gibberellin biosynthesis. J Biol Chem 272: 21706–21712 Kawamura C, Moriwaki J, Kimura N, Fujita Y, Fuji S, Hirano T, Koizumi S, Tsuge T (1997) The melanin biosynthesis genes of Alternaria alternata can restore pathogenicity of the melanin-deficient mutants of Magnaporthe grisea. Mol Plant Microbe Interact 10: 446–453 Kawamura C, Tsujimoto T, Tsuge T (1999) Targeted disruption of a melanin biosynthesis gene affects conidial development and UV tolerance in the Japanese pear pathotype of Alternaria alternata. Mol Plant Microbe Interact 12:59–63 Keller NP, Hohn TM (1997) Metabolic pathway gene clusters in filamentous fungi. Fungal Genet Biol 21:17–29 Keller NP, Turner G, Bennett JW (2005) Fungal secondary metabolism – from biochemistry to genomics. Nat Rev Microbiol 3:937–947 Khaldi N, Collemare J, Lebrun M-H, Wolfe KH (2008) Evidence for horizontal transfer of a secondary metabolite gene cluster between fungi. Genome Biol 9:R18 Khaldi N, Wolfe KH (2008) Elusive origins of the extra genes in Aspergillus oryzae. PLOS One 3:e3036 Kihara J, Moriwaki A, Ito M, Arase S, Honda Y (2004) Expression of THR1, a 1,3,8-trihydroxynaphthalene reductase gene involved in melanin biosynthesis in the phytopathogenic fungus Bipolaris oryzae, is enhanced by near-ultraviolet radiation. Pigment Cell Res 17:15–23 Kim KH, Cho Y, La Rota M, Cramer Jr. RA, Lawrence CB (2007) Functional analysis of the Alternaria brassicicola non–ribosomal peptide synthetase gene AbNPS2 reveals a role in conidial cell wall construction. Mol Plant Pathol 8:23–39 Kimura M, Tokai T, O’Donnell K, Ward TJ, Fujimura M, Hamamoto H, Shibata T, Yamaguchi I (2003) The trichothecene biosynthesis gene cluster of Fusarium graminearum F15 contains a limited number of essential pathway genes and expressed non-essential genes. FEBS Lett 539:105–110 Kimura M, Tokai T, Takahashi-Ando N, Ohsato S, Fujimura M (2007) Molecular and genetic studies of Fusarium trichothecene biosynthesis: pathways, genes, and evolution. Biosci Biotechnol Biochem 71: 2105–2123 Kimura N, Tsuge T (1993) Gene cluster involved in melanin biosynthesis of the filamentous fungus Alternaria alternata. J Bacteriol 175:4427–4435 Kodama M, Rose MS, Yang G, Yun SH, Yoder OC, Turgeon BG (1999) The translocation-associated tox1 locus of

250

Ines Teichert and Minou Nowrousian

Cochliobolus heterostrophus is two genetic elements on two different chromosomes. Genetics 151:585–596 Kono Y, Takeuchi S, Kawarada A, Daly JM, Knoche HW (1980) Studies on the host-specific pathotoxins produced by Helminthosporium maydis, race T. Agric Biol Chem 44:2613–2622 Koschorreck K, Schmid RD, Urlacher VB (2009) Improving the functional expression of a Bacillus licheniformis laccase by random and site-directed mutagenesis. BMC Biotechnol 9:12 Kroken S, Glass NL, Taylor JW, Yoder OC, Turgeon BG (2003) Phylogenomic analyis of type I polyketide synthase genes in pathogenic and saprobic ascomycetes. Proc Nat Acad Sci USA 100:15670–15675 Kubo Y, Suzuki K, Furusawa I, Ishida N, Yamamoto M (1982) Relation of appressorium pigmentation and penetration of nitrocellulose membranes by Colletotrichum lagenarium. Phytopathol 72:498–501 Kubo Y, Nakamura H, Kobayashi K, Okuno T, Furusawa I (1991) Cloning of a melanin biosynthetic gene essential for appressorial penetration of Colletotrichum lagenarium. Mol Plant Microbe Interact 4:440–445 Kubo Y, Takano Y, Endo N, Yasuda N, Tajima S, Furusawa I (1996) Cloning and structural analysis of the melanin biosynthesis gene SCD1 encoding scytalone dehydratase in Colletotrichum lagenarium. Appl Environ Microbiol 62:4340–4344 Landan G, Cohen G, Aharonowitz Y, Shuali Y, Graur D, Shiffman D (1990) Evolution of isopenicillin N synthase genes may have involved horizontal gene transfer. Mol Biol Evol 7:399–406 Langfelder K, Jahn B, Gehringer H, Schmidt A, Wanner G, Brakhage AA (1998) Identification of a polyketide synthase gene (pksP) of Aspergillus fumigatus involved in conidial pigment biosynthesis and virulence. Med Microbiol Immunol 187:79–89 Langfelder K, Streibel M, Jahn B, Haase G, Brakhage AA (2003) Biosynthesis of fungal melanins and their importance for human pathogenic fungi. Fungal Genet Biol 38:143–158 Lee BN, Kroken S, Chou DYT, Robbertse B, Yoder OC, Turgeon BG (2005) Functional analysis of all nonribosomal peptide synthetases in Cochliobolus heterostrophus reveals a factor, NPS6, involved in virulence and resistance to oxidative stress. Eukaryot Cell 4: 545–555 Liu GY, Nizet V (2009) Color me bad: microbial pigments as virulence factors. Trends Microbiol 17:406–413 Lorenz N, Haarmann T, Pazoutov S, Jung M, Tudzynski P (2009) The ergot alkaloid gene cluster: Functional analyses and evolutionary aspects. Phytochemistry 70:1822–1832 Machida M, Asai K, Sano M, Tanaka T, Kumagai T, Terai G, Kusumoto K-I, Arima T, Akita O, Kashiwagi Y, Abe K, Gomi K, Horiuchi H, Kitamoto K, Kobayashi T, Takeuchi M, Denning DW, Galagan JE, Nierman WC, Yu J, Archer DB, Bennett JW, Bhatnagar D, Cleveland TE, Fedorova ND, Gotoh O, Horikawa H, Hosoyama A, Ichinomiya M, Igarashi R, Iwashita K, Juvvadi PR, Kato M, Kato Y, Kin T, Kokubun A, Maeda H, Maeyama N, Maruyama J-i, Nagasaki H,

Nakajima T, Oda K, Okada K, Paulsen I, Sakamoto K, Sawano T, Takahashi M, Takase K, Terabayashi Y, Wortman JR, Yamada O, Yamagata Y, Anazawa H, Hata Y, Koide Y, Komori T, Koyama Y, Minetoki T, Suharnan S, Tanaka A, Isono K, Kuhara S, Ogasawara N, Kikuchi H (2005) Genome sequencing and analysis of Aspergillus oryzae. Nature 438:1157–1161 Martin F, Aerts A, Ahren D, Brun A, Danchin EGJ, Duchaussoy F, Gibon J, Kohler A, Lindquist E, Pereda V, Salamov A, Shapiro HJ, Wuyts J, Blaudez D, Buee M, Brokstein P, Canback B, Cohen D, Courty PE, Coutinho PM, Delaruelle C, Detter JC, Deveau A, DiFazio S, Duplessis S, Fraissinet-Tachet L, Lucic E, Frey-Klett P, Fourrey C, Feussner I, Gay G, Grimwood J, Hoegger PJ, Jain P, Kilaru S, Labbe J, Lin YC, Legue V, Le Tacon F, Marmeisse R, Melayah D, Montanini B, Muratet M, Nehls U, Niculita-Hirzel H, Secq MPO-L, Peter M, Quesneville H, Rajashekar B, Reich M, Rouhier N, Schmutz J, Yin T, Chalot M, Henrissat B, Kues U, Lucas S, Van de Peer Y, Podila GK, Polle A, Pukkila PJ, Richardson PM, Rouze P, Sanders IR, Stajich JE, Tunlid A, Tuskan G, Grigoriev IV (2008) The genome of Laccaria bicolor provides insights into mycorrhizal symbiosis. Nature 452:88–92 Martin F, Kohler A, Murat C, Balestrini R, Coutinho PM, Jaillon O, Montanini B, Morin E, Noel B, Percudani R, Porcel B, Rubini A, Amicucci A, Amselem J, Anthouard V, Arcioni S, Artiguenave F, Aury J-M, Ballario P, Bolchi A, Brenna A, Brun A, Buee M, Cantarel B, Chevalier G, Couloux A, Da Silva C, Denoeud F, Duplessis S, Ghignone S, Hilselberger B, Iotti M, Marcais B, Mello A, Miranda M, Pacioni G, Quesneville H, Riccioni C, Ruotolo R, Splivallo R, Stocchi V, Tisserant E, Viscomi AR, Zambonelli A, Zampieri E, Henrissat B, Lebrun M-H, Paolocci F, Bonfante P, Ottonello S, Wincker P (2010) Pe´rigord black truffle genome uncovers evolutionary origins and mechanisms of symbiosis. Nature 464:1033–1039 Martinez D, Challacombe J, Morgenstern I, Hibbett D, Schmoll M, Kubicek CP, Ferreira P, Ruiz-Duenas FJ, Martinez AT, Kersten P, Hammel KE, Vanden Wymelenberg A, Gaskell J, Lindquist E, Sabat G, Splinter BonDurant S, Larrondo LF, Canessa P, Vicuna R, Yadav J, Doddapaneni H, Subramanian V, Pisabarro ˜ n JL, Oguiza JA, Master E, Henrissat B, AG, LavA Coutinho PM, Harris P, Magnuson JK, Baker SE, ˜ ¼es U, Ramaiya Bruno K, Kenealy W, Hoegger PJ, KA P, Lucas S, Salamov A, Shapiro H, Tu H, Chee CL, Misra M, Xie G, Teter S, Yaver D, James T, Mokrejs M, Pospisek M, Grigoriev IV, Brettin T, Rokhsar D, Berka R, Cullen D (2009) Genome, transcriptome, and secretome analysis of wood decay fungus Postia placenta supports unique mechanisms of lignocellulose conversion. Proc Nat Acad Sci USA 106:1954–1959 Matzanke BF, Bill E, Trautwein AX, Winkelmann G (1987) Role of siderophores in iron storage in spores of Neurospora crassa and Aspergillus ochraceus. J Bacteriol 169:5873–5876 Mendgen K, Hahn M, Deising H (1996) Morphogenesis and mechanisms of penetration by plant pathogenic fungi. Annu Rev Phytopathol 34:367–386

Evolution of Genes for Secondary Metabolism in Fungi Metzger U, Schall C, Zocher G, Unso¨ld I, Stec E, Li S-M, Heide L, Stehle T (2009) The structure of dimethylallyl tryptophan synthase reveals a common architecture of aromatic prenyltransferases in fungi and bacteria. Proc Nat Acad Sci USA 106:14309–14314 Mironenko NV, Alekhina IA, Zhdanova NN, Bulat SA (2000) Intraspecific variation in gamma-radiation resistance and genomic structure in the filamentous fungus Alternaria alternata: a case study of strains inhabiting Chernobyl reactor no. 4. Ecotoxicol Environ Saf 45:177–187 Misiek M, Hoffmeister D (2007) Fungal genetics, genomics, and secondary metabolites in pharmaceutical sciences. Planta Med 73:103–115 Money NP, Howard RJ (1996) Confirmation of a link between fungal pigmentation, turgor pressure, and pathogenicity using a new method of turgor measurement. Fungal Genet Biol 20:217–227 Morris-Jones R, Youngchim S, Hextall JM, Gomez BL, Morris-Jones SD, Hay RJ, Casadevall A, Nosanchuk JD, Hamilton AJ (2004) Scytalidium dimidiatum causing recalcitrant subcutaneous lesions produces melanin. J Clin Microbiol 42:3789–3794 Morris-Jones R, Gomez BL, Diez S, Uran M, Morris-Jones SD, Casadevall A, Nosanchuk JD, Hamilton AJ (2005) Synthesis of melanin pigment by Candida albicans in vitro and during infection. Infect Immun 73: 6147–6150 Motoyama T, Imanishi K, Yamaguchi I (1998) cDNA cloning, expression, and mutagenesis of scytalone dehydratase needed for pathogenicity of the rice blast fungus, Pyricularia oryzae. Biosci Biotechnol Biochem 62:564–566 Nicholson MJ, Koulman A, Monahan BJ, Pritchard BL, Payne GA, Scott B (2009) Identification of two aflatrem biosynthesis gene loci in Aspergillus flavus and metabolic engineering of Penicillium paxilli to elucidate their function. Appl Env Microbiol 75: 7469–7481 Nierman WC, Pain A, Anderson MJ, Wortman JR, Kim HS, Arroyo J, Berriman M, Abe K, Archer DB, Bermejo C, Bennett J, Bowyer P, Chen D, Collins M, Coulsen R, Davies R, Dyer PS, Farman M, Fedorova N, Fedorova N, Feldblyum TV, Fischer R, Fosker N, Fraser A, Garcia JL, Garcia MJ, Goble A, Goldman GH, Gomi K, Griffith-Jones S, Gwilliam R, Haas B, Haas H, Harris D, Horiuchi H, Huang J, Humphray S, Jimenez J, Keller N, Khouri H, Kitamoto K, Kobayashi T, Konzack S, Kulkarni R, Kumagai T, Lafton A, Latge J-P, Li W, Lord A, Lu C, Majoros WH, May GS, Miller BL, Mohamoud Y, Molina M, Monod M, Mouyna I, Mulligan S, Murphy L, O’Neil S, Paulsen I, Penalva MA, Pertea M, Price C, Pritchard BL, Quail MA, Rabbinowitsch E, Rawlins N, Rajandream M-A, Reichard U, Renauld H, Robson GD, de Cordoba SR, Rodriguez-Pena JM, Ronning CM, Rutter S, Salzberg SL, Sanchez M, Sanchez-Ferrero JC, Saunders D, Seeger K, Squares R, Squares S, Takeuchi M, Tekaia F, Turner G, de Aldana CRV, Weidman J, White O, Woodward J, Yu J-H, Fraser C, Galagan JE, Asai K, Machida M, Hall N, Barrell B, Denning DW (2005) Genomic sequence of the pathogenic and allergenic

251

filamentous fungus Aspergillus fumigatus. Nature 438: 1151–1156 Nosanchuk JD, Gomez BL, Youngchim S, Diez S, Aisen P, Zancope-Oliveira RM, Restrepo A, Casadevall A, Hamilton AJ (2002) Histoplasma capsulatum synthesizes melanin-like pigments in vitro and during mammalian infection. Infect Immun 70:5124–5131 Nosanchuk JD, Casadevall A (2003) The contribution of melanin to microbial pathogenesis. Cell Microbiol 5: 203–223 Nosanchuk JD, van Duin D, Mandal P, Aisen P, Legendre AM, Casadevall A (2004) Blastomyces dermatitidis produces melanin in vitro and during infection. FEMS Microbiol Lett 239:187–193 Nosanchuk JD, Casadevall A (2006) Impact of melanin on microbial virulence and clinical resistance to antimicrobial compounds. Antimicrob Agents Chemother 50: 3519–3528 Nosanchuk JD, Yu JJ, Hung CY, Casadevall A, Cole GT (2007) Coccidioides posadasii produces melanin in vitro and during infection. Fungal Genet Biol 44: 517–520 Nowrousian M (2009) A novel polyketide biosynthesis gene cluster is involved in fruiting body morphogenesis in the filamentous fungi Sordaria macrospora and Neurospora crassa. Curr Genet 55:185–198 Nowrousian M, Stajich JE, Chu M, Engh I, Espagne E, Halliday K, Kamerewerd J, Kempken F, Knab B, Kuo HC, Osiewacz HD, Po¨ggeler S, Read ND, Seiler S, Smith KM, Zickler D, Ku¨ck U, Freitag M (2010) De novo assembly of a 40 Mb eukaryotic genome from short sequence reads: Sordaria macrospora, a model organism for fungal morphogenesis. PLoS Genet 6: e1000891 Oide S, Moeder W, Krasnoff S, Gibson D, Haas H, Yoshioka K, Turgeon BG (2006) NPS6, encoding a nonribosomal peptide synthetase involved in siderophore-mediated iron metabolism, is a conserved virulence determinant of plant pathogenic ascomycetes. Plant Cell 18:2836–2853 Oide S, Krasnoff SB, Gibson DM, Turgeon BG (2007) Intracellular siderophores are essential for ascomycete sexual development in heterothallic Cochliobolus heterostrophus and homothallic Gibberella zeae. Eukaryot Cell 6:1339–1353 Ozeki H, Ito S, Wakamatsu K, Ishiguro I (1997a) Chemical characterization of pheomelanogenesis starting from dihydroxyphenylalanine or tyrosine and cysteine. Effects of tyrosinase and cysteine concentrations and reaction time. Biochim Biophys Acta 1336: 539–548 Ozeki H, Wakamatsu K, Ito S, Ishiguro I (1997b) Chemical characterization of eumelanins with special emphasis on 5,6-dihydroxyindole-2-carboxylic acid content and molecular size. Anal Biochem 248:149–157 Panaccione DG, Scott-Craig JS, Pocard JA, Walton JD (1992) A cyclic peptide synthetase gene required for pathogenicity of the fungus Cochliobolus carbonum on maize. Proc Nat Acad Sci USA 89:6590–6594 Panaccione DG, Johnson RD, Wang J, Young CA, Damrongkool P, Scott B, Schardl CL (2001) Elimination of ergovaline from a grass–Neotyphodium endophyte

252

Ines Teichert and Minou Nowrousian

symbiosis by genetic modification of the endophyte. Proc Nat Acad Sci USA 98:12820–12825 Patron NJ, Waller RF, Cozijnsen AJ, Straney DC, Gardiner DM, Nierman WC, Howlett BJ (2007) Origin and distribution of epipolythiodioxopiperazine (ETP) gene clusters in filamentous ascomycetes. BMC Evol Biol 7:174 Pel HJ, de Winde JH, Archer DB, Dyer PS, Hofmann G, Schaap PJ, Turner G, de Vries RP, Albang R, Albermann K, Andersen MR, Bendtsen JD, Benen JAE, van den Berg M, Breestraat S, Caddick MX, Contreras R, Cornell M, Coutinho PM, Danchin EGJ, Debets AJM, Dekker P, van Dijck PWM, van Dijk A, Dijkhuizen L, Driessen AJM, d’Enfert C, Geysens S, Goosen C, Groot GSP, de Groot PWJ, Guillemette T, Henrissat B, Herweijer M, van den Hombergh JPTW, van den Hondel CAMJJ, van der Heijden RTJM, van der Kaaij RM, Klis FM, Kools HJ, Kubicek CP, van Kuyk PA, Lauber J, Lu X, van der Maarel MJEC, Meulenberg R, Menke H, Mortimer MA, Nielsen J, Oliver SG, Olsthoorn M, Pal K, van Peij NNME, Ram AFJ, Rinas U, Roubos JA, Sagt CMJ, Schmoll M, Sun J, Ussery D, Varga J, Vervecken W, van de Vondervoort PJJ, Wedler H, Wosten HAB, Zeng A-P, van Ooyen AJJ, Visser J, Stam H (2007) Genome sequencing and analysis of the versatile cell factory Aspergillus niger CBS 513.88. Nat Biotech 25:221–231 Penalva MA, Moya A, Dopazo J, Ramo´n D (1990) Sequences of isopenicillin N synthetase genes suggest horizontal gene transfer from prokaryotes to eukaryotes. Proc Biol Sci 241:164–169 Perpetua NS, Kubo Y, Takano Y, Furusawa I (1996) Cloning and characterization of a melanin biosynthetic THR1 reductase gene essential for appressorial penetration of Colletotrichum lagenarium. Mol Plant Microbe Interact 9:323–329 Perrin RM, Fedorova ND, Bok JW, Cramer RA, Wortman JR, Kim HS, Nierman WC, Keller NP (2007) Transcriptional Regulation of Chemical Diversity in Aspergillus fumigatus by LaeA. PLoS Pathog 3:e50 Pihet M, Vandeputte P, Tronchin G, Renier G, Saulnier P, Georgeault S, Mallet R, Chabasse D, Symoens F, Bouchara JP (2009) Melanin is an essential component for the integrity of the cell wall of Aspergillus fumigatus conidia. BMB Microbiol 9:177 Polacheck I, Kwon-Chung KJ (1988) Melanogenesis in Cryptococcus neoformans. J Gen Microbiol 134: 1037–1041 Prade RA, Cruz AK, Terenzi HF (1984) Regulation of tyrosinase during the vegetative and sexual life-cycles of Neurospora crassa. Archives of Microbiology 140: 236–242 Proctor RH, Hohn TM, McCormick SP (1995) Reduced virulence of Gibberella zeae caused by disruption of a trichothecene toxin biosynthetic gene. Mol Plant Microbe Interact 8:593–601 Proctor RH, McCormick SP, Alexander NJ, Desjardins AE (2009) Evidence that a secondary metabolic biosynthetic gene cluster has grown by gene relocation during evolution of the filamentous fungus Fusarium. Mol Microbiol 74:1128–1142

Ragnelli AM, Pacioni G, Aimola P, Lanza B, Miranda M (1992) Truffle melanogenesis: correlation with reproductive differentiation and ascocarp ripening. Pigment Cell Research 5:205–212 Ranocha P, Chabannes M, Chamayou S, Danoun S, Jauneau A, Boudet AM, Goffner D (2002) Laccase downregulation causes alterations in phenolic metabolism and cell wall structure in poplar. Plant Physiol 129: 145–155 Reverberi M, Ricelli A, Zjalic S, Fabbri AA, Fanelli C (2010) Natural functions of mycotoxins and control of their biosynthesis in fungi. Appl Microbiol Biotechnol 87: 899–911 Reyes-Dominguez Y, Bok JW, Berger H, Shwab EK, Basheer A, Gallmetzer A, Scazzocchio C, Keller N, Strauss J (2010) Heterochromatic marks are associated with the repression of secondary metabolism clusters in Aspergillus nidulans. Mol Microbiol in press: Riley PA (1997) Melanin. Int J Biochem Cell Biol 29: 1235–1239 Rohlfs M, Albert M, Keller NP, Kempken F (2007) Secondary chemicals protect mould from fungivory. Biol Lett 3:523–525 Romero-Martinez R, Wheeler M, Guerrero-Plata A, Rico G, Torres-Guerrero H (2000) Biosynthesis and functions of melanin in Sporothrix schenckii. Infect Immun 68:3696–3703 Rose MS, Yun S-H, Asvarak T, Lu S-W, Yoder OC, Turgeon BG (2002) A decarboxylase encoded at the Cochliobolus heterostrophus translocation-associated Tox1B locus is required for polyketide (T-toxin) biosynthesis and high virulence on T-cytoplasm maize. Mol Plant Microbe Interact 15:883–893 Rosewich UL, Kistler HC (2000) Role of horizontal gene transfer in the evolution of fungi. Annu Rev Phytopathol 38:326–363 Saika S, Nicholson MJ, Young C, Parker EJ, Scott B (2008) The genetic basis for indole-diterpene chemical diversity in filamentous fungi. Mycol Res 112:184–199 Saikia S, Parker EJ, Koulman A, Scott B (2006) Four gene products are required for the fungal synthesis of the indole-diterpene, paspaline. FEBS Lett 580:1625–1630 Sanchez-Ferrer A, Rodriguez-Lopez JN, Garcia-Canovas F, Garcia-Carmona F (1995) Tyrosinase: a comprehensive review of its mechanism. Biochim Biophys Acta 1247:1–11 Sandmann G (1994) Carotenoid biosynthesis in microorganisms and plants. Eur J Biochem 223:7–24 Schardl CL (2001) Epichloe¨ festucae and related mutualistic symbionts of grasses. Fungal Genet Biol 33:69–82 Schardl CL, Panaccione DG, Tudzynski P (2006) Ergot alkaloids – biology and molecular biology. Alkaloids Chem Biol 63:45–86 Schimek C, Wo¨stermeyer J (2009) Carotene derivatives in sexual communictaion of zygomycete fungi. Phytochemistry 70:1867–1875 Schrettl M, Winkelmann G, Haas H (2004) Ferrichrome in Schizosaccharomyces pombe – an iron transport and iron storage compound. Biometals 17:647–654 Schrettl M, Bignell E, Kragl C, Sabiha Y, Loss O, Eisendle M, Wallner A, Arst HN, Jr., Haynes K, Haas H (2007)

Evolution of Genes for Secondary Metabolism in Fungi Distinct roles for intra- and extracellular siderophores during Aspergillus fumigatus infection. PLoS Pathog 3:e128 Seshime Y, Juvvadi PR, Fujii I, Kitamoto K (2005) Discovery of a novel superfamily of type III polyketide synthases in Aspergillus oryzae. Biochem Biophys Res Commun 331:253–260 Shaaban M, Palmer JM, El-Naggar WA, El-Sokkary MA, Habib ELSE, Keller NP (2010) Involvement of transposon-like elements in penicillin gene cluster regulation. Fungal Genet Biol 47:423–432 Shwab EK, Bok JW, Tribus M, Galehr J, Graessle S, Keller NP (2007) Histone deacetylase activity regulates chemical diversity in Aspergillus. Eukaryot Cell 6: 1656–1664 Singaravelan N, Grishkan I, Beharav A, Wakamatsu K, Ito S, Nevo E (2008) Adaptive melanin response of the soil fungus Aspergillus niger to UV radiation stress at “Evolution Canyon”, Mount Carmel, Israel. PLoS One 3:e2993 Soler-Rivas C, Jolivet S, Arpin N, Olivier JM, Wichers HJ (1999) Biochemical and physiological aspects of brown blotch disease of Agaricus bisporus. FEMS Microbiol Rev 23:591–614 Song Z, Cox RJ, Lazarus CM, Simpson TJ (2004) Fusarin C biosynthesis in Fusarium moniliforme and Fusarium venenatum. Chembiochem 5:1196–1203 Spiering MJ, Moon CD, Wilkinson HH, Schardl CL (2005) Gene clusters for insecticidal loline alkaloids in the grass-endophytic fungus Neotyphodium uncinatum. Genetics 169:1403–1414 Starratt AN, Ross LM, Lazarovits G (2002) 1,8-Dihydroxynaphthalene monoglucoside, a new metabolite of Sclerotinia sclerotiorum, and the effect of tricyclazole on its production. Can J Microbiol 48:320–325 Staunton J, Weissman KJ (2001) Polyketide biosynthesis: a millenium review. Nat Prod Rep 18:380–416 Steffan N, Grundmann A, Yin WB, Kremer A, Li SM (2009) Indole prenyltransferases from fungi: a new enzyme group with high potential for the production of prenylated indole derivatives. Curr Med Chem 16: 218–231 Stipanovic RD, Bell AA (1976) Pentaketide metabolites of Verticillium dahliae. 3. Identification of ()-3, 4-dihydro-3,8-dihydroxy-1(2h)-naphtalenone[()vermelone] as a precursor to melanin. J Org Chem 41: 2468–2469 Sturley SL (2000) Conservation of eukaryotic sterol homeostasis: new insights from studies in budding yeast. Biochim Biophys Acta 1529:155–163 Stu¨ssi H, Rast DM (1981) The biosynthesis and possible function of gamma-glutaminyl-4-hydroxybenzene in Agaricus bisporus. Phytochemistry 20:2347–2352 Sugareva V, Hartl A, Brock M, Hubner K, Rohde M, Heinekamp T, Brakhage AA (2006) Characterisation of the laccase-encoding gene abr2 of the dihydroxynaphthalene-like melanin gene cluster of Aspergillus fumigatus. Arch Microbiol 186:345–355 Suryanarayanan TS, Ravishankar JP, Venkatesan G, Murali TS (2004) Characterization of the melanin

253

pigment of a cosmopolitan fungal endophyte. Mycol Res 108:974–978 Taborda CP, da Silva MB, Nosanchuk JD, Travassos LR (2008) Melanin as a virulence factor of Paracoccidioides brasiliensis and other dimorphic pathogenic fungi: a minireview. Mycopathologia 165:331–339 Takano Y, Kubo Y, Shimizu K, Mise K, Okuno T, Furusawa I (1995) Structural analysis of PKS1, a polyketide synthase gene involved in melanin biosynthesis in Colletotrichum lagenarium. Mol Gen Genet 249:162–167 Takano Y, Kubo Y, Kawamura C, Tsuge T, Furusawa I (1997) The Alternaria alternata melanin biosynthesis gene restores appressorial melanization and penetration of cellulose membranes in the melanin-deficient albino mutant of Colletotrichum lagenarium. Fungal Genet Biol 21:131–140 Tanabe K, Nishimura S, Kohmoto K (1988) Pathogenicity of pectic enzymes and melanin-deficient mutants of Alternaria alternata Japanese pear pathotype. Ann Phytopath Soc Jpn 54:364 Tanaka A, Tapper BA, Popay A, Parker EJ, Scott B (2005) A symbiosis expressed non-ribosomal peptide synthetase from a mutualistic fungal endophyte of perennial ryegrass confers protection to the symbiotum from insect herbivory. Mol Microbiol 57:1036–1050 Tanguay P, Bozza S, Breuil C (2006) Assessing RNAi frequency and efficiency in Ophiostoma floccosum and O. piceae. Fungal Genet Biol 43:804–812 Taylor BE, Wheeler MH, Szaniszlo PJ (1987) Evidence for pentaketide melanin biosynthesis in dematiaceous human pathogenic fungi. Mycologia 79:320–322 Thompson JE, Fahnestock S, Farrall L, Liao DI, Valent B, Jordan DB (2000) The second naphthol reductase of fungal melanin biosynthesis in Magnaporthe grisea – Tetrahydroxynaphthalene reductase. J Biol Chem 275: 34867–34872 Thurston CF (1994) The structure and function of fungal laccases. Microbiology 140:19–26 Tokousbalides MC, Sisler HD (1979) Site of inhibition by tricyclazole in the melanin biosynthetic pathway of Verticillium dahliae. Pest Biochem Phys 11:64–73 Tsai HF, Wang H, Gebler JC, Poulter CD, Schardl CL (1995) The Claviceps purpurea gene encoding dimethylallyltryptophan synthase, the commited step for ergot alkaloid biosynthesis. Biochem Biophys Res Commun 216:119–125 Tsai HF, Chang YC, Washburn RG, Wheeler MH, KwonChung KJ (1998) The developmentally regulated alb1 gene of Aspergillus fumigatus: its role in modulation of conidial morphology and virulence. J Bacteriol 180: 3031–3038 Tsai HF, Wheeler MH, Chang YC, Kwon-Chung KJ (1999) A developmentally regulated gene cluster involved in conidial pigment biosynthesis in Aspergillus fumigatus. J Bacteriol 181:6469–6477 Tsai HF, Fujii I, Watanabe A, Wheeler MH, Chang YC, Yasuoka Y, Ebizuka Y, Kwon-Chung KJ (2001) Pentaketide melanin biosynthesis in Aspergillus fumigatus requires chain-length shortening of a heptaketide precursor. J Biol Chem 276:29292–29298

254

Ines Teichert and Minou Nowrousian

Tudzynski B, Ho¨lter K (1998) Gibberellin biosynthetic pathway in Gibberella fujikuroi: evidence for a gene cluster. Fungal Genet Biol 25:157–170 Tudzynski B, Kawaide H, Kamiya Y (1998) Gibberellin biosynthesis in Gibberella fujikuroi: cloning and characterization of the copalyl diphosphate synthase gene. Curr Genet 34:234–240 Tudzynski B (2005) Gibberellin biosynthesis in fungi: genes, enzymes, evolution and impact on biotechnology. Appl Microbiol Biotechnol 66:597–611 Tudzynski P, Ho¨lter K, Correira T, Arntz C, Grammel N, Keller U (1999) Evidence for an ergot alkaloid gene cluster in Claviceps purpurea. Mol Gen Genet 261: 133–141 Unso¨ld IA, Li S-M (2005) Overproduction, purification and characterization of FgaPT2, a dimethylallyltryptophan synthase from Aspergillus fumigatus. Microbiology 151:1499–1505 Uthandi S, Saad B, Humbard MA, Maupin-Furlow JA (2010) LccA, an archaeal laccase secreted as a highly stable glycoprotein into the extracellular medium by Haloferax volcanii. Appl Environ Microbiol 76: 733–743 van den Berg MA, Albang R, Albermann K, Badger JH, Daran J-M, M Driessen AJ, Garcia-Estrada C, Fedorova ND, Harris DM, Heijne WHM, Joardar V, W Kiel JAK, Kovalchuk A, Martin JF, Nierman WC, Nijland JG, Pronk JT, Roubos JA, van der Klei IJ, van Peij NNME, Veenhuis M, von Dohren H, Wagner C, Wortman J, Bovenberg RAL (2008) Genome sequencing and analysis of the filamentous fungus Penicillium chrysogenum. Nat Biotech 26:1161–1168 van der Does HC, Rep M (2007) Virulence genes and the evolution of host specificity in plant-pathogenic fungi. Mol Plant Microbe Interact 20:1175–1182 Velasco AM, Leguina JI, Laczcano A (2002) Molecular evolution of the lysine biosynthetic pathways. J Mol Evol 55:445–459 Vershinin A (1999) Biological functions of carotenoids – diversity and evolution. Biofactors 10:99–104 Vidal-Cros A, Viviani F, Labesse G, Boccara M, Gaudry M (1994) Polyhydroxynaphthalene reductase involved in melanin biosynthesis in Magnaporthe grisea. Purification, cDNA cloning and sequencing. Eur J Biochem 219:985–992 Vining LC (1990) Functions of secondary metabolites. Annu Rev Microbiol 44:395–427 Walton JD (2000) Horizontal gene transfer and the evolution of secondary metabolite gene clusters in fungi: an hypothesis. Fungal Genet Biol 30:167–171 Wang YL, Aisen P, Casadevall A (1996) Melanin, melanin “ghosts,” and melanin composition in Cryptococcus neoformans. Infect Immun 64:2420–2424 Wapinski I, Pfeffer A, Friedman N, Regev A (2007) Natural history and evolutionary principles of gene duplication in fungi. Nature 449:54–61 Wheeler MH, Tolmsoff WJ, Bell AA, Mollenhauer HH (1978) Ultrastructural and chemical distinction of melanins formed by Verticillium dahliae from (þ)-scytalone, 1,8-dihydroxynaphthalene, catechol, and L-3,4-dihydroxyphenylalanine. Can J Microbiol 24:289–297

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

Evolution of Genes for Secondary Metabolism in Fungi Youngchim S, Hay RJ, Hamilton AJ (2005) Melanization of Penicillium marneffei in vitro and in vivo. Microbiology 151:291–299 Zhang A, Lu P, Dahl-Roshak AM, Paress PS, Kennedy S, Tkacz JS, An Z (2003) Efficient disruption of a polyketide synthase gene ( pks1) required for melanin synthesis through Agrobacterium-mediated transformation of Glarea lozoyensis. Mol Genet Genomics 268:645–655 Zhang S, Monahan BJ, Tkacz JS, Scott B (2004) Indolediterpene gene cluster from Aspergillus flavus. Appl Env Microbiol 70:6875–6883

255

Zhdanova NN, Lashko TN, Redchits TI, Vasilevskaia AI, Borisiuk LG, Siniavskaia OI, Gavriliuk VI, Muzalev PN (1991) The interaction of soil micromycetes with “hot” particles in a model system. Mikrobiol Zh 53:9–17 Zhu X, Williamson PR (2004) Role of laccase in the biology and virulence of Cryptococcus neoformans. FEMS Yeast Res 5:1–10 Zimmerman WC, Blanchette RA, Burnes TA, Farrell RL (1995) Melanin and perithecial development in Ophiostoma piliferum. Mycologia 87:857–863

.

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

258

Skander Elleuche

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

259

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

260

Skander Elleuche

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

Carbonic Anhydrases in Fungi and Fungal-Like Organisms

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

261

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

262

Skander Elleuche

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

Carbonic Anhydrases in Fungi and Fungal-Like Organisms

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

263

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

264

Skander Elleuche

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)

Carbonic Anhydrases in Fungi and Fungal-Like Organisms

265

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

266

Skander Elleuche

(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

Carbonic Anhydrases in Fungi and Fungal-Like Organisms

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

267

(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),

268

Skander Elleuche

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

Carbonic Anhydrases in Fungi and Fungal-Like Organisms

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

269

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

270

Skander Elleuche

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

Carbonic Anhydrases in Fungi and Fungal-Like Organisms

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.

References Aguilera J, Van Dijken JP, De Winde JH, Pronk JT (2005) Carbonic anhydrase (Nce103p): an essential biosynthetic enzyme for growth of Saccharomyces cerevisiae at atmospheric carbon dioxide pressure. Biochem J 391:311–316 Aspatwar A, Tolvanen ME, Parkkila S (2010) Phylogeny and expression of carbonic anhydrase-related proteins. BMC Mol Biol 11:25 Bahn YS, Cox GM, Perfect JR, Heitman J (2005) Carbonic anhydrase and CO2 sensing during Cryptococcus neoformans growth, differentiation, and virulence. Curr Biol 15:2013–2020 Bahn YS, Mu¨hlschlegel FA (2006) CO2 sensing in fungi and beyond. Curr Opin Microbiol 9:572–578 Bahn YS, Xue C, Idnurm A, Rutherford JC, Heitman J, Cardenas ME (2007) Sensing the environment: lessons from fungi. Nat Rev Microbiol 5:57–69 Barnett MJ, Fisher RF, Jones T, Komp C, Abola AP, BarloyHubler F, Bowser L, Capela D, Galibert F, Gouzy J, Gurjal M, Hong A, Huizar L, Hyman RW, Kahn D, Kahn ML, Kalman S, Keating DH, Long SR (2001) Nucleotide sequence and predicted functions of the entire Sinorhizobium meliloti pSymA megaplasmid. Proc Natl Acad Sci U S A 98:9883–9888 Chen Y Cann MJ, Litvin TN, Iourgenko V, Sinclair ML, Levin LR, Buck J (2000) Soluble adenylyl cyclase as an evolutionarily conserved bicarbonate sensor. Science 289:625–628

271

Chirica LC, Elleby B, Jonsson BH, Lindskog S (1997) The complete sequence, expression in Escherichia coli, purification and some properties of carbonic anhydrase from Neisseria gonorrhoeae. Eur J Biochem 244: 755–760 Cleves AE, Cooper DN, Barondes SH, Kelly RB (1996) A new pathway for protein export in Saccharomyces cerevisiae. J Cell Biol 133:1017–1026 Covarrubias AS, Bergfors T, Jones TA, Hogbom M (2006) Structural mechanics of the pH-dependent activity of beta-carbonic anhydrase from Mycobacterium tuberculosis. J Biol Chem 281:4993–4999 Cronk JD, Endrizzi JA, Cronk MR, O’Neill J W, Zhang KY (2001) Crystal structure of E. coli beta-carbonic anhydrase, an enzyme with an unusual pH-dependent activity. Protein Sci 10:911–922 Cronk JD, Rowlett RS, Zhang KY, Tu C, Endrizzi JA, Lee J, Gareiss PC, Preiss JR (2006) Identification of a novel noncatalytic bicarbonate binding site in eubacterial beta-carbonic anhydrase. Biochemistry 45:4351–4361 Demuth JP, Hahn MW (2009) The life and death of gene families. Bioessays 31:29–39 Eichinger L, Pachebat JA, Glo¨ckner G, Rajandream MA, Sucgang R, Berriman M, Song J, Olsen R, Szafranski K, Xu Q, Tunggal B, Kummerfeld S, Madera M, Konfortov BA, Rivero F, Bankier AT, Lehmann R, Hamlin N, Davies R, Gaudet P, Fey P, Pilcher K, Chen G, Saunders D, Sodergren E, Davis P, Kerhornou A, Nie X, Hall N, Anjard C, Hemphill L, Bason N, Farbrother P, Desany B, Just E, Morio T, Rost R, Churcher C, Cooper J, Haydock S, van Driessche N, Cronin A, Goodhead I, Muzny D, Mourier T, Pain A, Lu M, Harper D, Lindsay R, Hauser H, James K, Quiles M, Madan Babu M, Saito T, Buchrieser C, Wardroper A, Felder M, Thangavelu M, Johnson D, Knights A, Loulseged H, Mungall K, Oliver K, Price C, Quail MA, Urushihara H, Hernandez J, Rabbinowitsch E, Steffen D, Sanders M, Ma J, Kohara Y, Sharp S, Simmonds M, Spiegler S, Tivey A, Sugano S, White B, Walker D, Woodward J, Winckler T, Tanaka Y, Shaulsky G, Schleicher M, Weinstock G, Rosenthal A, Cox EC, Chisholm RL, Gibbs R, Loomis WF, Platzer M, Kay RR, Williams J, Dear PH, Noegel AA, Barrell B, Kuspa A (2005) The genome of the social amoeba Dictyostelium discoideum. Nature 435:43–57 Elleuche S, Po¨ggeler S (2008) A cyanase is transcriptionally regulated by arginine and involved in cyanate decomposition in Sordaria macrospora. Fungal Genet Biol 45:1458–1469 Elleuche S, Po¨ggeler S (2009a) Beta-carbonic anhydrases play a role in fruiting body development and ascospore germination in the filamentous fungus Sordaria macrospora. PLoS One 4:e5177 Elleuche S, Po¨ggeler S (2009b) Evolution of carbonic anhydrases in fungi. Curr Genet 55:211–222 Elleuche S, Po¨ggeler S (2010) Carbonic anhydrases in fungi. Microbiology 156:23–29 Emanuelsson O, Brunak S, von Heijne G, Nielsen H (2007) Locating proteins in the cell using TargetP, SignalP and related tools. Nat Protoc 2:953–971 Fabre N, Reiter IM, Becuwe-Linka N, Genty B, Rumeau D (2007) Characterization and expression analysis of

272

Skander Elleuche

genes encoding alpha and beta carbonic anhydrases in Arabidopsis. Plant Cell Environ 30:617–629 Felce J, Saier MH, Jr. (2004) Carbonic anhydrases fused to anion transporters of the SulP family: evidence for a novel type of bicarbonate transporter. J Mol Microbiol Biotechnol 8:169–176 Gogarten JP, Olendzenski L (1999) Orthologs, paralogs and genome comparisons. Curr Opin Genet Dev 9: 630–636 Go¨tz R, Gnann A, Zimmermann FK (1999) Deletion of the carbonic anhydrase-like gene NCE103 of the yeast Saccharomyces cerevisiae causes an oxygen-sensitive growth defect. Yeast 15:855–864 Gregor T, Fujimoto K, Masaki N, Sawai S (2010) The onset of collective behavior in social amoebae. Science 328:1021–1025 Gu Z, Steinmetz LM, Gu X, Scharfe C, Davis RW, Li WH (2003) Role of duplicate genes in genetic robustness against null mutations. Nature 421:63–66 Haas BJ, Kamoun S, Zody MC, Jiang RH, Handsaker RE, Cano LM, Grabherr M, Kodira CD, Raffaele S, TortoAlalibo T, Bozkurt TO, Ah-Fong AM, Alvarado L, Anderson VL, Armstrong MR, Avrova A, Baxter L, Beynon J, Boevink PC, Bollmann SR, Bos JI, Bulone V, Cai G, Cakir C, Carrington JC, Chawner M, Conti L, Costanzo S, Ewan R, Fahlgren N, Fischbach MA, Fugelstad J, Gilroy EM, Gnerre S, Green PJ, Grenville-Briggs LJ, Griffith J, Gru¨nwald NJ, Horn K, Horner NR, Hu CH, Huitema E, Jeong DH, Jones AM, Jones JD, Jones RW, Karlsson EK, Kunjeti SG, Lamour K, Liu Z, Ma L, Maclean D, Chibucos MC, McDonald H, McWalters J, Meijer HJ, Morgan W, Morris PF, Munro CA, O’Neill K, Ospina-Giraldo M, Pinzo´n A, Pritchard L, Ramsahoye B, Ren Q, Restrepo S, Roy S, Sadanandom A, Savidor A, Schornack S, Schwartz DC, Schumann UD, Schwessinger B, Seyer L, Sharpe T, Silvar C, Song J, Studholme DJ, Sykes S, Thines M, van de Vondervoort PJ, Phuntumart V, Wawra S, Weide R, Win J, Young C, Zhou S, Fry W, Meyers BC, van West P, Ristaino J, Govers F, Birch PR, Whisson SC, Judelson HS, Nusbaum C (2009) Genome sequence and analysis of the Irish potato famine pathogen Phytophthora infestans. Nature 461:393–398 Han KH, Chun YH, Figueiredo Bde C, Soriani FM, Savoldi M, Almeida A, Rodrigues F, Cairns CT, Bignell E, Tobal JM, Goldman MH, Kim JH, Bahn YS, Goldman GH, Ferreira ME (2010) The conserved and divergent roles of carbonic anhydrases in the filamentous fungi Aspergillus fumigatus and Aspergillus nidulans. Mol Microbiol 75:1372–1388 Hetherington AM, Raven JA (2005) The biology of carbon dioxide. Curr Biol 15:R406–R410 Hewett-Emmett D, Tashian RE (1996) Functional diversity, conservation, and convergence in the evolution of the alpha-, beta-, and gamma-carbonic anhydrase gene families. Mol Phylogenet Evol 5:50–77 Hibbett DS, Binder M, Bischoff JF, Blackwell M, Cannon PF, Eriksson OE, Huhndorf S, James T, Kirk PM, Lu¨cking R, Thorsten Lumbsch H, Lutzoni F, Matheny PB, McLaughlin DJ, Powell MJ, Redhead S, Schoch CL, Spatafora JW, Stalpers JA, Vilgalys R, Aime MC, Aptroot A, Bauer R, Begerow D, Benny GL, Castlebury

LA, Crous PW, Dai YC, Gams W, Geiser DM, Griffith GW, Gueidan C, Hawksworth DL, Hestmark G, Hosaka K, Humber RA, Hyde KD, Ironside JE, Ko˜ljalg U, Kurtzman CP, Larsson KH, Lichtwardt R, Longcore J, Miadlikowska J, Miller A, Moncalvo JM, MozleyStandridge S, Oberwinkler F, Parmasto E, Reeb V, Rogers JD, Roux C, Ryvarden L, Sampaio JP, Schu¨ssler A, Sugiyama J, Thorn RG, Tibell L, Untereiner WA, Walker C, Wang Z, Weir A, Weiss M, White MM, Winka K, Yao YJ, Zhang N (2007) A higher-level phylogenetic classification of the Fungi. Mycol Res 111:509–547 Ishida S, Muto S, Miyachi S (1993) Structural analysis of periplasmic carbonic anhydrase 1 of Chlamydomonas reinhardtii. Eur J Biochem 214:9–16 James TY, Kauff F, Schoch CL, Matheny PB, Hofstetter V, Cox CJ, Celio G, Gueidan C, Fraker E, Miadlikowska J, Lumbsch HT, Rauhut A, Reeb V, Arnold AE, Amtoft A, Stajich JE, Hosaka K, Sung GH, Johnson D, O’Rourke B, Crockett M, Binder M, Curtis JM, Slot JC, Wang Z, Wilson AW, Schu¨ssler A, Longcore JE, O’Donnell K, Mozley-Standridge S, Porter D, Letcher PM, Powell MJ, Taylor JW, White MM, Griffith GW, Davies DR, Humber RA, Morton JB, Sugiyama J, Rossman AY, Rogers JD, Pfister DH, Hewitt D, Hansen K, Hambleton S, Shoemaker RA, Kohlmeyer J, Volkmann-Kohlmeyer B, Spotts RA, Serdani M, Crous PW, Hughes KW, Matsuura K, Langer E, Langer G, Untereiner WA, Lu¨cking R, Bu¨del B, Geiser DM, Aptroot A, Diederich P, Schmitt I, Schultz M, Yahr R, Hibbett DS, Lutzoni F, McLaughlin DJ, Spatafora JW, Vilgalys R (2006) Reconstructing the early evolution of Fungi using a six-gene phylogeny. Nature 443:818–822 Jones NL (2008) An obsession with CO2. Appl Physiol Nutr Metab 33:641–650 Khaldi N, Wolfe KH (2008) Elusive origins of the extra genes in Aspergillus oryzae. PLoS One 3:e3036 Kim MS, Ko YJ, Maeng S, Floyd A, Heitman J, Bahn YS (2010) Comparative transcriptome analysis of the CO2 sensing pathway via differential expression of carbonic anhydrase in Cryptococcus neoformans. Genetics 185: 1207–1219 Kimber MS, Pai EF (2000) The active site architecture of Pisum sativum beta-carbonic anhydrase is a mirror image of that of alpha-carbonic anhydrases. Embo J 19:1407–1418 Kivela¨ AJ, Parkkila S, Saarnio J, Karttunen TJ, Kivela J, Parkkila AK, Bartosova M, Mucha V, Novak M, Waheed A, Sly WS, Rajaniemi H, Pastorekova S, Pastorek J (2005) Expression of von Hippel–Lindau tumor suppressor and tumor-associated carbonic anhydrases IX and XII in normal and neoplastic colorectal mucosa. World J Gastroenterol 11:2616–2625 Klengel T, Liang WJ, Chaloupka J, Ruoff C, Schro¨ppel K, Naglik JR, Eckert SE, Mogensen EG, Haynes K, Tuite MF, Levin LR, Buck J, Mu¨hlschlegel FA (2005) Fungal adenylyl cyclase integrates CO2 sensing with cAMP signaling and virulence. Curr Biol 15: 2021–2026 Ku¨ck U, Po¨ggeler S, Nowrousian M, Nolting N, Engh I (2009) Sordaria macrospora, a model system for

Carbonic Anhydrases in Fungi and Fungal-Like Organisms fungal development. In: Anke T, Weber D (eds) Physiology and Genetics, 1 st edn. Springer, Berlin, Heidelberg, pp 17–39 Lane TW, Saito MA, George GN, Pickering IJ, Prince RC, Morel FM (2005) Biochemistry: a cadmium enzyme from a marine diatom. Nature 435:42 Loftus BJ, Fung E, Roncaglia P, Rowley D, Amedeo P, Bruno D, Vamathevan J, Miranda M, Anderson IJ, Fraser JA, Allen JE, Bosdet IE, Brent MR, Chiu R, Doering TL, Donlin MJ, D’Souza CA, Fox DS, Grinberg V, Fu J, Fukushima M, Haas BJ, Huang JC, Janbon G, Jones SJ, Koo HL, Krzywinski MI, KwonChung JK, Lengeler KB, Maiti R, Marra MA, Marra RE, Mathewson CA, Mitchell TG, Pertea M, Riggs FR, Salzberg SL, Schein JE, Shvartsbeyn A, Shin H, Shumway M, Specht CA, Suh BB, Tenney A, Utterback TR, Wickes BL, Wortman JR, Wye NH, Kronstad JW, Lodge JK, Heitman J, Davis RW, Fraser CM, Hyman RW (2005) The genome of the basidiomycetous yeast and human pathogen Cryptococcus neoformans. Science 307:1321–1324 Longcore JE, Pessier AP, Nichols DK (1999) Batrachochytrium dendrobaditis gen et sp nov, a chytrid pathogenic to amphibians. Mycologia 91:219–227 Mitra M, Lato SM, Ynalvez RA, Xiao Y, Moroney JV (2004) Identification of a new chloroplast carbonic anhydrase in Chlamydomonas reinhardtii. Plant Physiol 135:173–182 Mitsuhashi S, Mizushima T, Yamashita E, Yamamoto M, Kumasaka T, Moriyama H, Ueki T, Miyachi S, Tsukihara T (2000) X-ray structure of beta-carbonic anhydrase from the red alga, Porphyridium purpureum, reveals a novel catalytic site for CO2 hydration. J Biol Chem 275:5521–5526 Mogensen EG, Janbon G, Chaloupka J, Steegborn C, Fu MS, Moyrand F, Klengel T, Pearson DS, Geeves MA, Buck J, Levin LR, Mu¨hlschlegel FA (2006) Cryptococcus neoformans senses CO2 through the carbonic anhydrase Can2 and the adenylyl cyclase Cac1. Eukaryot Cell 5:103–111 Mogensen EG, Mu¨hlschlegel FA (2008) CO2 sensing and virulence in Candida albicans. In: Brackhage AA, Zipfel PF (eds) Human and Animal Relationships, 2nd edn. Springer, Berlin, Heidelberg, pp 83–94 Moroney JV, Bartlett SG, Samuelsson G (2001) Carbonic anhydrases in plant and algae. Plant, Cell & Environment 24:141–153 Moroney JV, Ynalvez RA (2007) Proposed carbon dioxide concentrating mechanism in Chlamydomonas reinhardtii. Eukaryot Cell 6:1251–1259 Nishida H, Beppu T, Ueda K (2009) Symbiobacterium lost carbonic anhydrase in the course of evolution. J Mol Evol 68:90–96 Nowrousian M, Stajich JE, Chu M, Engh I, Espagne E, Halliday K, Kamerewerd J, Kempken F, Knab B, Kuo HC, Osiewacz HD, Po¨ggeler S, Read ND, Seiler S, Smith KM, Zickler D, Ku¨ck U, Freitag M (2010) De novo assembly of a 40 Mb eukaryotic genome from short sequence reads: Sordaria macrospora, a model organism for fungal morphogenesis. PLoS Genet 6: e1000891

273

Otto SP, Yong P (2002) The evolution of gene duplicates. Adv Genet 46:451–483 Parisi G, Perales M, Fornasari MS, Colaneri A, GonzlezSchain N, Go´mez-Casati D, Zimmermann S, Brennicke A, Araya A, Ferry JG, Echave J, Zabaleta E (2004) Gamma carbonic anhydrases in plant mitochondria. Plant Mol Biol 55:193–207 Park H, Song B, Morel FM (2007) Diversity of the cadmium-containing carbonic anhydrase in marine diatoms and natural waters. Environ Microbiol 9: 403–413 Rosenblum EB, Voyles J, Poorten TJ, Stajich JE (2010) The deadly chytrid fungus: a story of an emerging pathogen. PLoS Pathog 6:e1000550 Saran S, Meima ME, Alvarez-Curto E, Weening KE, Rozen DE, Schaap P (2002) cAMP signaling in Dictyostelium. Complexity of cAMP synthesis, degradation and detection. J Muscle Res Cell Motil 23:793–802 Sawaya MR, Cannon GC, Heinhorst S, Tanaka S, Williams EB, Yeates TO, Kerfeld CA (2006) The structure of beta-carbonic anhydrase from the carboxysomal shell reveals a distinct subclass with one active site for the price of two. J Biol Chem 281:7546–7655 Schlicker C, Hall RA, Vullo D, Middelhaufe S, Gertz M, Supuran CT, Mu¨hlschlegel FA, Steegborn C (2009) Structure and inhibition of the CO2-sensing carbonic anhydrase Can2 from the pathogenic fungus Cryptococcus neoformans. J Mol Biol 385:1207–1220 Shah GN, Hewett-Emmett D, Grubb JH, Migas MC, Fleming RE, Waheed A, Sly WS (2000) Mitochondrial carbonic anhydrase CA VB: differences in tissue distribution and pattern of evolution from those of CA VA suggest distinct physiological roles. Proc Natl Acad Sci U S A 97:1677–1682 Skamnioti P, Furlong RF, Gurr SJ (2008) The fate of gene duplicates in the genomes of fungal pathogens. Commun Integr Biol 1:196–198 Sly WS, Hu PY (1995) Human carbonic anhydrases and carbonic anhydrase deficiencies. Annu Rev Biochem 64:375–401 Smith KS, Ferry JG (2000) Prokaryotic carbonic anhydrases. FEMS Microbiol Rev 24:335–366 Smith KS, Ingram-Smith C, Ferry JG (2002) Roles of the conserved aspartate and arginine in the catalytic mechanism of an archaeal beta-class carbonic anhydrase. J Bacteriol 184:4240–4245 Smith KS, Jakubzick C, Whittam TS, Ferry JG (1999) Carbonic anhydrase is an ancient enzyme widespread in prokaryotes. Proc Natl Acad Sci U S A 96:15184– 15189 Strop P, Smith KS, Iverson TM, Ferry JG, Rees DC (2001) Crystal structure of the “cab”-type beta class carbonic anhydrase from the archaeon Methanobacterium thermoautotrophicum. J Biol Chem 276:10299–10305 Supuran CT (2008a) Carbonic anhydrases – an overview. Curr Pharm Des 14:603–614 Supuran CT (2008b) Carbonic anhydrases: novel therapeutic applications for inhibitors and activators. Nat Rev Drug Discov 7:168–181 Teng YB, Jiang YL, He YX, He WW, Lian FM, Chen Y, Zhou CZ (2009) Structural insights into the substrate

274

Skander Elleuche

tunnel of Saccharomyces cerevisiae carbonic anhydrase Nce103. BMC Struct Biol 9:67 Thompson JD, Gibson TJ, Plewniak F, Jeanmougin F, Higgins DG (1997) The CLUSTAL_X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res 25: 4876–4882 Tripp BC, Smith K, Ferry JG (2001) Carbonic anhydrase: new insights for an ancient enzyme. J Biol Chem 276: 48615–48618 Weber RWS (2009) Recent Developments in the Molecular Taxonomy in Fungi. In: Anke T, Weber D (eds) Physiology and Genetics, 1 st edn. Springer, Berlin, Heidelberg, pp 1–15 Wistrand PJ (1981) The importance of carbonic anhydrase B and C for the unloading of CO2 by the human erythrocyte. Acta Physiol Scand 113:417–426

Xu Y, Feng L, Jeffrey PD, Shi Y, Morel FM (2008) Structure and metal exchange in the cadmium carbonic anhydrase of marine diatoms. Nature 452:56–61 Ynalvez RA, Xiao Y, Ward AS, Cunnusamy K, Moroney JV (2008) Identification and characterization of two closely related beta-carbonic anhydrases from Chlamydomonas reinhardtii. Physiol Plant 133:15–26 Zimin AV, Delcher AL, Florea L, Kelley DR, Schatz MC, Puiu D, Hanrahan F, Pertea G, Van Tassell CP, Sonstegard TS, Marc¸ais G, Roberts M, Subramanian P, Yorke JA, Salzberg SL (2009) A whole-genome assembly of the domestic cow, Bos taurus. Genome Biol 10:R42 Zimmerman S, Innocenti A, Casini A, Ferry JG, Scozzafava A, Supuran CT (2004) Carbonic anhydrase inhibitors. Inhibition of the prokaryotic beta and gamma-class enzymes from Archaea with sulfonamides. Bioorg Med Chem Lett 14:6001–6006

.

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. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

277 278 278 279 279 281 281 282 283 284 285 285 286 287 287 288 288

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

278

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

279

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

280

Carrie A. Whittle and Hanna Johannesson

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

Evolution of Mating-Type Loci and Mating-Type Chromosomes

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

281

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

Carrie A. Whittle and Hanna Johannesson

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

Evolution of Mating-Type Loci and Mating-Type Chromosomes

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

283

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),

284

Carrie A. Whittle and Hanna Johannesson

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

Evolution of Mating-Type Loci and Mating-Type Chromosomes

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

285

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

286

Carrie A. Whittle and Hanna Johannesson

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

Evolution of Mating-Type Loci and Mating-Type Chromosomes

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

287

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

288

Carrie A. Whittle and Hanna Johannesson

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.

References Alexopoulos CJ, Mims CW, Blackwell M (1996) Introductory mycology, 4th edn. Wiley, New York Bachtrog D (2003a) Protein evolution and codon usage bias on the neo-sex chromosomes of Drosophila miranda. Genetics 165: 1221–1232 Bachtrog D (2003b) Adaptation shapes patterns of genome evolution on sexual and asexual chromosomes in Drosophila. Nat Genet 34: 215–219 Bachtrog D (2003c) Accumulation of spock and worf, two novel non-LTR retrotransposons on the neo-Y

Evolution of Mating-Type Loci and Mating-Type Chromosomes chromosome of Drosophila miranda. Mol Biol Evol 20: 173–181 Bachtrog D (2005) Sex chromosome evolution: Molecular aspects of Y-chromosome degeneration in Drosophila. Genome Res 15: 1393–1401 Bachtrog D (2006) Expression profile of a degenerating neo-Y chromosome in Drosophila. Curr Biol 16: 1694–1699 Bachtrog D, Charlesworth B (2002) Reduced adaptation of a non-recombining neo-Y chromosome. Nature 416: 323–326 Banham AHR, Asante-Owusu RN, Gottgens B, Thompson S, Kingsworth CS (1995) An N-terminal dimerization domain permits homeodomain proteins to choose compatible partners and initiate sexual development in the mushroom Coprinus cinereus. Plant Cell 7:773–783 Bartolome´ C, Charlesworth B (2006) Evolution of amino acid sequences and codon usage on the Drosophila miranda neo-sex chromosomes. Genetics 174: 2033– 2044 Barve MP, Arie T, Salimath SS, Muehlbauer FJ, Peever TL (2003) Cloning and characterization of the mating type (MAT) locus from Ascochyta rabiei (teleomorph: Didymella rabiei) and a MAT phylogeny of legume-associated Ascochyta spp. Fungal Genet Biol 39:151–167 Bobrowicz P, Pawlak R, Correa A, Bell-Pedersen D, Ebbole D J (2002) The Neurospora crassa pheromone precursor genes are regulated by the mating type locus and circadian clock. Mol Microbiol 45:795–804 Beatty NP, Smith ML, Glass NL (1994) Molecular characterization of mating-type loci in selected homothallic species of Neurospora, Gelasinospora and Anixiella. Mycol Res 98:1309–1316 Brasier CM (1988) Rapid changes in genetic structure of epidemic populations of Ophiostoma ulmi. Nature 332:538–541 Brooks LD (1988) The evolution of recombination rates. In: Michod RE, Levin BR (eds) The evolution of sex. Sinauer, Sunderland, pp 87–105 Bubnick M, Smulian AG (2007) The MAT1 locus of Histoplasma capsulatum is responsive in a mating type-specific manner. Eukaryot Cell 6:616–621 Carvalho AB (2002) Origin and evolution of the Drosophila Y chromosome. Curr Opin Genet Dev 12:664–668 Casselton LA (2002) Mate recognition in fungi. Heredity 99:142–147 Casselton A (2008) Fungal sex genes-searching for the ancestors. Bioessays 30:711–714 Casselton LA, Olesnicky NS (1998) Molecular genetics of mating recognition in basidiomycete fungi. Microbiol Mol Biol Rev 62:55–70 Charlesworth B (1996) The evolution of chromosomal sex determination and dosage compensation. Curr Biol 6:149–162 Charlesworth B, Charlesworth D (2000) The degeneration of Y chromosomes. Philos Trans R Soc Lond Ser B 355: 1563–1572 Charlesworth D, Charlesworth B, Marais G (2005) Steps in the evolution of heteromorphic sex chromosomes. Heredity 95:118–128

289

Charlesworth B, Coyne JA, Barton NH (1987) The relative rates of evolution of sex chromosomes and autosomes. Am Nat 130:113 Coppin E, Debuchy R, Arnaise S, Picard (1997) Mating types and sexual development in filamentous ascomycetes. Microbiol Mol Biol Rev 61:411–428 Debuchy R, Coppin E (1992) The mating types of Podospora anserina: functional analysis and sequence of the fertilization domains. Mol Gen Genet 233: 113–121 Debuchy R, Turgeon BG (2006) Mating-type structure, evolution, and function in Euascomycetes. In: Ku˝es U, Fischer R (eds) The Mycota I. Growth, differentiation and sexuality, Springer, Berlin Heidelberg New York, pp 293–323 Debuchy R, Arnaise S, Lecellier G (1993) The mat-allele of Podospora anserina contains three regulatory genes required for the development of fertilized female organs. Mol Gen Genet 241:667–673 Dettman JR, Taylor JW (2004) Mutation and evolution of microsatellite loci in Neurospora. Genetics 168: 1231–1248 Dettman J R., Jacobson DJ, Taylor J.W (2003) A multilocus genealogical approach to phylogenetic species recognition in the model eukaryote Neurospora. Evolution 57:2703–2720 Dettman JR, Jacobson DJ, Taylor JW ( 2006) Multilocus sequence data reveal extensive phylogenetic species diversity within the Neurospora discreta complex. Mycologia 98:436–446 Dodge BO (1927) Nuclear phenomena associated with heterothallism and homothallism in the ascomycete Neurospora. J Agric Res 35:289–305 Duret L (2000) tRNA gene number and codon usage in the C. elegans genome are coadapted for optimal translation of highly expressed genes. Trends Genet 16: 287–289 Duret L, Mouchiroud D (1999) Expression pattern and, surprisingly, gene length shape codon usage in Caenorhabditis, Drosophila, and Arabidopsis. Proc Natl Acad Sci USA 96:4482–4487 Dyer PS (2008) Evolutionary biology: genomic clues to original sex in fungi. Curr Biol 18:R207–R209 Ellegren H (2000) Evolution of the avian sex chromosome and their role in sex determination. Trends Ecol Evol 15:188–192 Ellegren H, Parsch J (2007) The evolution of sex-biased genes and sex-biased gene expression. Nat Rev Genet 8:689–698 Ferreira AVB, Saupe S, Glass NL (1996) Transcriptional analysis of the mtA idiomorph of Neurospora crassa identifies two genes in addition to mt A-1. Mol Gen Genet 250:767–774 Ferreira AVB, An ZQ, Metzenberg RL, Glass NL (1998) Characterization of mat A-2, mat A-3 and Delta matA mating-type mutants of Neurospora crassa. Genetics 148:1069–1079 Fraser JA, Heitman J (2004) Evolution of fungal sex chromosomes. Mol Microbiol 51:299–306 Fraser JA, Heitman J (2005) Chromosomal sex-determining regions in animals, plants and fungi. Curr Opin Genet Dev 15:645–651

290

Carrie A. Whittle and Hanna Johannesson

Felsenstein J (1974) The evolutionary advantage of recombination. Genetics 78:737–756 Fraser JA, Diezmann S, Subaran RL, Allen A, Lengeler KB, Dietrich FS, Heitman J (2004) Convergent evolution of chromosomal sex-determining regions in the animal and fungal kingdoms. PLoS Biol 2:2243–2255 Fraser JA, Stajich JE, Tarcha EJ, Cole GT, Inglis DO, Sil A, Heitman J (2007) Evolution of the mating type locus: insights gained from the dimorphic primary fungal pathogens Histoplasma capsulatum, Coccidioides immitis, and Coccidioides posadasii. Eukaryot Cell 6:622–629 Gallegos A, Jacobson DJ, Raju NB, Skupski MP, Natvig DO (2000) Suppressed recombination and a pairing anomaly on the mating-type chromosome of Neurospora tetrasperma. Genetics 154:623–633 Galagan JE, Calvo SE, Cuomo C, Ma LJ, Wortman JR, Batzoglou S, Lee SI, Bas¸tu¨rkmen M, Spevak CC, Clutterbuck J (2005) Sequencing of Aspergillus nidulans and comparative analysis with A. fumigatus and A. oryzae. Nature 438:1105–1115 Glass NL, Smith M L (1994) Structure and function of a mating-type gene from the homothallic species Neurospora africana. Mol Gen Genet 244:401–409 Glass NL, Vollmer SJ, Staben C, Grotelueschen J, Metzenberg RL, Yanofsky C (1990a) DNAs of the two mating-type alleles of Neurospora crassa are highly dissimilar. Science 241:570–573 Glass NL, Grotelueschen J, Metzenberg RL (1990b) Neurospora crassa A mating-type region. Proc Natl Acad Sci USA 87:4912–4816 Haber JE (1998) Mating-type switching in Saccharomyces cerevisiae. Annu Rev Genet 32:561–599 Handley LL, Ceplitis H, Ellegren H (2004) Evolutionary strata on the chicken Z chromosome: implications for sex chromosome evolution. Genetics 167:367–376 Hedrick PW (2007) Sex: Differences in mutation, recombination, selection, gene flow and genetic drift. Evolution 61:2750–2771 Heitman J, Kronstad JW, Taylor JW, Casselton LA (2007). Sex in fungi: molecular determination and evolutionary implications. ASM Press, Washington, DC Hood ME (2002) Dimorphic mating-type chromosomes in the fungus Microbotryum violaceum. Genetics 160: 457–461 Hood ME, Antonovics J, Koskella B (2004) Shared forces of sex chromosomes evolution in haploid-mating and diploid mating organisms: Microbotryum violaceum and other model organisms. Genetics 168:141–146 Hsueh Y-P, Idnurm A, Heitman J (2006) Recombination hotspots flank the Cryptococcus mating-type locus: implications for the evolution of a fungal sex chromosome. PLoS Genet 2:e184 Hull CM, Johnson AD (1999) Identification of a mating type-like locus in the asexual pathogen Candida albicans. Science 285:1271–1275 Idnurm A, Walton FJ, Floyd A, Heitman J (2008) Identification of the sex genes in an early diverged fungus. Nature 451:193–196 Jacobson DJ (1995) Sexual dysfunction associated with outcrossing in Neurospora tetrasperma, a pseudohomothallic ascomycete. Mycologia 87:604–617

Jacobson DJ (2005) Blocked recombination along the mating-type chromosomes of Neurospora tetrasperma involves both structural heterozygosity and autosomal genes. Genetics 171:839–843 James TY, Kauff F, Schoch CL, Matheny PB, Hofstetter V, Cox CJ, Celio G, Gueidan C, Fraker E, Miadlikowska J et al. (2006) Reconstructing the early evolution of Fungi using a six-gene phylogeny. Nature 443: 758–761 Kang S, Chumley FG, Valent B (1994) Isolation of the mating-type genes of the phytopathogenic fungus Magnaporthe grisea using genomic subtraction. Genetics 138:289–296 Karlsson M, Nygren K, Johannesson H (2008) The evolution of the pheromonal signal system and its potential role for reproductive isolation in heterothallic Neurospora. Mol Biol Evol 25:168–178 Khakhlova O, Bock R (2006) Elimination of deleterious mutations in plastid genomes by gene conversion. Plant J 46:85–94 Kim H, Borkovich KA (2004) A pheromone receptor gene, pre-1,is essential for mating type-specific directional growth and fusion of trichogynes and female fertility in Neurospora crassa. Mol Microbiol 52:1781–1798 Kim H, Borkovich KA (2006) Pheromones are essential for male fertility and sufficient to direct chemotropic polarized growth of trichogynes during mating in Neurospora crassa. Eukaryot Cell 5:544–554 Kothe E (1996) Tetrapolar fungal mating types: sexes by the thousands. FEMS Microbiol Rev 18:65–87 Kronstad JW, Staben C (1997) Mating type in filamentous fungi. Annu Rev Genet 31:245–276 Lahn BT, Page DC (1999) Four evolutionary strata on the human X chromosome. Science 286:964–967 Lahn BT, Pearson NM, Jegalian K (2001) The human Y chromosome, in the light of evolution. Nat Rev Genet 2:207–216 Lee N, Bakkeren G, Wong K, Sherwood JE, Kronstad JW (1999) The mating-type and pathogenicity locus of the fungus Ustilago hordei spans a 500-kb region. Proc Natl Acad Sci USA 96:15026–15031 Lengeler KB, Fox DS, Fraser JA, Allen A, Forrester K, Dietrich FS, Heitman J (2002) Mating-type locus of Cryptococcus neoformans: a step in the evolution of sex chromosomes. Eukaryot Cell 1:704–718 Li W-H, Yi S, Makova K (2002) Male-driven evolution. Curr Opin Genet Dev 12:650–656 Mandel MA, Barker BM, Kroken S, Rounsley SD, Orbach MJ (2007) Genomic and population analyses of the mating type loci in Coccidioides species reveal evidence for sexual reproduction and gene acquisition. Eukaryot Cell 6:1189–1199 Mank JE, Axelsson E, Ellegren H (2007) Fast-X on the Z: rapid evolution of sex-linked genes in birds. Genome Res 17:618–624 Marais GA, Nicolas M, Bergero R, Chambrier P, Kejnovsky E, Mone´ger F, Hobza R, Widmer A, Charlesworth D (2008) Evidence for degeneration of the Y chromosome in the dioecious plant Silene latifolia. Curr Biol 18:545–549

Evolution of Mating-Type Loci and Mating-Type Chromosomes Menkis A, Whittle CA, Johannesson H (2010) Gene genealogies indicates abundant gene conversions and independent evolutionary histories of the mating-type chromosomes in the evolutionary history of Neurospora tetrasperma. BMC Evol Biol 10:234 Menkis A, Bastiaans E, Jacobson DJ, Johannesson H (2009) Phylogenetic and biological species diversity within the Neurospora tetrasperma complex. J Evol Biol 22: 1923–1936 Menkis A, Jacobson DJ, Gustafsoon T, Johannesson H (2008) The mating-type chromosome in the filamentous asycomycete Neurospora tetrasperma represents a model for early evolution of sex chromosomes. PLoS Genet 4:e1000030 Merino ST, Nelson MA, Jacobson DJ, Natvig DO (1996) Pseudohomothallism and evolution of the matingtype chromosome in Neurospora tetrasperma. Genetics 143:789–799 Nei M (1970) Accumulation of nonfunctional genes on sheltered chromosomes. Am Nat 104:311–322 Nicolas M, Marais G, Hykelova V, Janousek B, Laporte V, Vyskot B, Mouchiroud D, Negrutiu I, Charlesworth D, Mone´ger (2004) A gradual process of recombination restriction in the evolutionary history of the sex chromosomes in dioecious plants. PLoS Biol 3:e4 Ohta T (1989) The mutational load of a multigene family with uniform members. Genet Res 53:141–145 O’Shea SF, Chaure PT, Halsall JR, Olesnicky NS, Leibbrandt A, Connerton IF, Casselton LA (1998) A large pheromone and receptor gene complex determines multiple B mating type specificities in Coprinus cinereus. Genetics 148:1081–1090 Pamilo P, Nei M (1988) Relationships between gene trees and species trees. Mol Biol Evol 5:568–583 Paoletti M, Rydholm C, Schwier EU, Anderson MJ, Szakacs G, Lutzoni F, Debeaupuis JP, Latge´ JP, Denning DW, Dyer PS (2005) Evidence for sexuality in the opportunistic fungal pathogen Aspergillus fumigatus. Curr Biol 15:1242–1248 Paoletti M, Buck KW, Brasier CM (2006) Selective acquisition of novel mating type and vegetative incompatibility genes via interspecies gene transfer in the globally invading eukaryote Ophiostoma novo-ulmi. Mol Ecol 15:249–262 Po¨ggeler S (1999) Phylogenetic relationships between mating-type sequences from homothallic and heterothallic ascomyctes. Curr Genet 36:222–231 Po¨ggeler S, Ku¨ck U (2000) Comparative analysis of the mating-type loci from Neurospora crassa and Sordaria macrospora: identification of novel transcribed ORFs. Mol Gen Genet 263:292–301 Po¨ggeler S, Risch S, Ku¨ck U, Osiewacz HD (1997) Matingtype genes from the homothallic fungus Sordaria macrospora are functionally expressed in a heterothallicascomycete. Genetics 147:567–580 Powell AJ, Jacobson DJ and Natvig DI (2001) Allelic diversity at the het-c locus in Neurospora tetrasperma confirms outcrossing in nature and reveals an evolutionary dilemma for pseudohomothallic ascomycetes. J Mol Evol 52:94–102

291

Raju NB, Perkins DD (1994) Diverse programs of ascus development in pseudohomothallic species of Neurospora, Gelasinospora, and Podospora. Dev Genet 15:104–118 Rappold GA (1993) The pseudoautosomal regions of the human sex chromosomes. Hum Genet 92:315–324 Repping S, Skaletsky H, Brown L, van Daalen SK, Korver CM, Pyntikova T, Kuroda-Kawaguchi T, de Vries JW, Oates RD, Silber S, van der Veen F, Page DC, Rozen S (2003) Polymorphism for a 1.6-Mb deletion of the human Y chromosome persists through balance between recurrent mutation and haploid selection. Nat Genet 35:247–251 Rice WR (1987) Genetic hitch-hiking and the evolution of reduced genetic activity of the Y sex chromosome. Genetics 116:161–167 Sandstedt SA, Tucker PK (2004) Evolutionary strata on the mouse X chromosome correspond to strata on the human X chromosome. Genome Res 14:267–272 Scott MP, Matsudaira P, Lodish H, Darnell J, Zipursky L, Kaiser CA, Berk A, Krieger M (2004) Molecular cell biology, 5th edn. Freeman, New York Sharon A, Yamaguchi K, Christiansen S, Horwitz BA, Yoder OC, Turgeon BG (1996) An asexual fungus has the potential for sexual development. Mol Gen Genet 251:60–68 Shiu PK, Glass NL (2000) Cell and nuclear recognition mechanisms mediated by mating type in filamentous ascomycetes. Curr Opin Microbiol 3:183–188 Singh G, Ashby AM (1998) Cloning of the mating type loci from Pyrenopeziza brassicae reveals the presence of a novel mating type gene within a discomycete MAT 1–2 locus encoding a putative metallothionein-like protein. Mol Microbiol 30:799–806 Skaletsky H., Kuroda-Kawaguchi T, Minx P, Cordum H, Hillier L, Brown L, Repping S, Pyntikova T, Ali, J, Bieri T, et al. (2003). The male-specific region of the human Y chromosome is a mosaic of discrete sequence classes. Nature 423:825–837 Spatafora JW, Johnson D, Sung G-H, Hosaka K, O’Rourke B, Serdani M, Spotts R, Lutzoni F, Hofstetter V, Fraker E (2006) A five-gene phylogenetic analysis of the Pezizomycotina. Mycologia 98:1020–1030 Staben C, Yanofsky C (1990) Neurospora crassa A mating-type region. Proc Natl Acad Sci USA 87: 4917–4921 Stoletzki N, Eyre-Walker A (2006) Synonymous codon usage in Escherichia coli: selection for translational accuracy. Mol Biol Evol 24:374–381 Strandberg R, Nygren K, Menkis A, James TY, Wik L, Stajich JE, Johannesson H (2010) Conflict between reproductive gene trees and species phylogeny among heterothallic and pseudohomothallic members of the filamentous ascomycete genus Neurospora. Fungal Genet Biol 47: 869–878 Sugiyama J, Hosaka K, Suh, S-O (2006) Early diverging Ascomycota: phylogenetic divergence and related evolutionary enigmas. Mycologia 98:996–1005 Swanson, W.J., Vacquier, VD (2002) The rapid evolution of reproductive proteins. Nat Rev Genet 3:137–143

292

Carrie A. Whittle and Hanna Johannesson

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

294

Christine Schimek

(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,

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

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.

295

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

296

Christine Schimek

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

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

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

297

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.

298

Christine Schimek

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

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

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).

299

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%,

300

Christine Schimek

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

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

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).

301

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

302

Christine Schimek

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,

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

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

303

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

304

Christine Schimek

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

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

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

305

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

306

Christine Schimek

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

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

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

307

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

308

Christine Schimek

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

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

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

309

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.

310

Christine Schimek

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

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

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).

311

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.

312

Christine Schimek

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

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

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

313

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

314

Christine Schimek

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

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

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

315

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.

316

Christine Schimek

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

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

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.

317

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

318

Christine Schimek

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

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

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.

319

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

320

Christine Schimek

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

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

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-

321

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

322

Christine Schimek

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.

References Abe Y, Ono C, Hosobuchi M, Yoshikawa H (2002) Functional analysis of mlcR, a regulatory gene for ML-236B (compactin) biosynthesis in Penicillium citrinum. Mol Genet Genom 268:352–361 Aharoni A, Gaidukov L, Khersonsky O, McQ Gould S; Roodveld C, Tawfik DS (2005) The ‘evolvability’ of promiscuous protein functions. Nat Genet 37:73–76 Aharonowitz Y, Cohen G, Martin JF (1992) Penicillin and cephalosporin biosynthetic genes: structure, organization, regulation and evolution. Annu Rev Microbiol 46:461–495 Ahn J-H, Walton JD (1996) Chromosomal organization of TOX2, the complex locus controlling host-selective toxin biosynthesis in Cochliobolus carbonum. Plant Cell 8:887–897 Akagi Y, Akamatsu H, Otani H, Kodama M (2009) Horizontal chromosome transfer, a mechanism for evolution and differentiation of a plant-pathogenic fungus. Eukaryot Cell 8:1732–1738 Akhmanova A, Voncken FGI, Harhangi H, Hosea KM, Vogels GD, Hackstein JHP (1998) Cytosolic enzymes with a mitochondrial ancestry from the anaerobic chytrid Piromyces sp. E2. Mol Microbiol 30:1017–1027 Andersson JO (2009) Horizontal gene transfer between microbial eukaryotes. Meth Mol Biol 532: 473–487 Bailey AM, Cox RJ, Harley K, Lazarus CM, Simpson TJ, Skellam E (2007) Characterization of 3-methylorcinaldehyde synthase (MOS) in Acremonium strictum: first observation of a reductive release mechanism during polyketide biosynthesis. Chem Commun 394053–4055 Batada NN, Urrutia AO, Hurst LD (2007) Chromatin remodelling is a major source of coexpression of linked genes in yeast. Trends Genet 23:480–484 Batada NN, Hurst LD (2007) Evolution of chromosome organization driven by selection for reduced gene expression noise. Nat Genet 39:945–949 Bayram O, Krappman S, Ni M, Bok JW, Helmstaedt, K, Valerius O, Braus-Stromeyer S, Kwon NJ, Keller NP, Yu JH, Braus GH (2008) VelB/VeA/LaeA complex coordinates light signal with fungal development and secondary metabolism. Science 320:1504–1506 Belbahri L, Calmin G, Mauch F, Andersson JO (2008) Evolution of the cutinase gene family: evidence for lateral transfer of a candidate Phytophthora virulence factor. Gene 408:1–8 Bobrowicz P, Wysocki R, Owsianik G, Goffeau A, Ulaszewski S (1997) Isolation of three contiguous genes, ACR1, ACR2 and ACR3, involved in resistance to arsenic compounds in the yeast Saccharomyces cerevisiae. Yeast 13:819–828 Bok JW, Keller NP (2004) LaeA, a regulator of secondary metabolism in Aspergillus spp. Eukaryot Cell 3:527–535 Bok JW, Noordermeer D, Kale S, Keller NP (2006) Metabolic gene cluster silencing in Aspergillus nidulans. Mol Microbiol 61:1636–1645 Bo¨hnert HU, Fudal I, Dioh W, Tharreau D, Notteghem J-L, Lebrun M-H (2004) A putative polyketide synthase/

Evolution of Special Metabolism in Fungi: Concepts, Mechanisms, and Pathways peptide synthetase from Magnaporthe grisea signals pathogen attack to resistant rice. Plant Cell 16. 2499–513 Bo¨mke C, Rojas MC, Gong F, Hedden P, Tudzynski B (2008a) Isolation and characterization of the gibberellin biosynthetic gene cluster in Sphaceloma manihoticola. Appl Environ Microbiol 74:5325–5339 Bo¨mke C, Rojas MC, Hedden P, Tudzynski B (2008b) Loss of gibberellin production is due to a deletion in the gibberellic acid gene cluster in Fusarium verticilloides (G. Fujikuroi MP-A). Appl Environ Microbiol 74:7790–7801 Bo¨mke C, Tudzynski B (2009) Diversity, regulation, and evolution of the gibberellin biosynthetic pathway in fungi compared to plants and bacteria. Phytochemistry 70:1876–1893 Borges-Walmsley ML, Turner G, Bailey AM Brown J, Lehmbeck J, Clausen IG (1995) Isolation and characterization of genes for sulphate activation and reduction in Aspergillus nidulans: implications for evolution of an allosteric control region by gene duplication. Bornscheuer UT, Kazlauskas RJ (2004) Catalytic promiscuity in biocatalysis: using old enzymes to from new bonds and follow new pathways. Angew. Chem Int Ed Engl 43:6032–6040 Bouhired S, Weber M, Kempf-Sontag A, Keller NP, Hoffmeister D (2007) Accurate prediction of the Aspergillus nidulans terrequinone gene cluster boundaries using the transcriptional regulator LaeA. Fungal Genet Biol 44:1134–1145 Brakhage AA, Al-Abdallah Q, Tu¨ncher A, Spro¨te P (2005) Evolution of b-lactam biosynthesis genes and recruitment of trans-acting factors. Phytochemistry 66:1200–1210 Brakhage AA, Spro¨te P, Al-Abdallah Q, Gehrke A, Plattner H, Tu¨ncher A (2004) Regulation of penicillin biosynthesis in filamentous fungi. Adv Biochem Eng Biotechnol 88:45–90 Brakhage AA, Tho¨n M, Spro¨te P, Scharf DH, Al-Abdallah Q, Wolke SM, Hortschansky P (2009) Aspects on evolution of fungal b-lactam biosynthesis clusters and recruitment of trans-acting factors. Phytochemistry 70:1801–1811 Bringmann G, Irmer A, Feineis D, Gulder TAM, Fiedler HP (2009) Convergence in the biosynthesis of acetogenic natural products from plants, fungi, and bacteria. Phytochemistry 70:1776–1788 Brown DW, Yu JH, Kelkar HS, Fernandes M, Nesbitt TC, Keller NP, Adams TH, Leonard TJ (1996) Twenty-five coregulated transcripts define a sterigmatocystin gene cluster in Aspergillus nidulans. Proc Natl Acad Sci USA 93:1418–1422 Burmester A, Richter M, Schultze K, Voelz K, Schachtschabel D, Boland W, Wo¨stemeyer J, Schimek C (2007) Cleavage of b-carotene as the first step in sexual hormone synthesis in zygomycetes is mediated by a trisporic acid regulated b-carotene oxygenase. Fungal Genet Biol 44:1096–1108 Caetano-Anolles G, Yafremava L, Gee H, Caetano-Anolles D, Kim HS, Mittenthal JE (2009) The origin and evolution of modern metabolism. Int J Biochem Cell Biol 41:285–297

323

Calvo AM (2008) The VeA regulatory system and its role in morphological and chemical development in fungi. Fungal Genet Biol 45:1053–1061 Calvo AM, Bok J, Brooks W, Keller NP (2004) VeA is required for toxin and sclerotial production in Aspergillus parasiticus. Appl Environ Microbiol 70:4733– 4739 Calvo AM, Gardner HW, Keller NP (2001) Genetic connection between fatty acid metabolism and sporulation in Aspergillus nidulans. J Biol Chem 276:25766–25774 Calvo AM; Hinze LL, Gardner HW, Keller NP (1999) Sporogenic effect of polyunsaturated fatty acids on developments of Aspergillus spp. Appl Environ Microbiol 65:3668–3673 Caruso ML, Litzka O, Martic G, Lottspeich F, Brakhage AA (2002) A novel basic-region helix-loop-helix transcription factor (AnBH1) of Aspergillus nidulans counteracts the CCAAT-binding complex AnCF in the promoter of a penicillin biosynthesis gene. J Mol Biol 323:425–439 Cary JW, Ehrlich KC (2006) Aflatoxigenicity in Aspergillus: molecular genetics, phylogenetic relationships and evolutionary implications. Mycopathologia 162:167– 177 Cavener DR (1992) GMC oxidoreductases – a newly defined family of homologous proteins with diverse catalytic activities. J Mol Biol 223:811–814 Challis GL, Hopwood DA (2003) Synergy and contingency as driving forces for the evolution of multiple secondary metabolite production by Streptomyces species. Proc Natl Acad Sci USA 100:14555–14561 Chang PK, Cary JW, Bhatnagar D, Cleveland TE, Bennett JW, Linz JE, Woloshuk CP, Payne GA (1993) Cloning of the Aspergillus parasiticus apa-2 gene associated with the regulation of aflatoxin biosynthesis. Appl Environ Microbiol 59:3273–3279 Chang PK, Matsushima K, Takahashi T, Yu J, Abe K, Bhatnagar D, Yuan GF, Koyama Y, Cleveland TE (2007) Understanding nonaflatoxigenicity of Aspergillus sojae: a windfall of aflatoxin biosynthesis research. Appl Microbiol Biotechnol 76:977–984 Chague V, Elad Y, Barakat R, Tudzynski P, Sharon A (2002) Ethylene biosynthesis in Botrytis cinera. FEMS Microbiol Ecol 40:143–149 Chapman DJ, Regan MA (1980) Evolution of a biochemical pathway: evidence from comparative biochemistry. Annu Rev Plant Physiol 31:639–645 Christianson DW (2006) Structural biology and chemistry of the terpenoid cyclases. Chem Rev 106:3412–3442 Clay K, Schardl CL (2002) Evolutionary origins and ecological consequences of endophyte symbioses with grasses. Am Nat 160:S99–S127 Cohen BA, Mitra RD, Hughes JD, Church GM (2000) A computational analysis of whole-genome expression data reveals chromosomal domains of gene expression. Nat Genet 26:183–186 Collemare J, Pianfetti M, Houlle AE, Morin D, Camborde L, Gagey MJ, Barbisan C, Fudal I, Lebrun MH, Bohnert HU (2008) Magnaporthe grisea avirulence gene ACE1 belongs to an infection-specific gene cluster involved in secondary metabolism. New Phytol 179:196–208

324

Christine Schimek

Cooper TG (1996) Regulation of allantoin catabolism in Saccharomyces cerevisiae. In: Marzluf GA (ed) The Mycota, vol 3. Biochemistry and Molecular Biology. Springer, Berlin, pp 139–169 Correia T, Grammel N, Ortel I, Keller U, Tudzynski P (2003) Molecular cloning and analysis of the ergopeptine assembly system in the ergot fungus Claviceps purpurea. Chem Biol 10:1281–1292 Cox RJ (2007) Polyketides, proteins and genes in fungi: programmed nano-machines begin to reveal their secrets. Org Biomol Chem 5:2010–2026 Czapek F (1922–1925) Biochemie der Pflanzen. 3rd ed., Fischer, Jena Czempinski K, Kruft V, Wo¨stemeyer J, Burmester A (1996) 4-Dihydromethyltrisporate dehydrogenase from Mucor mucedo, an enzyme of the sexual hormone pathway: purification, and cloning of the corresponding gene. Microbiology 142:2647–2654 Damrongkool P, Sedlock AB, Young CA, Johnson RD, Goetz KE, Scott B, Schardl CL, Panaccione DG (2005) Structural analysis of a peptide synthetase gene required for ergopeptine production in the endophytic fungus Neotyphodium lolii. DNA Seq 16:379–385 de Vos M, Kim JH, Jander G (2007) Biochemistry and molecular biology of Arabidopsis–aphid interactions. BioEssays 29:871–883 Dreyer J, Eichhorn H, Friedlin E, Kurnsteiner H, Ku¨ck U (2007) A homologue of the Aspergillus velvet gene regulates both cephalosporin C biosynthesis and hyphal fragmentation in Acremonium chrysogenum. Appl Environ Microbiol 73:3412–3422 Duncan K, Edwards RM, Coggins JR (1987) The pentafunctional arom enzyme of Saccharomyces cerevisiae is a mosaic of monofunctional domains. Biochem J 246:375–386 Ehrlich KC, Montalbano BG, Cary JW (1999) Binding of the C6-zinc cluster protein, AFLR, to the promoters of the aflatoxin pathway biosynthesis genes in Aspergillus parasiticus. Gene 230:249–257 Ehrlich KC, Yu J, Cotty PJ (2005) Aflatoxin biosynthesis gene clusters and flanking regions. J Appl Microbiol 99:518–527 Fairhead C, Dujon B (2006) Structure of Kluyveromyces lactis subtelomeres. FEMS Yeast Res 6:428–441 Ferguson BJ, Mathesius U (2003) Signaling interactions and nodule development. J Plant Growth Regul 22:47–72 Fernandes M, Keller NP, Adams TH (1998) Sequence-specific binding by Aspergillus nidulans AflR, a C6 zinc cluster protein regulating mycotoxin biosynthesis. Mol Microbiol 28:1355–1365 Firn RD, Jones CG (2000) The evolution of secondary metabolism – a unifying model. Mol Microbiol 37:989–994 Firn RD, Jones CG (2009) A Darwinian view of metabolism: molecular properties determine fitness. J Exp Bot 60:719–726 Fleetwood DJ, Scott B, Lane GA, Tanaka A, Johnson RD (2007) A complex ergovaline gene cluster in Epichloe¨ endophytes of grasses. Applied Environ Microbiol 73:2571–2579

Fothergill-Gilmore LA, Michaels PAM (1993) Evolution of glycolysis. Prog Biophys Mol Biol 59:105–235 Fox EM, Howlett BJ (2008) Biosynthetic gene clusters for epipolythiodioxopiperazines in filamentous fungi. Mycol Res 112:162–169 Fraser PD, Bramley PM (2004) The biosynthesis and nutritional uses of carotenoids. Prog Lipid Res 43:228–265 Friesen TL, Stukenbrock EH, Liu Z, Meinhardt S, Ling H, Faris JD, Rasmussen JD, Solomon PS, McDonald BA, Oliver RP (2006) Emergence of a new disease as a result of interspecific virulence gene transfer. Nat Gen 38:953–956 Galagan JE, Calvo SE, Cuomo C, Ma LJ, Wortman JR et al. (2005) Sequencing of Aspergillus nidulans and comparative analysis with A. fumigatus and A. oryzae. Nature 438:1105–1115 Garcia-Vallve S, Romeu A, Palau J (2006) Horizontal gene transfer of glycosyl hydrolases of the rumen fungi. Mol Biol Evol 17:352–361 Gardiner DM, Cozijnsen AJ, Wilson LM, Pedras MS, Howlett BJ (2004) The sirodesmin biosynthetic gene cluster of the plant pathogenic fungus Leptosphaeria maculans. Mol Microbiol 53:1307–1318 Gardiner DM, Howlett BJ (2005) Bioinformatic and expression analysis of the putative gliotoxin biosynthetic gene cluster of Aspergillus fumigatus. FEMS Microbiol Lett 248:241–248 Georgianna Dr, Payne GA (2009) Genetic regulation of aflatoxin biosynthesis: from gene to genome. Fungal Genet Biol 46:113–125 Gilbert HJ, Hazlewood GP, Laurie JI, Orpin CG, Xue GP (1992) Homologous catalytic domains in a rumen fungal xylanase – Evidence for gene duplication and prokaryotic origin. Mol Microbiol 6:2065–2072 Greenhagen BT, O’Maille PE, Noel JP, Chappell J (2006) Identifying and manipulating structural determinates linking catalytic specificities in terpene synthases. Proc Natl Acad Sci USA 103:9826–9831 Haarmann T, Machado C, Lu¨bbe Y, Correia T, Schardl CL, Panaccione DG, Tudzynski P (2005) The ergot alkaloid gene cluster in Claviceps purpurea: extension of the cluster sequence and intra species evolution. Phytochemistry 666:1312–1320 Haarmann T, Ortel I, Tudzynski P, Keller U (2006) Identification of the cytochrome P450 monooxygenase that bridges the clavine and ergoline alkaloid pathways. ChemBioChem 7:645–652 Haas H, Marzluf GA (1995) NRE, the major nitrogen regulatory protein of Penicillium chrysogenum binds specifically to elements in the intergenic promoter regions of nitrate assimilation and penicillin biosynthetic gene clusters. Curr Genet 28:177–183 Hall C, Dietrich FS (2007) The reacquisition of biotin prototrophy in Saccharomyces cerevisiae involves horizontal gene transfer, gene duplication and gene clustering. Genetics 177:2293–2307 Hallberg BM, Hendriksson G, Petterson G, Divne C (2002) Crystal structure of the flavoprotein domain of the extracellular flavocytochrome cellobiose dehydrogenase. Structure 8:79–88 Halme A, Bumgarner S, Styles C, Fink GR (2004) Genetic and epigenetic regulation of the FLO gene family

Evolution of Special Metabolism in Fungi: Concepts, Mechanisms, and Pathways generates cell-surface variation in yeast. Cell 116:405– 415 Harhangi HR, Akhmanova AS, Emmens R, van der Drift C, de Laat WTAM, van Dijken JP, Jetten MSM, Pronk JT, Op den Camp HJM (2003) Xylose metabolism in the anaerobic fungus Piromyces sp. strain E2 follows the bacterial pathway. Arch Microbiol 180:134–141 Hartmann T (2007) From waste products to ecochemicals: Fifty years research of plant secondary metabolism. Phytochemistry 68:2831–2846 Hedden P, Phillips AL, Rojas MC, Carrera E, Tudzynski B (2002) Gibberellin biosynthesis in plants and fungi: a case of convergent evolution? J Plant Growth Regul 20:319–331 Hemmi H, Ikejiri S, Nakayama T, Nishino T (2003) Fusion-type lycopene b-cyclase from a thermoacidophilic archaeon Sulfolobus solfataricus. Biochem Biophys Res Commun 305:586–591 Hiltunen JK, Wenzel B, Beyer A, Erdmann R, Fossa A, Kunau WH (1992) Peroxisomal multifunctional beta-oxidation protein of Saccharomyces cerevisiae. Molecular analysis of the fox-2 gene and gene product. J Biol Chem 267:6646–6653 Hittinger CT, Rokas A, Carroll SB (2004) Parallel inactivation of multiple GAL pathway genes and ecological diversification in yeasts. Proc Natl Acad Sci USA 101:14144–14149 Hof H (2008) Mycotoxins: Pathogenicity factors or virulence factors? Mycoses 51:93–94 Hoffmeister D, Keller NP (2007) Natural products of filamentous fungi: enzymes, genes, and their regulation. Nat Prod Rep 24:393–416 Hohn TM, McCormick SP, Desjardins AE (1993) Evidence for a gene cluster involving trichothecene-pathway biosynthetic genes in Fusarium sporotrichiodes. Curr Genet 24:291–295 Hottiger T, Boller T (1991) Ethylene biosynthesis in Fusarium oxysporum sp. tulipae proceeds from glutamate/ 2-oxoglutarate and requires oxygen and ferrous ions in vivo. Arch Microbiol 157:18–22 Hrmova M, Fincher GB (2001) Plant enzyme structure. Explaining substrate specificity and the evolution of function. Plant Physiol 125:54–57 Hsiao NH, Kirby R (2008) Comparative genomics of Streptomyces avermitilis, Streptomyces cattleya, Streptomyces maritimus and Kitasatospora aureofaciens using a Streptomyces coelicolor microarray system. A van Leeuw 93:1–25 Hurst LD, Williams EJ, Pal C (2002) Natural selection promotes the conservation of linkage of co-expressed genes. Trends Genet 18:604–606 Hurst, LD, Pal C, Lercher MJ (2004) The evolutionary dynamics of eukaryotic gene order. Nat Rev Genet 5:299–310 Jeffries TW (1983) Utilization of xylose by bacteria, yeast, and fungi. Adv Biochem Eng Biotechnol 27:1–32 Jekosch K, Ku¨ck U (2000) Glucose dependent transcriptional expression of the cre1 gene in Acremonium chrysogenum strains showing different levels of cephalosporin C production. Curr Genet 37:388–395 Johnson R, Voisey C, Johnson L, Pratt J, Fleetwood D, Khan A, Bryan G (2007) Distribution of NRPS gene

325

families within the Neotyphodium/Epichloe¨ complex. Fungal Genet Biol 44:1180–1190 Ka¨mper J, Kahmann R, Bo¨lker M, Ma L-J, Brefort T, Saville BJ, Banuett F, Kronstad JW, Mu¨ller O (2006) Insights from the genome of the biotrophic fungal plant pathogen Ustilago maydis. Nature 444:97–101 Kato N, Brooks W, Calvo AM (2003) The expression of sterigmatocystin and penicillin genes in Aspergillus nidulans is controlled by veA, a gene required for sexual development. Eukaryot Cell 2:1178–1186 Kawaide H (2006) Biochemical and molecular analyses of gibberellin biosynthesis in fungi. Biosci Biotechnol Biochem 70:583–590 Keller NP, Adams TH (1995) Analysis of a mycotoxin gene cluster in Aspergillus nidulans. SAAS Bull Biochem Biotechnol 8:14–21 Keller NP, Hohn TM (1997) Metabolic pathway gene clusters in filamentous fungi. Fungal Genet Biol 21: 17–29 Keller NP, Turner G, Bennet JW (2005) Fungal secondary metabolism: from biochemistry to genomics. Nat Rev Microbiol 3:937–947 Kennedy J, Auclair K, Kendrew SG, Park C, Vederas JC, Hutchinson CR (1999) Modulation of polyketide synthase activity by accessory proteins during lovastatin biosynthesis. Science 284:1368–1372 Khaldi N, Collemare J, Lebrun MH, Wolfe K (2008) Evidence for horizontal transfer of a secondary metabolite gene cluster between fungi. Genome Biol 9:R18 (doi:10.1186/gb-2008-9-1-r18) Kloer DP, Ruch S, Al-Babili S, Beyer P, Schulz GE (2005) The structure of a retinal-forming carotenoid oxygenase. Science 308:267–269 Kloer DP, Schulz GE (2006) Structural and biological aspects of carotenoid cleavage. Cell Mol Life Sci 63:2291–2303 Klutts JS, Yoneda A; Reilly MC, Bose I, Doering TL (2005) Glycosyltransferases and their products: cryptococcal variations on fungal themes. FEMS Yeast Res 6:499– 512 ¨ ber die chemische Zusammensetzung Ko¨ssel A (1891) U der Zelle. Du Bois-Reymond’s Archiv/Arch Anat Physiol Physiol Abt:181–186 Kremer A, Li SM (2008) Tryptophan aminopeptidase activity of several indole prenyltransferases from Aspergillus fumigatus. Chem Biol 15:729–738 Kremer A, Westrich L, Li SM (2007) A 7-dimethylallyltryptophan synthase from Aspergillus fumigatus: overproduction, purification and biochemical characterization. Microbiology 153:3409–3416 Kroken S, Glass NL, Taylor JW, Yoder OC, Turgeon BG (2003) Phylogenomic analysis of type I polyketide synthase genes in pathogenic and saprobic ascomycetes. Proc Natl Acad Sci USA 100:15670–15675 Kurihara T, Ueda M, Okada H, Kamasawa N, Anito N, Osumi M, Tanaka A (1992) Beta-oxidation of butyrate, the short-chain-length fatty acid, occurs in peroxisomes in the yeast Candida tropicalis. J Biochem 111:783–787 Kutil BL, Greenwald C, Liu G, Spiering MJ, Schardl CL, Wilkinson HH (2007) Comparison of loline alkaloid gene clusters across fungal endophytes: predicting the

326

Christine Schimek

co-regulatory sequence motifs and the evolutionary history. Fungal Genet Biol 44:1002–1010 Lacroix B, Tzfira T, Vainstein A, Citovsky V (2006) A case of promiscuity: Agrobacterium’s endless hunt for new partners. Trends Genet 22:29–37 Lawrey JD (1995) The chemical ecology of lichen mycoparasites: a review. Can J Bot 73:S603–S608 Li SM (2009) Evolution of aromatic prenyltransferases in the biosynthesis of indole derivatives. Phytochemistry 70:1746–1757 Li Y, Walton DC (1990) Violaxanthin is an abscisic acid precursor in water-stressed, dark-grown bean leaves. Plant Physiol 92:551–559 Linnemansto¨ns P, Prado MM, Fernandez-Martin R, Tudzynski B, Avalos J (2002) A carotenoid biosynthesis gene cluster in Fusarium fujikuroi: the genes carB and carRA. Mol Genet Genom 267:593–602 Liras P, Martin JF (2006) Gene clusters for b-lactam antibiotics and control of their expression: why have clusters evolved, and from where did they originate? Int Microbiol 9:9–19 Litzka O, Papagiannopoulos P, Davis MA; Hynes MJ, Brakhage AA (1998) The penicillin regulator PENR1 of Aspergillus nidulans is a HAP-like transcriptional complex. Eur J Biochem 251:758–767 Litzka O, Then Bergh K, Brakhage AA (1996) The Aspergillus nidulans penicillin-biosynthesis gene aat (penDE) is controlled by a CCAAT-containing DNA element. Eur J Biochem 238:675–682 Lorenz N, Haarmann T, Pazoutova S, Jung M, Tudzynski P (2009) The ergot alkaloid gene cluster: functional analyses and evolutionary aspects. Phytochemistry 70:1822–1832 Lorenz N, Wilson EV, Machado C, Schardl CL, Tudzynski P (2007) Comparison of ergot alkaloid biosynthesis gene clusters in Claviceps species indicates loss of late pathway steps in evolution of C. fusiformis. Appl Environ Microbiol 73:7185–7191 Ma LJ, Ibrahim AS, Skory C, Grabherr MG, Burger G, et al. (2009) Genomic analysis of the basal lineage fungus Rhizopus oryzae reveals a whole-genome duplication. PloS Genet 5:e1000549 Machida M, Machida M, Asai K, Sano M, Tanaka T, et al. (2005) Genome sequencing and analysis of Aspergillus oryzae. Nature 438:1157–1161 Madern D (2002) Molecular evolution within the L-malate and L-acetate dehydrogenase super-family. J Mol Evol 54:825–840 Maggio-Hall LA, Keller NP (2004) Mitochondrial betaoxidation in Aspergillus nidulans. Mol Microbiol 54:1173–1185 Malonek S, Rojas MC, Hedden P, Gaskin P, Hopkins P, Tudzynski B (2005a) Functional characterization of two cytochrome P450 monooxygenase genes, P450-1 and P450-2, of the gibberellic acid gene cluster in Fusarium proliferatum (Gibberella fujikuroi MP-D). Appl Environ Microbiol 71:1462–1472 Malonek S, Rojas MS, Hedden P, Hopkins P, Tudzynski B (2005b) Restoration of gibberellin production in Fusarium proliferatum by functional complementation of enzymatic blocks. Appl Environ Microbiol 71:6014–6025

Malonek S, Bo¨mke C, Bornberg-Bauer E, Rojas MC, Hedden P, Hopkins P, Tudzynski B (2005c) Distribution of gibberellin biosynthetic genes and gibberellin production in the Gibberella fujikuroi species complex. Phytochemistry 66:1296–1311 Mannhaupt G, Montrone C, Haase D, Mewes HW, Aign V, Hoheisei JD, Fartmann B, Nyakatura G, Kempken F, Maier J, Schulte U (2003) What’s in the genome of a filamentous fungus? Analysis of the Neurospora genome sequence. Nucl Acids Res 31:1944–1954 Marumo S, Katayama M, Komori E, Ozaki Y, Natsume M, Kondo S (1982) Microbial production of abscisic acid by Botrytis cinera. Agric Biol Chem 46:1967–1968 Moise AR, von Lintig J, Palczewski K (2005) Related enzymes solve evolutionary recurrent problems in the metabolism of carotenoids. Trend Plant Sci 10:178–186 Morris PF, Schlosser LR, Onasch KD, Wittenschlaeger T, Austin R, Provart N (2009) Multiple horizontal gene transfer events and domain fusions have created novel regulatory and metabolic networks in the oomycete genome. PLoS One 4:e6133 Nierman WC, Pain A, Anderson MJ, Wortman JR, Kim HS, et al. (2005) Genomic sequence of the pathogenic and allergenic filamentous fungus Aspergillus fumigatus. Nature 438:1151–1156 O’Brien PJ, Herschlag (1999) Catalytic promiscuity and evolution of new enzymatic activities. Chem Biol 6:91–105 Oritani T, Kiyota H (2003) Biosynthesis and metabolism of abscisic acid and related compounds. Nat Prod Rep 20:414–425 Oritani T, Yamashita K (1985) Biosynthesis of (+)-abscisic acid in Cercospora cruenta. Agric Biol Chem 49:245– 249 Pail M, Peterbauer T, Seiboth B, Hametner C, Druzhinina I Kubicek CP (2004) The metabolic role and evolution of L-arabinitol 4-dehydrogenase of Hypocrea jecorina. Eur J Biochem 271:1864–1872 Pal C, Hurst LD (2003) Evidence for co-evolution of gene order and recombination rate. Nat Genet 33:392–395 Papp B, Pal C, Hurst LD (2004) Metabolic network analysis of the causes and evolution of enzyme dispensability in yeast. Nature 429:661–664 Partida-Martinez LP, de Looss CF, Ishida K, Ishida M, Roth M, Buder K, Hertweck C (2007a) Rhizonin, the first mycotoxin isolated from the zygomycota, is not a fungal metabolite but is produced by bacterial endosymbionts. Appl Environ Microbiol 73:793–797 Partida-Martinez LP, Groth I, Schmitt I, Richter W, Roth M, Hertweck C (2007b) Burkholderia rhizoxinica sp. nov. and Burkholderia endofungorum sp. nov., bacterial endosymbionts of the plant-pathogenic fungus Rhizopus microsporus. Int J Syst Evol Microbiol 57:2583–2590 Patron NJ, Waller RF, Cozijnsen AJ, Straney DC, Gardiner DM, Nierman WC, Howlett BJ (2007) Origin and distribution of epipolythiodioxopiperazine (ETP) gene clusters in filamentous ascomycetes. BMC Evolut Biol 7:174 Payne GA, Brown MP (1998) Genetics and physiology of aflatoxin biosynthesis. Annu Rev Phytopathol 36:329–362

Evolution of Special Metabolism in Fungi: Concepts, Mechanisms, and Pathways Peck RF, Johnson EA, Krebs MP (2002) Identification of a lycopene beta-cyclase required for bacteriorhodopsin biogenesis in the archaeon Halobacterium salinarum. J Bacteriol 184:2889–2897 Pedley KF, Walton JD (2001) Regulation of cyclic peptide biosynthesis in a plant pathogenic fungus by a novel transcription factor. Proc Natl Acad Sci USA 98:14174–14179 Penalva MA, Moya A, Dopazo J, Ramon D (1990) Sequences of isopenicillin N synthetase suggest horizontal gene transfer from prokaryotes to eukaryotes. Proc Biol Sci 241:164–169 Perrin RM, Fedorova ND, Bok JW, Cramer RA, Wortman JR, Kim HS, Nierman WC, Keller NP (2007) Transcriptional regulation of chemical diversity in Aspergillus fumigatus by LaeA. PloS Pathog 3:508–517 Prado-Cabrero A, Scherzinger D, Avalos J, Al-Babili S (2007) Retinal biosynthesis in fungi: characterization of the carotenoid oxygenase CarX from Fusarium fujikuroi. Eukaryot Cell 6:650–657 Price MN, Arkin AP, Alm EJ (2006) The life-cycle of operons. PloS Genet 2:0859–0873 Price MS, Yu J, Nierman WC, Kim HS, Pritchard B, Jacobus CA Bhatnagar D, Cleveland TE, Payne GA (2006) The aflatoxin pathway regulator AflR induces gene transcription inside and outside of the aflatoxin biosynthetic cluster. FEMS Microbiol Lett 255: 275–279 Proctor RH, Hohn TM, McCormick SP, Desjardins AE (1995) Tri6 encodes an unusual zinc finger protein involved in the regulation of trichothecene biosynthesis in Fusarium sporotrichiodes. Appl Environ Microbiol 61:1923–1930 Rademacher W (1994) Gibberellin formation in microorganisms. Plant Growth Regul 15: 303–314 Ragan MA, Beiko RG (2009) Lateral genetic transfer: open issues. Phil Trans R Soc B 364:2241–2251 Rehmeyer C, Li W, Kusaba M, Kim YS, Brown D, Staben C, Dean R, Framan M (2006) Organization of chromosome ends in the rice blast fungus, Magnaporthe oryzae. Nucl Acids Res 34:4685–4701 Ricchetti M, Dujon B, Fairhead C (2003) Distance from the chromosome end determines the efficiency of double strand break repair in subtelomeres of haploid yeast. J Mol Biol 328:847–862 Richards TA, Dacks LB, Jenkinson JM, Thornton CR, Talbot NJ (2006) Evolution of filamentous plant pathogens: gene exchange across eukaryotic kingdoms. Curr Biol 16: 1857–1864 Richards TA, Soanes DM, Foster PG, Leonard D, Thornton CR, Talbot NJ (2009) Phylogenetic analysis demonstrates a pattern of rare an ancient horizontal gene transfer between plants and fungi. Plant Cell 21:1897– 1911 Rim SO, Lee JH, Choi WY, Hwang SK, Suh SJ, Lee IJ, Rhee IK, Kim JG (2005) Fusarium proliferatum KGL0401 as a new gibberellin–producing fungus. J Microbiol Biotechnol 15:809–814 Robyr D, Suka J, Xenarios I, Kurdistani SK, Wamd A, Suka N, Grundstein M (2002) Microarray deacetylation maps determine genome-wide functions for yeast histone deacetylases. Cell 109:437–446

327

Rohlfs M, Albert M, Keller NP, Kempken F (2007) Secondary chemicals protect mould from fungivory. Biol Lett 3:523–525 Rojas MC, Urrutia O, Cruz C, Tudzynski B, Hedden P (2004) Kaurenolides and fujenoic acid are side products of the P450-1 monooxygenase in Gibberella fujikuroi. Phytochemistry 65:821–830 Rosewich UL, Kistler HC (2000) Role of horizontal gene transfer in the evolution of fungi. Annu Rev Pathopathol 38:325–363 Saks V, Beraud N, Wallimann T (2008) Metabolic compartmentation – a system level property of muscle cells: real problems of diffusion in living cells. Int J Mol Sci 9:751–767 Sandmann G (2001) Carotenoid biosynthesis and biotechnological application. Arch Biochem Biophys 385:4–12 Sandmann G (2002) Combinatorial biosynthesis of carotenoids in a heterologous host: a powerful approach for the biosynthesis of novel structures. ChemBioChem 3:629–635 Schardl CL (2001) Epichloe¨ festucae and related mutualistic symbionts of grasses. Fungal Genet Biol 33:59–82 Schardl CL, Panacchione DG, Tudzynski P (2006) Ergot alkaloids – biology and molecular biology. Alkaloids Chem Biol 63:45–86 Schimek C, Petzold A, Schultze K, Wetzel J, Wolschendorf F, Burmester A, Wo¨stemeyer J (2005) 4-Dihydromethyltrisporate dehydrogenase, an enzyme of the sex hormone pathway in Mucor mucedo, is constitutively transcribed but its activity is differently regulated in (+) and () mating types. Fungal Genet Biol 42:804–812 Schimek C, Wo¨stemeyer J (2006) Pheromone action in the fungal groups Chytridiomycota, and Zygomycota, and in the Oomycota. In: Ku¨es U, Fischer R (eds) The Mycota, vol 1, 2nd edn, Springer, Berlin Heidelberg New York, pp 215–231 Schimek C, Wo¨stemeyer J (2009) Carotene derivatives in sexual communicaton of zygomycete fungi. Phytochemistry 70:1867–1875 Schmitt EK; Bunse A, Janus D, Hoff B, Friedlin E, Kurnsteiner H, Ku¨ck U (2004) Winged helix transcription factor CPCR1 is involved in regulation of beta-lactam biosynthesis in the fungus Acremonium chrysogenum. Eukaryot Cell 3:121–134 Schmitt EK, Hoff B, Ku¨ck U (2004) Regulation of Cephalosporin biosynthesis. Adv Biochem Engin Biotechnol 88:1–43 Schroeckh V, Scherlach K, Nu¨tzmann H-W, Shelest E, Schmidt-Heck W, Schuemann J, Martin K, Hertweck C, Brakhage AA (2009) Intimate bacterial–fungal interaction triggers biosynthesis of archetypal polyketides in Aspergillus nidulans. Proc Natl Acad Sci USA 106:14558–14563 Schultze K, Schimek C, Wo¨stemeyer J, Burmester A (2005) Sexuality and parasitism share common regulatory pathways in the fungus Parasitella parasitica. Gene 348:33–44 Schwab W (2003) Metabolome diversity: too few genes, too many metabolites. Phytochemistry 62:837–849 Seymour FA, Cresswell JE, Fisher PJ, Lappin-Scott HM, Haag H, Talbot NJ (2004) The influence of genotypic

328

Christine Schimek

variation on metabolite diversity in populations of two endophytic fungal species. Fungal Genet Biol 41:721–734 Shen YQ, Burger G (2009) Plasticity of a key metabolic pathway in fungi. Funct Integr Genomics 9:145–151 Shaaban M, Palmer JM, El-Naggar WA, El-Sokkary MA, Habib ESE, Keller NP (2010) Involvement of transposon-like elements in penicillin gene cluster regulation. Fungal Genet Biol 47: 423–432 Shwab EK, Bok JW, Tribus M, Galehr J, Graessle S, Keller NP (2007) Histone deacetylase activity regulates chemical diversity in Aspergillus. Eukaryot Cell 6:1656–1664 Shwab EK, Keller NP (2008) Regulation of secondary metabolite production in filamentous ascomycetes. Mycol Res 112:225–230 Siewers V, Kokkelink L, Smedsgaard J, Tudzynski P (2006) Identification of an abscisic acid gene cluster in the grey mold Botrytis cinerea. Appl Environ Microbiol 72:4619–4626 Slot JC, Hibbett DS (2007) Horizontal transfer of a nitrate assimilation gene cluster and ecological transitions in fungi: a phylogenetic study. PLoS One 10:e1097 Slot JC, Rokas A (2010) Multiple GAL pathway gene clusters evolved independently and by different mechanisms in fungi. Proc Natl Acad Sci USA 107:10136– 10141 Smith DJ, Burnham MKR, Bull JH, Hodgson JE, Ward JM, Browne P. Brown J, Barton B, Earl AJ, Turner G (1990) b-Lactam antibiotic biosynthetic genes have been conserved in clusters in prokaryotes and eukaryotes. EMBO J 9:741–747 Spro¨te P, Brakhage AA (2007) The light-dependent regulator velvet A of Aspergillus nidulans acts as a repressor of the penicillin biosynthesis. Arch Microbiol 188: 69–79 Spro¨te P, Hynes MJ, Hortschansky P, Shelest E, Scharf DH, Wolke SM, Brakhage AA (2008) Identification of the novel penicillin biosynthesis gene aatB of Aspergillus nidulans and its putative evolutionary relationship to this fungal secondary metabolism gene cluster. Mol Microbiol 70:445–461 Steidl S, Papagiannopoulos P, Litzka O, Andrianopoulos A, Davis MA, Brakhage AA (1999) AnCF, the CCAAT binding complex of Aspergillus nidulans contains products of the hapB, hapC and hapE genes and is required for activation by the pathway specific regulatory gene, amdR. Mol Cell Biol 19:99–106 Subramaniam SS, Nagalla SR, Renganathan V (1999) Cloning and characterization of a thermostable cellobiose dehydrogenase from Sporotrichum thermophile. Arch Biochem Biophys 365:223–230 Sutter RP, Dadok J, Bothner-By, AA, Smith RR, Mishra PK (1989) Cultures of separated mating types of Blakeslea trispora make D and E forms of trisporic acids. Biochemistry 28:4060–4066 Talbot NJ, Vincent P, Wildman HG (1996) The influence of genotype and environment on the physiological and metabolic diversity of Fusarium compactum. Fungal Genet Biol 20:254–267 Tanaka A, Shiotani H, Yamamoto M, Tsuge T (1999) Insertional mutagenesis and cloning of the genes

required for biosynthesis of the host-specific AK toxin in the Japanese pear pathotype of Alternaria alternata. Mol Plant-Microbe Interact 12:691–702 Tanaka A, Tapper BA, Popay A, Parker EJ, Scott B, Scott B (2005) A symbiosis expressed non-ribosomal peptide synthetase from a mutualistic fungal endophyte of perennial ryegrass confers protection to the symbiotum from insect herbivory. Mol Microbiol 57: 1036–1050 Tanaka T, Tateno Y, Gojobori T (2004) Evolution of vitamin B6 (pyridoxine) metabolism by gain and loss of genes. Mol Biol Evol 22:243–250 Teichmann SA (2002) The constraints protein–protein interactions place on sequence divergence. J Mol Biol 324:399–407 Temporini ED, Van Etten HD (2004) An analysis of the pea pathogenicity genes of Nectria haematococca MPVI supports the hypothesis of their origin by horizontal transfer and uncovers a potentially new pathogen of garden pea: Neocosmospora boniensis. Curr Genet 46: 29–36 Then Bergh K, Brakhage AA (1998) Regulation of Aspergillus nidulans penicillin biosynthesis gene acvA (pcbAB) by amino acids: Implication for involvement of transcription factor PACC. Appl Environ Microbiol 64:843–849 Thewes S, Prado-Cabrero A, Prado MM, Tudzynski B, Avalos J (2005) Characterization of a gene in the car cluster of Fusarium fujikuroi that codes for a protein of the carotenoid oxygenase family. Mol Genet Genom 274:217–228 Tilburn J, Sarkar S, Widdick DA, Espeso EA, Orejas M, Mongroo J, Penalva MA, Arst Jr HN (1995) The Aspergillus PacC zinc finger transcription factor mediates regulation of both acidic- and alkaline-expressed genes by ambient pH. EMBO J 14:779–790 Tokai T, Fujimura M, Inoue H, Aoki T, Ohta K, Shibata T, Yamaguchi I, Kimura M (2005) Concordant evolution of trichothecene 3-O-acetyltransferase and an rDNA species phylogeny of trichothecene-producting and non-producting fusaria and other ascomycetous fungi. Microbiology 151:509–519 Tsai HF, Wang H, Gebler JC, Poulter CD, Schardl CL (1995) The Claviceps purpurea gene encoding dimethylallyltryptophan synthase, the committed step for ergot alkaloid biosynthesis. Biochem Biophys Res Commun 216:119–125 Tsitsigiannis DI, Kowieski TM, Zarnowski R, Keller NP (2005) Three putative oxylipin biosynthetic genes integrate sexual and asexual development in Aspergillus nidulans. Microbiology 151:1809–1821 Tsitsigiannis DI, Keller NP (2006) Oxylipins act as determinants of natural product biosynthesis and seed colonization in Aspergillus nidulans. Mol Microbiol 59:882–892 Tudzynski B, Holter K (1998) Gibberellin biosynthetic pathway in Gibberella fujikuroi: evidence for a gene cluster. Fungal Genet Biol 25:157–170 Velayos A, Eslava AP, Iturriaga EA (2000) A bifunctional enzyme with lycopene cyclase and phytoene synthase activities is encoded by the carRP gene of Mucor circinelloides. Eur J Biochem 267:5509–5519

Evolution of Special Metabolism in Fungi: Concepts, Mechanisms, and Pathways Verdoes JC, Krubasik P, Sandmann G, van Ooyen AJJ (1999) Isolation and functional characterization of a novel type of carotenoid biosynthetic gene from Xanthophyllomyces dendrorhous. Mol Gen Genet 262:453–461 von Lintig J, Vogt K (2004) Vitamin A formation in animals: molecular identification and functional characterization of carotene-cleaving enzymes. J Nutr 134:251S–256S Walton JD (2000) Horizontal gene transfer and the evolution of secondary metabolite gene clusters in fungi: a hypothesis. Fungal Genet Biol 30:167–171 Walton JD (2006) HC-toxin. Phytochem Anal 67:1406– 1413 Ward TJ, Bielawski JP, Kistler HG, Sullivan E, O’Donnell K (2002) Ancestral polymorphism and adaptive evolution in the trichothecene mycotoxin gene cluster of phytopathogenic Fusarium. Proc Natl Acad Sci USA 99:9278–9283 Werkman BA (1976) Localization and partial characterization of a sex-specific enzyme in homothallic and heterothallic Mucorales. Arch Microbiol 109:209–213 Wetzel J, Scheibner O, Burmester A, Schimek C, Wo¨stemeyer J (2009) 4-Dihydrotrisporin-dehydrogenase, an enzyme of the sex hormone pathway of Mucor mucedo: purification, cloning of the corresponding gene, and developmental expression. Eukaryot Cell 8:88–95 Wink M (2003) Evolution of secondary metabolites from an ecological and molecular phylogenetic perspective. Phytochemistry 64:3–19 Woloshuk CP, Fount KR, Brewer JF, Bhatnagar D, Cleveland TE, Payne GA (1994) Molecular characterization of alfR, a regulatory locus for aflatoxin biosynthesis. Appl Environ Microbiol 60:2408–2414 Woloshuk CP, Prieto R (1998) Genetic organization and function of the aflatoxin B1 biosynthetic genes. FEMS Microbiol Lett 160:169–176 Wong S, Wolfe KH (2005) Birth of a metabolic gene cluster in yeast by adaptive gene relocation. Nat Genet 37:777–782 Wo¨stemeyer J, Schimek C (2007) Trisporic acid and mating in zygomycetes. In: Heitman J, Kronstad JW, Taylor JW, Casselton L (eds) Sex in fungi: molecular determination and evolutionary implications. ASM Press, Washington DC, pp 431–443 Wo¨stemeyer J, Wo¨stemeyer A, Burmester A, Czempinski K (1995) Relationships between sexual processes and parasitic interactions in the host–pathogen system Absidia glauca – Parasitella parastica. Can J Bo. 73: S243–S250 Xu M, Wilderman PR, Peters RJ (2007) Following evolution’s lead to a single residue switch for diterpene synthase product outcome. Proc Natl Acad Sci USA 104:7397–7401 Yabe K, Ando J, Hamaski T (1988) Biosynthetic relationships among aflatoxins B1, B2, G1 and G2. Appl Envı´ron Microbiol 54:2101–2106 Yabe K, Nakajima H (2004) Enzyme reactions and genes in aflatoxin biosynthesis. Appl Microbiol Biotechnol 64:745–755

329

Yamamoto H, Inomata M, Tsuchiya S, Nakamura M, Oritani T (2000a) Metabolism of chiral ionylideneacetic acids on the abscisic acid biosynthesis pathway in Cercospora. Biosci Biotechnol Biochem 64:2644–2650 Yamamoto H, Inomata M, Tsuchiya S, Nakamura M, Uchiyama T, Oritani T (2000b) Early biosynthetic pathway to abscisic acid in Cercospora cruenta. Biosci Biotechnol Biochem 64:2075–2082 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 Yin WB, Ruan HL, Westrich L, Grundmann A, Li SM (2007) CdpNPT, an N-prenyltransferase from Aspergillus fumigatus: overproduction, purification and biochemical characterization. ChemBioChem 8:1154–1161 Young C, McMillan L, Telfer E, Scott B (2001) Molecular cloning and genetic analysis of an indolediterpene gene cluster from Penicillium paxilli. Mol Microbiol 39:754–764 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, 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 Gen Genom 274:13–29 Yu JH, Butchko RA, Fernandes M, Keller NP, Leonard TJ Adams TH (1996) Conservation of structure and function of the aflatoxin regulatory gene aflR from Aspergillus nidulans and A. flavus. Curr Genet 29:549–555 Yu J, Chang PK, Ehrlich KC, Cary JW, Bhatnagar D, Cleveland TE, Payne GA, Linz JE, Woloshuk CP, Bennett JW (2004) Clustered pathway genes in aflatoxin biosynthesis. Appl Environ Microbiol 70:1253–1262 Zamocky M, Hallberg M, Ludwig R, Divne C, Haltrich D (2004) Ancestral gene fusion in cellobiose dehydrogenases reflects a specific evolution of GMC oxidoreductases in fungi. Gene 338:1–14 Zhang DX, Stromberg AJ, Spiering MJ, Schardl CL (2009) Coregulated expression of loline alkaloid-biosynthesis genes in Neotyphodium unicatum cultures. Fungal Genet Biol 46:517–530 Zhang S et al. (2004) An indole-diterpene gene cluster from Aspergillus xavus. Appl Environ Microbiol 70:6875–6883 Zhang S, Schwelm A, Jin H, Collins LJ, Bradshaw RE (2007) A fragmented aflatoxin-like gene cluster in the forest pathogen Dothistroma septosporum. Fungal Genet Biol 44:1342–1354 Zhou LQ, Xue GP, Orpin CG, Black GW, Gilbert HJ, Hazlewood GP (1994) Intronless Celb from the anaerobic fungus Neocallimastix patriciarum encodes a modular family A endoglucanase. Biochem J 297:359–364

.

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