Plant Genome: Biodiversity and Evolution: Lower Groups

PLANT GENOME: BIODIVERSITY AND EVOLUTION Volume 2, Part B Lower Groups

Plant Genome Biodiversity and Evolution Series Editors A.K. Sharma and Archana Sharma Department of Botany University of Calcutta, Kolkata India

Advisory Board J. Dolezel, Laboratory of Molecular Cytogenetics and Cytometry, Institute of Experimental Botany, Sokolovska, Czech Republic K. Fukui, Department of Biotechnology, Graduate School of Engineering, Osaka University, Osaka, Japan R.N. Jones, Institute of Biological Sciences, Ceredigion, Scotland, UK G.S. Khush, 416 Cabrillo Avenue, Davis, California 95616, USA Ingo Schubert, Institut fur Pflanzengenetik Kulturpflanzenforschung, Gatersleben, Germany Canio G. Vosa, Scienze Botanische, Pisa, Italy

Volume 1, 1, 2, 2,

Part Part Part Part

A B A B

: : : :

Phanerogams Phanerogams Lower Groups Lower Groups

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Plant Genome

Biodiversity and Evolution Volume 2, Part B Lower Groups

Editors

A.K. SHARMA and A. SHARMA

Science Publishers Enfield (NH)

Jersey

Plymouth

SCIENCE PUBLISHERS An Imprint of Edenbridge Ltd., Channel Islands, British Isles.

Post Office Box 699 Enfield, New Hampshire 03748 Website: http://www.scipub.net [email protected] (marketing department) [email protected] (editorial department) [email protected] (for all other enquiries) ISBN 1-57808-413-x © 2006, Copyright reserved All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying or otherwise, without the prior permission. This book is sold subject to the condition that it shall not by way of trade or otherwise be lent, re-sold, hired out, or otherwise circulated without the publisher’s prior consent in any form of binding or cover other than that in which it is published and without a similar condition including this condition being imposed on the subsequent purchaser. Published by Science Publishers, Inc. Enfield, NH, USA An Imprint of Edenbridge Ltd. Printed in India

Preface to the Series “Plant Genome”

The term genome, the basic gene complement of an individual, is almost synonymous with the chromosome complement of both nucleus and organelles. Refinements in cellular, genetic and molecular methods in recent years have opened up unexplored avenues in genome research. The modern tools of gene and genome analyses, coupled with analysis of finer segments of gene sequences in chromosomes utilizing molecular hybridization, are now applied on a wider scale in different groups of plants, ranging from algae to angiosperms. This synergistic approach has made the study of biodiversity highly fascinating, permitting a deep insight into the molecular basis of genetic diversity. Simultaneous to the enrichment of fundamentals in systematics and phylogeny, the plant system, because of its inherent flexibility, has permitted genetic engineering and horizontal transfer of genes with immense importance in agriculture, horticulture and medicine. Despite the fact that the data on plant genomics with its impact on the assessment of biodiversity and evolution show a logarithmic increase, a comprehensive series on the aspect covering all groups of plant kingdom is sadly lacking. In view of this lacuna, the present series on Plant Genomics: Biodiversity and Evolution has been planned. It aims to cover, in successive volumes, comprehensive reviews, concepts and

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discussions on the results of genome analysis and their impact on systematics, taxonomy, phylogeny and evolution of all plant groups. We have not gone out of our way to seek original articles, but in course of reviews and discussions, research articles, if any, are welcome. February, 2003

A.K. Sharma Archana Sharma Series Editors

Preface to this Edition

In the previous volume of this series on Lower Groups, algal genomics was represented in Rhodophyceae in detail. Genomics of fungi was presented in the phytopathogen Leptosphaerea, powdery mildew and oyster mushroom. In addition, comparative genomics of several phytopathogens and genomc redundancy in yeast were critically reviewed. The genomics of lichens, liverwort sex chromosomes, the functional genomics of mosses and organellar transcription of Bryophytes formed the core matter of the other chapters. The coverage of the present volume which fills up the lacuna of and is supplementary to the earlier one is very wide, starting with the most emerging study in Genomics—the RNA world–ancient to modern–a metabolic story in detail. The structural features of all the different ribozymes and their functioning as well as all the non coding RNAs–their role, chemical basis of action and catalytic mechanisms have been critically reviewed in the backdrop of genome structure and evolution. The subsequent chapters on genomics of algae deal with groups which are of great phylogenetic significance and biodiversity. These include origin of plastids, diversity, origin and evolution of dinoflagellates and diatoms from the molecular perspective. The fungal genomics is relation to phylogeny has been covered in hemi, archi and euascomycete with special reference to comparative

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genomics and evolution, The biodiversity in basidiomycetous yeast such as Ustilaginales, Uredinales and Hymenomycetes has been critically reviewed in terms of DNA sequence analysis. In the yeast group, a special feature is the analysis of cactophilic yeast growing on decaying cactus tissue, from an evolutionary and ecological standpoint. The molecular data of these habitat and insect specific yeast show clear interstrain and interspecific variations and throw light on the intercommunity relationship. The molecular phylogeny of the genus Aspergillus with the two sub genera–the source of drugs and enzymes has been presented in detail along with the evolution of mycotoxin production, which is presumed to have evolved several times in course of evolution. The genomic study of Fusarium – a highly destructive fungus has been focussed towards its phylogenetic analysis and its detection using molecular tools. The host pathogen co–evolution with the aid of nuclear DNA and microsatellite markers in two co–colonizing fungi Philaphora and Plectosporium and a negative correlation between the two pathogens could be established. The articles on Bryophytes deal with a comprehensive analysis on role of nuclear and organellar DNA in evolution and biodiversity. The importance of ITS spacers of nuclear ribosomal DNA and the role of intergenic spacers of chloroplast DNA in the analysis of phylogeny and plant systematics and evolution of this group have been discussed. In Plagiochila–the Jungermaniales, the data on phylogeny and interactions between species of intercontinental ranges have been presented. The molecular evidences indicate interconnecting link between the taxa of neotropics and Africa with the Atlantic, Europe and the Americas and the Asian Plagiochila with the Holarctic. This volume thus provides an in depth and comprehensive analysis on genomics in evolution from the ancient RNA world to the modern day land plants passing through all hierarchy of algae, fungi and bryophytes. It will be of immense use to all engaged in the molecular studies of different groups with their impact on genetics, evolution and agriculture. Arun Kumar Sharma Archana Sharma

Contents

Preface to the Series Preface to this Edition List of Contributors 1. From Ancient to Modern RNA world: A Metabolic Story Yan Li Li, Sylvia Tobé, Annabelle Chabauty and Marie-Christine Maurel

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2. Plastid Origin: A Driving Force for the Evolution of Algae Alexis Miguel Bellorin and Mariana Cabral de Oliveira

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3. Evolution and Diversity of Dinoflagellates: Molecular Perspectives Michael J. Bennett and Joseph T.Y. Wong

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4. Evolution of the Diatoms Wiebe H.C.F. Kooistra, Victor Chepurnov, Linda K. Medlin, Mario De Stefano, Koen Sabbe and David G. Mann 5. Ascomycota: Introduction to Biodiversity, Evolutionary Genomics and Systematics Wolfgang Schweigkofler

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6. Yeast Biodiversity and Evolution Vu Nguyen Thanh 7. Evolutionary Relationships among Economically Important Species of Aspergillus Subgenera Aspergillus and Fumigati János Varga, Krisztina Rigó, Sándor Kocsubé, Károly Pál, Beáta Tóth, Robert A. Samson and Zotia Kozaki Ewicz 8. Polymerase Chain Reaction-based Methods in Fusarium Taxonomy Youssuf A.M.H. Gherbawy, Mohamed A.El-Naghy and Aabdullah Altalhi 9. Use of Molecular Markers to Study Host-pathogen Co-evolution Weidong Chen and Craig R. Grau 10. Utility of the Internal Transcribed Spacers of the 18S-5.8S-26S Nuclear Ribosomal DNA in Land Plant Systematics, with Special Emphasis on Bryophytes Alain Vanderpoorten, B. Goffinet and D. Quandt

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11. Molecular Evolution of the atpB-rbcL Intergenic Spacer in Bryophytes Michael Stech and Dietmar Quandt

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12. Molecular Phylogeny and Biogeography of Plagiochila (Jungermanniidae: Plagiochilaceae) Jochen Heinrichs

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13. Genomes Studies of Cactus Yeast Miguel de Barros Lopes and Philip F. Ganter

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Author Index

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Detailed Contents

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List of Contributors

Yan Li Li Biochemistry of Evolution and Molecular Adaptability Laboratory, Jacques-Monod Institute-CNRS, Pierre-et-Marie Curie, Paris 6, Paris, France Sylvia Tobé Biochemistry of Evolution and Molecular Adaptability Laboratory, Jacques-Monod Institute-CNRS, Pierre-et-Marie Curie, Paris 6, Paris, France Anabelle Chabauty Biochemistry of Evolution and Molecular Adaptability Laboratory, Jacques-Monod Institute-CNRS, Pierre-et-Marie Curie, Paris 6, Paris, France Marie-Christine Maurel Biochemistry of Evolution and Molecular Adaptability Laboratory, Jacques-Monod Institute-CNRS, Pierre-et-Marie Curie, Tower 43, 2 Place Jussieu, 75251 Paris Cedex 05, France; E-mail: [email protected] Alexis Miguel Bellorin Dept. Biologia, Escuela de Ciencias, University of Oriente (UDO), Av. Universidad, Cumana, Venezuela AP 245; E-mail: [email protected] Mariana Cabral de Oliveira Dept. Botânica, Inst. De Biociências, University of São Paulo (USP), R. do Matão, travessa 14 n. 321, São Paulo, SP cep 05508-900, Brazil; E-mail: [email protected], [email protected]

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Michael J. Bennett Department of Biology, Hong Kong University of Science and Technology, Clearwater Bay, Kowloon, Hong Kong SAR, People’s Republic of China Joseph T.Y. Wong Department of Biology, Hong Kong University of Science and Technology, Clearwater Bay, Kowloon, Hong Kong SAR, People’s Republic of China; E-mail: [email protected] Weibe H.C.F. Kooistra Stazione Zoologica “Anton Dohrn,” Villa Comunale, I-80121, Naples, Italy; E-mail: [email protected] Victor Chepurnov Laboratory for Protistology and Aquatic Ecology, Department of Biology, Ghent University, Krijgslaan 281 – S8, B-9000 Ghent, Belgium Linda K. Medlin Alfred-Wegener –Institute for Polar and Marine Research, Am Handelshafen 12, D-27570 Bremerhaven, Germany Mario De Stefano Stazione Zoologica “Anton Dohrn,” Villa Comunale, I-80121, Naples, Italy Koen Sabbe Laboratory for Protistology and Aquatic Ecology, Department of Biology, Ghent University, Krijgslaan 281 – S8, B-9000 Ghent, Belgium David G. Mann Royal Botanic Garden, Edinburgh EH3 5LR, Scotland, UK Wolfgang Schweigkofler Department of Plant Protection, Research Center for Agriculture and Forestry, Laimburg, I-39040 Auer (BZ), Italy; E-mail: [email protected] Vu Nguyen Thanh Department of Microbiology, Food Industries Research Institute, 301 Nguyen Trai, Thanh Xuan, Hanoi, Vietnam; E-mail: [email protected]

List of Contributors

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János Varga Department of Microbiology, Faculty of Sciences, University of Szeged, P.O. Box 533, H-6701 Szeged, Hungary; E-mail: [email protected] Krisztina Rigó Department of Microbiology, Faculty of Sciences, University of Szeged, P.O. Box 533, H-6701 Szeged, Hungary Sándor Kocsubé Department of Microbiology, Faculty of Sciences, University of Szeged, P.O. Box 533, H-6701 Szeged, Hungary Károly Pál Department of Microbiology, Faculty of Sciences, University of Szeged, P.O. Box 533, H-6701 Szeged, Hungary Beáta Tóth Cereal Research non-Profit Company, Szeged, Hungary Robert A. Samson Centraalbureau voor Schimmelcultures, Utrecht, The Netherlands Zofia Kozaki - Ewics CABI Bioscience, UK Centre, Bakeham Lane, Surrey 9TY, UK Youssuf A.M.H. Gherbawy Botany Department, Faculty of Sciences, South Valley University, Qena 83523, Egypt; E-mail: [email protected] Mohammed A.El-Naghy Botany Department, Faculty of Science, El-Minia University, ElMinia, Egypt Abdullah Altalhi Biology Department, Faculty of Science, Taif Universy, Taif, Saudi Arabia Weidong Chen USDA_ARS, Grain Legume Genetics and Physiology Research, 303 Johnson Hall, Washington State University, Pullman, WA 99164, USA; E-mail: [email protected]

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Craig R. Grau Department of Plant Pathology, University of Wisconsin, 1607 Linden Drive, Madison, WI 53706, USA Alain Vanderpoorten Research Associate of the Belgian Funds for the Scientific Research at University of Liège, Department of Life Sciences, B-22 Sart Tilman, B-4000 Liège, Belgium; E-mail: [email protected] Bernard Goffinet University of Connecticut, Department of Ecology and Evolutionary Sciences, Storrs, CT, USA Dietmar Quandt Institute of Botany, Technical University of Dresden, Zellescher Weg 22, D-01062 Dresden, Germany Michael Stech Institut für Biologie – Systematische Botanik und Pflanzengeography, Freie Universität Berlin, Altensteinstraße 6, D-14195, Berlin, Germany; E-mail: [email protected] Jochen Heinrichs Department of Systematic Botany, Albrecht von Haller Institute of Plant Sciences, Georg August University, Untere Karspüle 2, D37073 Gottingen, Germany; E-mail: [email protected] Miguel de Barros Lopes School of Pharmacy and Medical Sciences, University of South Australia, City East Campus, North Terrace, Adelaide, SA 5000, Australia Philip F. Ganter Biology Department, Tennessee State University, 3500 John Merritt Blvd., Nashville, TN 37209, USA

CH AP TE R



From Ancient to Modern RNA World: A Metabolic Story YAN LI LI, SYLVIA TOBÉ, ANNABELLE CHABAUTY and MARIE-CHRISTINE MAUREL * Biochemistry of Evolution and Molecular Adaptability Laboratory, Jacques-Monod Institute-CNRS, Pierre-et-Marie-Curie, Paris 6, Paris, France

ABSTRACT The RNA world theory suggests that modern life arose from molecular ancestors in which RNA molecules stored genetic information as well as catalyzed chemical reactions. This theory is largely supported by the importance of RNA molecules in essential contemporary cellular processes: RNA is involved in the decoding of genetic information, DNA replication, and chromosome-end maintenance; it mediates interference to defend cells against molecular parasites, and an increasing number of non-coding RNAs have recently been discovered that are involved in various biological activities. In addition, individual ribonucleotides and their coenzyme derivatives are highly involved in central metabolism. Moreover, the discovery of catalytic RNAs strongly reinforced the RNA world hypothesis. Seven naturally occurring classes of ribozymes have been identified till date, including the four classes of small self-cleaving RNAs. The latter are found on plant pathogenic RNA genomes, and are essentially involved in the pathogenicity by mediating the viral replication. This chapter succinctly presents the arguments in favor of the ancient RNA world, followed by a description of the world of non-coding RNAs and natural ribozymes, and finally focuses on small self-cleaving RNAs by manifesting their structural and functional characteristics. Key Words: RNA world, ribozymes, non-coding RNAs, small self-cleaving RNAs * Address for correspondence: Marie-Christine Maurel, Biochemistry of Evolution and Molecular Adaptability Laboratory, Jacques-Monod Institute-CNRS, Pierre-et-Marie-Curie, Tower 43, 2 Place Jussieu, 75251 Paris Cedex 05, France. Tel: 33-1-44-27-40-21, Fax: 33-144-27-99-16 E-mail: [email protected].

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INTRODUCTION

In 1986, Walter Gilbert established the RNA world hypothesis, according to which a single molecular species, the RNA molecule, was able to carry out informational and catalytic activities at the same time [25, 50, 106, 150]. In this theory, modern life arose from molecular ancestors in which RNA molecules stored genetic information, as well as catalyzed chemical reactions [51, 95]. Evidences for the “RNA world”, an episode of life on Earth during which RNA was the genetic component of bio-catalysts, are found in contemporary metabolism [89, 90], natural ribozymes [37], and in the ribosome [107, 156]. From a fundamental point of view, the RNA world hypothesis gains support from the wide implication of RNA in the most ancient and conserved cellular processes: the RNA molecule is involved in a very large range of functions in the contemporary biology of the three domains of life (Fig. 1). In living cells, desoxyribonucleotides, necessary to DNA synthesis, are continuously synthesized from ribonucleotides. The DNA specific base, thymine, is obtained by methylation of a RNA specific base: uracil. Furthermore, the DNA replication process starts with the synthesis of a RNA primer, suggesting that DNA replication could be a “modified” transcription process. The checkpoints in the decoding of genetic information are performed by mRNA, rRNA and tRNA, and RNA itself catalyzes the processing of precursor messenger RNAs. In addition, RNA has recently been shown to mediate protein synthesis, and it also plays a role in chromosome-end maintenance (telomeres). RNA interference and other RNA silencing phenomena reflect an elaborate cellular apparatus that eliminates defective mRNAs and defends against molecular parasites, such as transposons and viruses. Finally, individual ribonucleotides are essential signalling molecules and their coenzyme derivatives are highly involved in central metabolism [17, 65, 72] (Fig. 2). Beyond the wide range of RNA natural functions, the RNA world hypothesis was recently reinforced by the increasing number of artificial ribozymes obtained by in vitro evolution: the repertoire of chemical reactions catalyzed by RNA seems far more promising than previously anticipated [18, 54, 91, 132]. It is now usual to point out the modern RNA world: from the ribosome to the spliceosome, from viruses to catalytic RNAs.

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Fig. 1.

Phylogenetic distribution of RNAs.

FAD CoA-SH

NADP NAD

Fig. 2.

+

+

Essential coenzymes derived from ribonucleotides involved in central metabolism.

Seven naturally occurring classes of ribozymes have been identified to date: the four classes of the small self-cleaving RNAs family, group I and II introns, and Ribonuclease P. RNA is also the only catalyst in the ribonucleoprotein enzymes of the ribosome [107, 156] and within the spliceosome [105, 143].

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Most of the small self-cleaving RNAs are found on RNA viruses, satellite RNAs and viroïds. Although the details of some of these plant pathogenics biological functions are still not clear, they constitute excellent tools to study several biochemical mechanisms, such as ribozymic activities during replication. This chapter covers the following: (1) brief presentation of the non-coding RNAs and their metabolic functions; (2) focus on the natural ribozymes, particularly small endonucleolytic RNAs involved in plant viruses pathogenicity; and (3) description of their individual structural and catalytic properties. NON-CODING RNAS

For quite a long time, mRNA, rRNA and tRNA were thought to be the only existing types of RNA, all involved in protein synthesis. However, during the last few years, a number of small noncoding RNAs have been discovered, as presented below [38, 88]. The Small Nuclear RNAs (snRNAs)

Zieve, in 1981, isolated several RNAs which were different from canonical RNAs. They were rich in uridine, and were thus first called URNAs. In fact, these RNAs are associated to proteins in order to compose a Ribonucleoprotein (RNP) complex in the nucleus that is involved in splicing of mRNAs. The URNAs are thus spliceosome components, namely, U1, U2, U4, U5 and U6 small nuclear RNAs (snRNA). They play a key role in the recognition of the exon/intron junctions and in the splicing specificity [159]. The Small Nucleolar RNAs (snoRNAs)

The snoRNAs are biochemically isolated from nucleolar parts. They are 70 to 250 nucleotides in length, and play a key role in rRNAs processing and modification by guiding specific enzymes through their complementarity with target sequences. These RNAs are separated into two families, the C/D box and H/ACA box RNAs. The C/D box snoRNAs family is involved in site-specific 2’-O-ribose methylations of rRNAs. The reaction is catalysed by an associated methylase protein. The H/ACA snoRNA family, on the other hand, is

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involved in site-specific pseudouridylation of rRNAs. The catalytic function depends on an associated pseudo-U synthetase [39]. The snoRNAs have also been shown to mediate some modifications in tRNA and in snRNA. The Transfer Messenger RNAs (tmRNAs)

The tmRNAs are alanine tRNAs-like found only in procaryotes. They play a role in “non stop mRNA decay”. When a mRNA lacks a stop codon, the ribosome remains blocked on it. To solve this problem in procaryote cells, a tmRNA enters the P site of the ribosome, bringing an alanine that will be linked onto the nascent peptide. The tmRNA contains a specific coding frame that is then read and translated by the ribosome. This allows the formation of a chimeric protein that will be digested by proteases (Fig. 3). This mechanism is essential for bacterial cells: it allows the release of the blocked ribosome and makes it available for a new translation cycle [53, 67]. Blocked ribosome

alanine

reading frame

tmRNA 1

2

3

4

5

Fig. 3. Mechanism of action of tmRNAs.

Chimeric protein

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The RNA Signal Recognition Particle (RNA-SRP)

The RNA-SRP (or 7SL) is located in the signal recognition particle (SRP) complex. This complex is conserved in all types of cells. It allows the translocation of secreted proteins across the endoplasmic reticulum membrane in Eukarya, and the plasmic membrane in Archaebacteria and Eubacteria. The SRP recognizes a signal sequence on the nascent peptide and binds it. This complex interacts with the SR receptor on the membrane, allowing the protein to cross it [83, 147]. According to phylogenetic studies, the SRP was present in LUCA (Last Unique Common Ancestor) [89]. Vault RNA

The vault RNA belongs to the vault ribonucleoprotein particle (vRNP). Even if its function remains unclear, it is strongly suggested that this complex plays an important role in compartmentation and nuclear transport [145]. The Small Interfering RNAs (siRNA)

The siRNAs are small RNAs (21-23 nucleotides) that interfere with specific mRNAs, thus regulating target gene expression by inhibiting translation or altering mRNAs stability [141, 157]. They were first discovered in plants in 1990 by the Napoli team [101]. With the objective of increasing the coloration of petunia flowers, they inserted a transgene coding for chalcone synthase, an enzyme involved in the anthocyanin biosynthesis. Against all expectations, they obtained prominent proportion of purple-white flowers; instead of an increased amount of purple pigment, they observed its disappearance. This experience showed that there was a suppression of transgene and endogenous gene expression. This phenomenon was called PTGS (Post Transcriptional Gene Silencing). In 1994, research demonstrated that it occurred through deletion of the non endogenous-genes mRNA products [30]. While RNA interference (RNAi) was first observed in plants, it was later discovered in Caenorhabditis elegans by Fire [47, 135], which allowed a better understanding of the RNAi mechanism. During the first step of the process, the dsRNA ‘input is processed’ into small interfering RNAs of about 21-23 nt in length by a ribonuclease: DICER. These primary

Yan Li Li et al.

Fig. 4.

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Mechanism of RNA interference.

siRNAs are then recruited by a ribonuclease complex, RISC (RNA Induced Silencing Complex), which in turn mediates the cleavage of the target mRNA [58, 94, 146] (Fig. 4). The Micro RNAs (miRNAs)

The miRNAs play a role in RNA interference as well. They are transcribed from endogenous genes, are single-stranded and 21 to 25 nt long. They are involved in the spatial and temporal regulation of gene expression, particularly in response to stress answer or at specific cellular cycle stages. Thus, their expression varies at various developmental stages and in different tissues. MiRNAs operate the same way as siRNAs [78, 81]. The Small Temporal RNAs (stRNA)

The stRNAs are expressed at specific stages in development. The first stRNAs discovered, LET-7 and LIN-4, were genetically identified in the nematode Caenorhabditis elegans . They are respectively 21 and 22 nt in length, but are obtained from longer precursors transcribed from the

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LET-7 and LIN-4 heterochromatic genes. They are post-transcriptional negative expression regulators of genes encoding proteins such as Lin-41 and Lin-42 (for LET-7), Lin-14 and Lin-28 (for LET-4) [115]. The X Inactive-Specific Transcript RNAs (Xist RNA)

Dosage compensation is a remarkable example of connection between gene regulation and chromosome architecture. This system evolved so that the expression of genes carried by the X chromosome is identical in males and females. Thus, in placental female mammals, one of the two Xs is inactivated randomly. Sex-specific factors are responsible for X inactivation by regulating the expression of chromatinic structure genes (Fig. 5). The X inactivation takes place following X chromosome “counting” by the blocking factors: these factors are just in sufficient amount to prevent the inactivation of one X chromosome. The inactivation of the other chromosome starts at the X inactivation center (Xic), recognized by the Xist RNA (X inactive-specific transcript). Specific proteins then bind to the Xist RNA so that the inactivation can spread [3, 21, 153]. Xic X chromosomes Xist gene 1

Blocking Factor

Xist RNA 2

3

Protein

Fig. 5.

Mechanism of X chromosome inactivation by Xist RNA (See text for details).

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MAJOR NATURALLY OCCURRING RIBOZYMES

Naturally occurring ribozymes can be divided into seven groups on the basis of their functions and mechanisms : the ribonuclease P, the group I and II self-splicing introns, the hairpin, hammerhead, Varkud satellite and hepatitis delta virus ribozymes that constitute the small self-cleaving RNAs family. The Ribonuclease P (RNase P)

The ribonuclease P, discovered by Altman in 1982, is a ribonucleoprotein complex found in all cells [114]. It catalyzes a site-specific hydrolysis responsible for the maturation of precursors of tRNAs, 5S rRNAs and the signal recognition particle RNAs. It is considered a true catalyst because it catalyzes multiple turnover trans-reactions [49, 97]. The RNA component of RNase P is conserved in all organisms and is responsible, at least in bacteria, for catalytic activity. It is about 300 to 400 nt in length, and comprises two domains forming the substraterecognition site and the ribozyme active site. The human RNase P complex is larger, comprises multiple protein components, and its RNA is not catalytically active in the absence of protein. The respective implications of RNA and/or proteins in its catalysis, therefore, remain unclear [84, 154]. The Class I Introns

The self-splicing in vivo of both classes I and II introns occurs in presence of associated proteins. However, in vitro, the reaction can take place without any protein. The first ribozymes discovered were the group I introns, identified in the Cech laboratory (Univ. of Colorado) in 1982 [71]. They are present in organelle genes and nuclear ribosomic genes of several protozoans (algae, fungi, Eubacteria, bacteriophages, and eucaryotic viruses). This intron class utilizes an exogenous guanosine cofactor in catalysis. The splicing reaction requires two successive transesterifications. First, the 3'-OH of the exogenous guanosine acts as a nucleophile, attacks the phosphate at the 5' exon-intron junction, and covalently binds to the excised intron. This step requires metal ions for folding and catalysis. To complete splicing, the 3'-OH of the released 5'exon then attacks the phosphate at the 3' junction. After reaction, the exons are ligated and a free linear intron, with the G nucleoside attached at its 5' end, is released. This intron molecule eventually self-circularizes after splicing [19, 20] (Fig. 6A).

Fig. 6. Mechanism of group I (A) and group II (B) self-splicing introns.

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The Class II Introns

The class II introns have been described in organelle genes of fungi, algae, protists, plants, and cyanobacteria. Self-splicing of these introns occurs via a lariat intermediate and two transesterification reactions similar to the ones occurring for nuclear pre-mRNA. In the first step, a bulged adenosine attacks the 5' splice site, resulting in cleavage of the 5' exon and formation of a lariat intermediate. In the second step, the 5' and 3' exons are ligated together and the intron is released as a lariat (Fig. 6B) [20, 82]. THE SMALL SELF-CLEAVING RNAs FAMILY

The ribozymes of the small self-cleaving RNAs family are 50-150 nucleotides long, naturally present in viral, virusoid, or satellite RNA genomes (Table 1). They include the hammerhead, hairpin, Varkud satellite, and Hepatitis delta virus ribozymes [11, 34, 137]. Viroids, viroid-like and some satellite RNAs are known to follow the same replication pathway, called the rolling circle replication mechanism (Fig. 7). The small autocatalytic sequences harbored by viral and virusoid genomes are thought to catalyze the self-cleavage and self-ligation reactions producing the monomeric linear and circular intermediates of this replication process [8, 9, 33, 36, 133, 134]. The cleavage reaction catalyzed by such ribozymes is a site-specific reversible transesterification that follows an associative nucleophilic substitution (SN2) pathway, inducing an inversion of configuration of the scissile phosphate. The reaction starts with a nucleophilic attack of phosphorus by the adjacent 2’-oxygen, resulting in the formation of a pentavalent trigonal bipyramidal transition state. Rupture of the 5’oxygen-phosphorus bond generates products with 5’-hydroxyl and 2’-3’cyclic phosphate termini [41] (Fig. 8). The method by which reaction is catalyzed by the small self-cleaving ribozymes remains unclear; catalysis could occur through deprotonation of the 2’-hydroxyl group, facilitation of the trajectory into the in-line transition state, charge stabilization in the transition state, or stabilization of the cleaving group. It could involve the nucleobases as acid-base catalysts, or use metal ions in an acid-base and/or electrophilic catalysis [123].

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Table 1. Summary of the distribution of small self-cleaving ribozymes among viroids and other related RNAs RNA species Viroids Avocado sunblotch viroid (ASBVd) Peach latent mosaic viroid (PLMVd) Satellite RNAs Luteovirus Barley yellow dwarf virus satellite RNA (sBYDV) Nepovirus Arabis mosaic virus satellite RNA (sARMV) Chicory yellow mottle virus satellite RNA S1 (sCYMV) Tobacco ringspot virus satellite RNA (sTRSV) Sobemovirus Lucerne transient streak virus satellite RNA Subterranean clover mottle virus satellite RNA Solanum nodiflorum mottle virus satellite RNA Velvet tobacco mottle virus satellite RNA Other related RNAs Carnation stunt associated viroid Newt satellite 2 transcript Hepatitis delta virus Varkud Satellite*

Catalytic motifs Positive strand Negative strand Hammerhead Hammerhead

Hammerhead Hammerhead

» Hammerhead

Hammerhead

Hammerhead Hammerhead

Hairpin Hairpin

Hammerhead

Hairpin

Hammerhead Hammerhead Hammerhead Hammerhead

Hammerhead

Hammerhead Hammerhead Hammerhead Delta Delta Varkud

*: The VS RNA is not a virus satellite: it is transcribed from the VS DNA of Neurospora mitochondria. » hammerhead: for the sequence flanking the cleavage site in the encapsidated (+) strand of sBYDV sits most of the consensus rules for formation of a hammerhead structure, but differs from other naturally occurring hammerheads.

Although small self-cleaving ribozymes have very similar functions and generate the same reaction products, they do not share any structural characteristics. Comparison of their respective biochemical requirements also shows that ribozymes, like protein enzymes, can exploit more than one catalytic strategy. Each ribozyme adopts a unique structure and follows distinct kinetic and catalytic pathways, as if each motif was a different evolutionary answer for the same biological function [137].

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Fig. 7. Rolling circle replication mechanism. A: Symmetric pathway: the circular positive RNA strand is copied into an oligomeric negative strand, which is cleaved at specific sites to give unit-length negative strands. These linear molecules are then circularized and transcribed into oligomeric positive strands, processed as the formation of unit length negative strands and circularized to produce the progeny genomes. B: Asymmetric pathway: the rolling circle transcription occurs only once. Adapted from Symmons et al., 1987 [134].

O

R1

O

B

O

R1

OH

O

O

R1

B

O

O

B

O P

O

P O

O R2

OH

O

O

O

O B

O

Fig. 8.

O

+ P

O

B

O

O

OH

R2

OH B

O

O

OH

R2

Reaction mechanism of small ribozymes self-cleavage [41].

In laboratories, because of their small size and relative simplicity, small self-cleaving RNAs are especially suitable to RNA structurefunction studies, and may help to understand how RNA sequences can assemble to form functional structures, and the individual roles played by each specific group in the steps of assembly and catalysis.

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Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

The Hairpin Ribozyme

The hairpin ribozyme was first observed in the negative strand of the tobacco ringspot virus satellite RNA ((-)sTRSV) [14, 15, 144], and has since then also been found in two other pathogenic plant satellite viruses: the arabis mosaic virus and the chicory yellow mottle virus satellites (sARMV and sCYMV) [60, 137]. It can catalyze both cleavage and ligation reactions efficiently. Ligation is a simple reversal of the cleavage reaction, with the 5’-oxygen being the nucleophile [43] (Fig. 8). This ligation activity of the ribozyme suggests that it mediates both the processing of linear multimers and the circularization of monomers during in vivo rolling circle replication. However, this would require a very precise regulation mechanism that is not yet understood. Structure

In its natural context, the hairpin ribozyme consists of four stem regions (A, B, C, and D) connected through a four way junction, the scissile phosphate being on one of the strands of stem A (Fig. 9). Mutagenesis and modification interference studies showed that the essential nucleotides reside in stems A and B [121], and it is possible to reconstitute the catalytic activity in vitro by synthesizing both stems separately and mixing them together [13, 55, 129]. Catalysis by the hairpin ribozyme requires a rearrangement of the molecule which occurs in the presence of two molar equivalents of divalent cations: stems A and B are brought into close proximity, allowing the active site formation. Indeed, when the junction between domains A and B is blocked in a coaxial orientation preventing the folding of the molecule, the catalytic activity of the ribozyme is lost [73]. High hydrostatic and osmotic presssures experiments confirmed this Mg2+-dependent docking of the C

D

D

2+

2M

A

C B

A

B

Fig. 9. Global structure of the Hairpin ribozyme: schematic representation of the ribozyme and its docking in presence of divalent cations. Adapted from Ferré-D’Amaré, 2003 [45].

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Yan Li Li et al.

ribozyme, and helped in the calculation of the activation volume of the reaction and the number of water molecules removed from the molecule as it docks [139]. The structures of isolated stems A [16] and B [12] were first characterized by NMR spectroscopy. Both included an irregular helical central portion with non-canonical base pairs, flanked on both sides by standard base-paired helices. The fact that the hairpin ribozyme catalysis uses an associative SN2 mechanism implies that the nucleophile 2’-oxygen, the electrophile phosphate, and the leaving 5’-oxygen must be aligned for the reaction to take place. Nonetheless, NMR analysis showed that association of stems A and B is necessary to force the substrate to adopt this reactive in-line conformation: the reactive atoms are not aligned in the isolated stem A alone [16], and the conformations of the isolated stems A and B remain unchanged in presence of divalent cations. In addition, the isolated stem B NMR spectra suggested that the stem was able to adopt spontaneously a conformation different from its predominant one [12]. The hairpin ribozyme structure was further analyzed by crystallization of a ribozyme construct bound to an inhibitor of the cleavage reaction. The crystal revealed dramatic rearrangements for both stems A and B during docking, including base-pairing modifications in the noncanonical parts, as well as three-dimensional modifications. Interestingly, the substrate strand on stem A appears distorted in the docked ribozyme, and the reactive atoms present the in-line conformation required for the reaction (Fig. 10). The structure of stem B within the crystal is consistent with its NMR spectra: the minor conformation observed in NMR analysis reflects the stacking, in the docked molecule, of three purine bases that A

B

N–1

N+1 N–1

O N+1

O

O O O

O

Fig. 10. Structure of stem A in the cleavage site region: A: In the isolated stem, the reactive atoms (open spheres) are not aligned. B: In the docked ribozyme, the distortion of the substrate strand forces the reactive atoms in an in-line conformation. Adapted from Ferré-D’Amaré, 2003 [45].

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are separated by two uridines in the free stem. This base stacking results in a narrowing of the major groove, where two divalent metal ions are observed on electron density maps. Directed mutagenesis experiments showed that the ribozyme absolutely requires a guanyl residue after the scissile phosphate. The crystal structure revealed that this residue (G+1) is extruded from stem A during docking, but specifically accommodated in a pocket formed by the rearranged stem B, with which it interacts through an interhelical base pairing [12]. Interactions within the G+1 fixation pocket represent about one half of the free energy of docking [68, 69]. To visualize the hairpin ribozyme in its active conformation, a ribozyme construct without the scissile phosphate was crystallized in the presence of ammonium vanadate [45, 120]. Vanadate is selectively bound to the active sites of enzymes catalyzing phosphate transesterification. It coordinates five oxygen atoms in a trigonal bipyramidal geometry, thus mimicking the pentavalent transition state of the reaction. In the hairpin ribozyme, vanadate formed a complex with the ribose moieties of nucleosides, and established two apical coordinations to the nucleophile and leaving oxygen atoms. Catalytic mechanism

Mutagenesis and modification interference experiments showed that four purines of the active site (G8, A9, A10 and A38) have functional groups involved in catalysis. According to crystallographic analysis, the docking of stems A and B induces a distortion of the substrate that allows the in-line conformation of the reactive atoms. This distortion is comparable to that observed in a ribonuclease A-inhibitor complex. Therefore, it seems possible that the hairpin ribozyme employs, like ribonuclease A, an acid-base catalysis in which the bases G8, A38 and A10 of the ribozyme would play the same role as the histidine 12, histidine 119, and lysine 41 of the ribonuclease A, respectively [45] (Fig. 11). However, several observations suggest that the hairpin ribozyme does not function in a manner totally analogous to the ribonuclease A. Since the ribozyme catalyzes both cleavage and ligation reactions efficiently, if G8 and A38 function as base and acid, respectively, in the cleavage reaction, they must act as acid and base during ligation. However, this

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Yan Li Li et al.

A

B O

R1

A10 (1/8)

B O

R1 O

K41 (1/90,000)

–

O

P

OH

–

B

O

A38 H119 (1/400,000)

O R2

O

P

A9 (1/9)

OH

O

O

O

B

H12 (1/700,000) O

O

G8 (1/350) B

O

OH O

OH

R2

Fig. 11. Representation of essential residues involved in catalysis in ribonuclease A (A) and hairpin ribozyme (B). In the ribonuclease A active site, histidines 12 and 119 function as acid-base catalysts, whereas the lysine 41 stabilizes charges in the pentavalent transition state. Numbers in parentheses indicate the loss of catalytic activity upon deletion of the residue. Adapted from Ferré-D’Amaré, 2003 [45].

would imply a very large shift of their pKas (3-4 pH units) towards neutrality that has not been demonstrated yet. Furthermore, experiments with abasic ribozyme constructs in which the purines of the essential residues were removed showed that punctual deletions do not decrease the catalytic rates significantly compared to the effects of comparable deletions in ribonuclease A [80]. The reaction rates for cleavage and ligation by the hairpin ribozyme change less than five-fold between pH 4.9 and pH 10.1, suggesting that catalysis is independent of pH [70]. However, this does not completely rule out the possibility of an acid-base catalysis: a pH dependence of the reaction could be masked by slow rate-determining conformational changes, or if steps requiring protonation and deprotonation are both partially rate-limiting and the pKas of the corresponding functional groups are very different [103, 144]. Several methods were used to estimate the specific implication of cations in catalysis. Sulfur substitutions of non-bridging oxygens of the reactive phosphate have been particularly useful in evaluating specific interactions between these oxygens and metal cations [103, 144, 149, 155]. The “hard” Mg 2+ cation interacts strongly with hard ligands, such as oxygen, but very poorly with “soft” ligands like sulfur, while the Mn2+

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Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

cation interacts equally well with both types of ligands. Thus, a variation in cation specificity accompanying sulfur substitutions can reflect direct metal coordination to specific substituted oxygens. However, in the case of the hairpin ribozyme, the catalysis kinetics are very similar with either phosphorothioates or regular phosphates, and the reaction rates are virtually the same with either Mg2+ or Mn2+. Furthermore, experiments with [Co(NH3)6]3+ showed that it can promote the catalytic activity more efficiently than Mg2+ [144, 155], even though [Co(NH3)6]3+ interacts with RNA only through electrostatic and outer sphere hydrogen bonding, and cannot coordinate atoms directly. Catalytic activity is also maintained in presence of a wide variety of cations, including NH 4+, polyamines, and aminoglycoside antibiotics. It seems, therefore, that divalent cations stabilize the functional structure of the ribozyme through non-specific electrostatic interactions, but are not required for catalytic chemistry itself and are not involved in any direct, specific coordination of reactive atoms [144]. Varkud Satellite Ribozyme

The Varkud satellite (VS) ribozyme is harbored by the VS RNA, which is transcribed from the VS plasmid DNA and found in the mitochondria of certain Neurospora natural isolates [123, 124]. Most VS RNAs are circular monomers, but linear monomeric and multimeric forms have also been isolated. Since the VS ribozyme performs the same type of cleavage and ligation as the other ribozymes of the family, it is thought to process multimeric VS RNAs into monomeric linear and circular products. The VS ribozyme is about 150 nt in length, and constitutes the largest of the small nucleolytic ribozymes. It is the only one for which no crystal structure has yet been resolved [23, 75, 76]. Structure

The secondary structure of the ribozyme comprises six helical parts. Stem-loop I is the substrate domain and carries the cleavage site within its internal loop. The five remaining helices (II-VI) constitute the core of the ribozyme and form a H-shaped structure organized around two threeway junctions. The A730 loop, on helix VI, was shown to be essential for cleavage [4, 113] and is believed to constitute the active site of the ribozyme (Fig. 12A).

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Yan Li Li et al. A

B IV IV

I

V V III III

5¢ 3¢

II

730 VI

II I

VI

Fig. 12. Schematic representation of the cis-acting VS ribozyme. A: Secondary structure: The cleavage site is indicated by an arrowhead, and the arrow designates the tertiary kissing interaction between loops I and V. B: Three dimensional aspects of the molecule. Adapted from Collins 2002 [23].

Helical junctions were previously shown to have a great implication in the overall fold of the hairpin and hammerhead ribozymes [26, 70]. Therefore, the global structure of the VS ribozyme was thought to be essentially determined by its two junctions as well, and both junctions were analyzed by a combination of comparative gel electrophoresis, FRET (Fluorescence Resonance Energy Transfer) and mutagenesis [85]. Interestingly, these analyses showed that both junctions undergo conformational changes upon non-cooperative binding of magnesium ions, and mutations in and around the junctions affect this folding as well as catalytic activity. Further studies of the junctions docking combined with the calculation of the dihedral angle between them allowed to predict a tertiary structure of the ribozyme, in which the substrate stemloop I is located in a cleft between helices II and VI [74, 75] (Fig. 12B). A major characteristic of the Neurospora VS ribozyme is that it does not recognize its substrate in the same manner as other ribozymes of the family. For all small self-cleaving ribozymes, binding of the substrate implies the formation of extensive Watson-Crick basepairing between sequences at one or both sides of the scissile phosphate and nucleotides on the ribozyme, inducing the creation of a helix around the cleavage site. Fixation of the substrate on the VS ribozyme, on the other hand, does not involve direct base-pairing between the substrate and the ribozyme. The substrate stem-loop can be separated from the rest of the

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Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

ribozyme for a trans-acting reaction with multiple turnover; it binds tightly to the catalytic domain, mainly through tertiary interactions, with a low Km of about 0.13 mM [56]. Several interactions are required between the VS substrate and the ribozyme. The substrate stem-loop I must be in close proximity to the active site on the A-730 loop of helix VI, and it is connected to the distal end of helix II in the cis-reaction. It is also involved in a Mg 2+dependant kissing interaction with loop V. This loop-loop interaction induces conformational modifications in the substrate internal loop and places it in the right position relative to the rest of the ribozyme [2]. The substrate stem-loop I undergoes secondary structure rearrangements upon binding to the ribozyme, thus going from an inactive « unshifted » conformation to an active « shifted » one. This shift is accompanied by the unpairing of C634 (thus available for the kissing interaction with loop V), and by rearrangements within the socalled internal loop of the substrate. Both active [63] and inactive [48, 96] substrate structures have been characterized using NMR spectroscopy (Fig. 13). The inactive substrate is constituted by a stable hexanucleotide internal loop flanked on both sides by an A-type helical region. The major differences between the active and inactive conformations include rearrangements of the basepairings within this internal loop, resulting in the formation of a RNA fold composed exclusively of sheared G-A base pairs : G638 forms two shared sheared base-pairs with A622 and A621, and lies between the planes defined by these two adenines. In addition, the bases of G620 and A639 are coplanar in the active conformation and a Watson-Crick base-pairing is

Fig. 13. Structure of the internal loop of the inactive (A) and active (B) substrate of the VS ribozyme. Adapted from Hoffmann 2003 [66].

Yan Li Li et al.

21

established between C637 and G623, reducing the size of the internal loop from six to five nucleotides [63]. NMR study of the active substrate also revealed that it contains two divalent metal ion binding sites. However, only one of these sites seems essential for catalysis, since it appears only in the active form and, unlike the other site, is included in the minimal substrate [63]. Bioinformatic analysis also allowed to identify a tertiary interaction motif on the active substrate internal loop structure. This motif, involved in long-range tertiary interactions called canonical ribose zippers, was found in two rRNAs: one from helix 44 of 16S rRNA of Thermus thermophilus, and the other from helix 25 of 23S rRNA of Haloarcula marismortui. This suggests that the active substrate establishes a ribose zipper with either helix II or VI of the ribozyme [63]. Catalytic Mechanism

Mutagenesis experiments showed that most mutations affecting the catalytic activity of the Varkud satellite ribozyme did so by altering the global structure of the molecule. The only exceptions found were punctual mutations in the A730 loop on helix VI: most of those modifications led to an important loss of activity in the cis and in the trans cleavage reaction, even though none of them induced structural changes on the molecule [77, 131]. Furthermore, the cleavage site appears in close proximity to the A730 loop in low resolution structural models of the ribozyme [48], suggesting that the active site is located within this loop. Substitutions of one particular nucleotide, A756, within the A730 loop, dramatically reduce the rates of both cleavage and ligation reactions without affecting the affinity of the substrate for the ribozyme [77, 92]. Additional systematic functional group modifications analysis showed that the base of A756 is particularly important for cleavage, especially on its Watson-Crick face [92]. The loss of activity associated with substitutions or abasic site at this position could not be restored by the addition of exogenous imidazole. However, this result does not rule out the importance of the A756 base, and might simply reflect the sterical inaccessibility of the active site, since the addition of adenine did not rescue the catalytic activity either [78]. UV-light irradiation of a ribozyme containing a 4-thiouridine at a position adjacent to the cleavage site led to a cross-link with A756, showing the proximity of this

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residue to the cleavage site [62]. More recently, nucleotide analogue interference mapping (NAIM) experiments performed on the whole ribozyme demonstrated that the greatest interferences observed with the ligation reaction are associated with modifications at this position [64]. In the hypothesis of an acid-base catalysis, the nucleobase of A756 clearly appears as a strong candidate. Interestingly, McLeod et al. recently observed a pH dependence of the ligation reaction corresponding to a pKa value of 5.6 [92], and NAIM experiments [64] showed that catalysis of the ligation reaction requires the protonation of the A756 base. Alternatively, catalysis could take place through stabilization of the pentavalent transition state of the reaction. This could occur through the formation of differential hydrogen bonds with the ribozyme [86]. The transition state could also be stabilized by a charged nucleobase of the ribozyme, as well as metal ions. However, high concentrations of monovalent metal ions support the cleavage reaction efficiently [99], suggesting that, as in the case of the hairpin ribozyme, metal ions are not directly involved in catalytic chemistry, but rather just required to support the global folding of the molecule. The Hepatitis Delta Virus Ribozyme

The hepatitis delta virus (HDV), discovered in 1977, is a viroid-like satellite virus of the hepatitis B virus (HBV). Coinfection of HDV and HBV results in intensification of the disease symptoms associated with HBV [57]. Although HDV is an animal virus, it shares many features with certain plant pathogenic subviral RNAs. These features include an apparent rolling-circle mechanism for replication and the ability for both the genomic and antigenomic forms to undergo an autocatalyzed selfcleavage reaction in vitro [73]. Remarkably, the HDV ribozyme is the fastest of the known naturally occurring self-cleaving RNAs. It also stands out among ribozymes because of its stability to denaturants [44]. Structure

The HDV ribozyme is thought to form secondary structures, since it requires only 1 nucleotide at 5’ and 82 nucleotides at 3’ to the break site [109]. The sequences of both the genomic and antigenomic ribozymes can be folded into a similar secondary structure consisting of five basepaired elements, joining sequences, and hairpin loops. Four of the paired regions (P1, P2, P3, and P4) were predicted on the basis of

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Yan Li Li et al.

sequence variation between the two ribozymes [50]. The resolution of the ribozyme crystal structure revealed a fifth base-paired duplex, P1.1, unexpectedly formed by nucleotides of J1/4 and loop 3. This helix consists of only two G-C (Guanine-Cytosine) pairs, but its formation introduces a second pseudoknot nested within the P1-P2 pseudoknot [5, 22]. These two pseudoknots contribute significantly to the stability of the ribozyme folding, and any mutation or disruption to either one of them induces a marked loss of activity. Furthermore, the interactions that produce P1.1 are considered essential to constrain the ribozyme into its unique three-dimensional fold by bracing the four other helices and the J4/2 region which is functionally important (Fig. 14). 3¢

A

B 3¢

P2

J1/2

5¢ G

P3

P1 G

x n n 5¢

P1

J4/2

L3

P1.1

G G

J1.1/4

P2 U C L3 C

P3 J4/2

P4

J1/4

U1RBD P4

L4

Fig. 14. Global structure of the genomic HDV ribozyme. A: Pseudoknotted secondarystructure model of the HDV ribozyme proposed by Perrotta and Been [109]. P1 to P4 are the base-paired regions ; L3 and L4 are the corresponding loops ; J1/2, J1/4 and J4/2 are the single-stranded sequences joining helices. X denotes the nucleotide preceding the cleavage site, which can have any base. B: Three-dimensional aspects in the complex of genomic HDV ribozyme bound to U1A-RBD (U1A spliceosomal protein). Nucleotides of the U1Abinding site engineered into helix P4, and which do not participate in the ribozyme structure, are shown in outlined lower-case letters; those making close contacts with the U1A-RBD face a light grey semicircle. Adapted from Ferré-D’Amaré, 1998 [44].

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The five helical segments form two parallel co-axial stacks: P1, P1.1 and P4 are oriented co-axially to form the first stack, whereas P2 and P3 form the second one. These two helical stacks are joined side-by-side by five strand-crossovers, located at the joining region J1/4 and loop L3 [5]. P1 is the substrate-carrying duplex. It is seven base-pairs in length and includes a G•U wobble (G1•U37) at its 5’ end. Mutagenesis studies demonstrated that except for the GU•U37 wobble pair, the composition of P1 can be altered freely when the total length and the overall basepairing of the stem are maintained [151]. Similarly, P2 also lacks sequence requirements, and seems to stabilize the global structure of the ribozyme rather than forming part of the catalytic core. P3 is part of a small hairpin composed of a 3-bp stem and a hairpin loop [128]. It is contiguous on the 5’ side with P2 and on the 3’ side with P1, but has no sequence-specific contacts with the rest of the ribozyme. However, with the unpaired nucleotides U20, C24 and G25, the minor groove face of L3 forms a niche that flanks the active site. Crystallization, guided by previous biochemical work [6], demonstrated that the P4 stem could be freely altered or shortened without adversely affecting the in vitro activity of the ribozyme [128]. This indicates that P4 stabilizes the active structure without participating directly in catalysis [148]. Catalytic mechanism

The cleavage site was mapped by primer extension between C901 and G900 [148]. The structure of a self-cleaved form of a genomic HDV ribozyme was determined by X-ray crystallography, and showed that the ribozyme adopts an intricate fold that buries the active site deep within a catalytic cleft. The crystal also revealed that the C75 is projected deep into the core of the ribozyme and hydrogen bonded through its N4 with the pro-Rp oxygen of C22 that localizes in the loop of helix P3 and thus, C75 is in the proximity of the 5’-OH leaving group in the active site cleft [44, 46] (Fig. 15). Subsequent kinetic studies provided evidence that C75 acts either as a general base catalyst to activate the 2’-OH group of nucleotide -1 for nucleophilic attack [87], or as a general acid catalyst to protonate the 5’oxyanion leaving group [128]. Furthermore, C75’s pKa value is shifted toward basicity in an intermediate state of the transesterification reaction. Presumably, this change is due to the local structure around the

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Yan Li Li et al.

C75

G1 O O

O

O G22

O

O O

N

O

O

N P

O

O

U37

N O O P

G38 O

G39 P

Fig. 15. Structural components of the HDV ribozyme active site. The substrate-bearing nucleotide G1 forms a wobble pair with U37 which stacks on P1.1. The 5'-hydroxyl group is within hydrogen-bonding distance of the Watson–Crick face of C75. Adapted from Ferré D’Amaré, 1998 [44].

base, specifically the hydrogen bond between its exocyclic amino group and the negatively charged pro-Rp phosphate oxygen of C22 [110]. This local structure is stabilized by the essential short helix P1.1, and it buttresses the cleavage site G1 U37 wobble pair through stacking interactions in the tightly packed catalytic core [46]. The HDV ribozyme requires divalent cations for its activity, but in a non-specific manner: cleavage is observed even at very low (<0.1mM) concentrations of Ca2+, Mg2+, Mn2+ or Sr2+ [73, 109, 127] and in the presence of monovalent cations at very high concentrations (for example, 1M NaCl or 2.5 M NH4Cl) and at low temperature [108]. The studies of Wu indicated that the rate of the reaction is relatively insensitive to pH over a broad range [152]. However, there are more surprising findings showing that the rate of self-cleavage is stimulated by denaturants [116], suggesting that the rate-limiting step may be refolding prior to cleavage. The substrate specificity of the trans-acting antigenomic ribozyme was thoroughly investigated using a collection of substrates that varied in either the length or the nucleotide sequence of their P1 stems. It was then observed that the substrate is also important for the cleavage activity, not only for the base pairing with the ribozyme, but probably also for its contribution to the conformation change yielding a transition complex [1, 138].

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Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

The Hammerhead Ribozyme

The hammerhead ribozyme was identified as a sequence motif responsible for self-cleavage in RNAs of a number of plant pathogenic viroids and virusoids. It is the smallest of the nucleolytic ribozymes [37], and probably the most extensively studied: its structural simplicity makes it a very reliable tool, not only for structural studies [111, 125, 126], but also for therapeutic applications to target RNA viruses and oncogene messenger RNAs [117]. One remarkable characteristic of the hammerhead ribozyme is its occurrence in very divergent species. In vitro selection experiments showed that the hammerhead motif is in fact the simplest self-cleaving RNA structure, suggesting that this motif might have appeared independently several times in the course of evolution [122]. Sixteen hammerhead motifs have been identified in several plant pathogenics genomes [61], including three members of the Avsunviroidae family (ASBVd (avocado sunbloth viroid), PLMVd (peach latent mosaic viroid) and CChMVd (chrysanthemum chlorotic mottle viroid)) [102], nine viroid-like satellite RNAs of plants [10, 24, 66], and two other small circular RNAs from cherry and carnation [28, 34, 61]. Besides these plant RNAs, hammerhead structures have also been found in DNA satellite transcripts of certain families of caudate amphibians [40, 158], and in the satellite DNAs of three Schistosome species [43]. Structure

The study of the global structure of the hammerhead ribozyme shows that it contains three stems: stem I, stem II and stem III. The structure of the ribozyme was resolved by crystallography independently in the laboratories of Mckay [111] and Scott [125]. The three helical regions are variable and arranged in a ‘Y-shape’ structure. Stems II and III are coaxially aligned to form the domain 2, whereas stem I, with the C3-U4G5-A6 sequence (uridine turn) at the 5’ end of the long single-stranded section, forms the domain 1, located in the same quadrant as stem II [59] (Fig. 16). The uridine turn is identical in sequence and conformation to that found in the anti-codon loop of tRNAphe [111]. Together with the cytosine at the cleavage site, this uridine turn constitutes the catalytic pocket of the ribozyme and is absolutely necessary for catalytic activity [118]. Its importance may be understood in terms of both structure and

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Yan Li Li et al.

A

B

III

ai

m

do

III

n

domain II

I I

12 G

A

14 A

II 9A II

G U 8 7

17 C I C3 U4 G A 5 6 U-turn

Fig. 16. Schematic view of the structure of the Hammerhead ribozyme. A: Model of the crystal structure. Domain 2 represents the coaxial alignment of helices II and III, while domain 1 is the uridine turn surrounding C17 at the cleavage site. B: Secondary structure of the hammerhead ribozyme. The ribozyme comprises three helical stems, I, II and III, with a central core of unpaired nucleotides. Cleavage occurs at the position indicated by the large arrow. Adapted from Hammann, 2002 [59].

function. It carries a catalytic metal ion binding site, and also functions as a stable purine-dominated stacking platform that interacts with the reactive oxygen. Stem II, together with two absolutely conserved reversed-Hoogsteen GA base pairs (G12-A9 and A13-G8) and a singly hydrogen-bonded AU base pair (A14-U7), forms a so-called augmented stem II helix. This augmented helix stacks directly upon the helix of stem III, forming one long pseudocontinuous, distorted A-form helix. This stack forces the cleavage site C17, which joins stem III to stem I, outward to stack upon the end of stem I, in the proximity of the conserved uridine turn [125]. Stem III links stem I and stem II augmented helix through two conserved pairs (A15.1- U16.1 and C15.2-G16.2). These pairs lie adjacent to C17 and stack upon the A14-U7 non-Watson-Crick base pair of the abutting augmented stem III helix. The branching architecture of stem I and the conformation of the three-strand junction are further stabilized by a number of true longrange RNA tertiary interactions, including some hydrogen bonds. These bonds can also facilitate the interaction of the substrate with the active site, especially in the uridine turn [125].

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Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

In stem I, a hydrogen bond is formed between A15.1 and U16.1 next to the cleavage site C17, which locally broadens the minor groove. From the results of mutagenesis experiments, it has been proved that this broadening of the minor groove is crucial to catalytic activity, because of the high conservation of the A15.1-U16.1 pair [118]. Catalytic mechanism

The hypothesis that a great conformational change is necessary for catalysis was verified by the resolution of the crystal structures of the hammerhead ribozyme [100, 111, 125, 126]. It showed that, at the active site, the torsional backbone rotations are required to bring the hammerhead into a reactive conformation in which the reactive atoms are aligned [140]. Using a series of X-ray crystallographic freeze-trapping experiments and several other structural and biochemical probes, four different conformational states of the hammerhead ribozyme were captured on the cleavage reaction pathway: the initial-state structure; an “early” confomational intermediate in which a movement of the scissile phosphate group occurs in conjunction with movement of the cleavage site base and ribose; a larger or “later” conformational change, in which the 2’-oxygen nucleophile begins to align with the scissile phosphate group; and the structure of the cleaved hammerhead RNA, in the form of an enzyme-product complex [37]. As revealed by these structures, during the reaction, the nucleobase at the cleavage site (C17) is not involved in hydrogen bonding or proper stacking. The bend at the junction of stems I and III turns this residue out toward the solution to attain a near in-line attack conformation [93]. The structure folding of the ribozyme, initiated by the addition of one or several types of catalytic divalent metal ions such as Mg2+, Ca2+, or Mn2+, is now well-defined and follows a twostage process [112, 142]. In the first step, the binding of a first metal ion induces the formation of domain 2. At this stage, the ribozyme is not yet active. During the second stage, called the second transition, domain 1 is formed with the noncooperative binding of a second ion (Fig. 17). With the formation of domain 1, the final structure is, or at least considered, to be catalytically competent, for the Mg2+ concentration needed for this stage is very similar to that over which catalytic activity is acquired. It has been suggested that the metal hydroxides bind specifically to the hammerhead RNA to abstract the proton from the 2’-hydroxyl at the

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Yan Li Li et al. III

III

III

I

do

G U

A

G

2+

Mg

2+

I

C

Mg

n

I

ai

C

II A

m

A

domain II

G

A

I

U II

II

Fig. 17. A schematic illustration of the Mg2+-induced two-stage folding of the Hammerhead ribozyme. The binding of a metal ion induces a first structural transition corresponding to the formation of Domain II. Domain I is formed during the second stage, following the noncooperative binding of a second divalent metal ion. At this stage the structure is considered active and is predominant in the crystal. Adapted from Hammann, 2002 [59].

cleavage site [26, 27]. Indeed, when other counterions, such as spermine or sodium, are present to stabilize the folded structure, divalent metal ions are still required for the reaction [27]. Alternatively, these metal ions could stabilize the pentacoordinated phosphate transition state, or act by giving a proton to the leaving oxygen group [136]. The cleavage rate of the hammerhead ribozyme increases exponentially with pH over a broad pH range. Furthermore, this pH-rate profile is in correlation with the Mg2+ pKa in water, suggesting that Mg2+ is bound in close proximity to the cleavage site and could act as a base in the reaction [26]. In addition to the central conserved cores of natural hammerheads, the peripheral regions surrounding them also play a significant role in catalysis, suggesting that tertiary interactions are essential between these peripheral regions during the self-cleavage reaction [29]. Besides the Mg2+-dependent hammerhead ribozyme, a novel Zn2+catalyzed cleavage site between nucleotides C3 and U4 has been characterized in the catalytic core of the hammerhead ribozyme. This cleavage site has an unusual pH dependence, because the U4 cleavage products are only observed above pH 7.9 and reach a maximum yield at about pH 8.5 [7].

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Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

CONCLUSION

The non-coding RNAs and natural ribozymes presented here give a glimpse of the numerous functions that the RNA molecule can carry out. RNA is highly involved in the most conserved metabolic processes, and recent scientific discoveries keep on extending its functional repertoire. With the conception of in vitro selection experiments, the catalytic capacities of RNA have been greatly broadened, and the RNA world theory is more and more conceivable: it seems now plausible that RNAs were the molecular precursors of our DNA- and protein- based modern world, providing both the support of genetic information and the catalysis of metabolic processes. Small self-cleaving ribozymes already considerably helped to understand the way a RNA molecule can assemble to form functional and catalytic structures. In the future, further structural and functional studies on RNA will hopefully demonstrate that the RNA molecule can assist more complex processes, such as its own macromolecular replication. It has recently been shown that RNA might act as a cofactor in the conversion of the normal prion protein into its infectious form, and would thus play a role in the pathogenesis of prion disease [31]. This discovery illustrates how surprising and varied the RNA functions are, even though many more are probably yet to be discovered. Acknowledgments

This work was supported by CNES (Centre National d’Etudes Spatiales), CNRS (Centre National de Recherche Scientifique) and UPMC (Université Pierre-et-Marie-Curie). Yan Li Li was supported by EAKE (European Knowledge Economy Company). Sylvia Tobé, Anabelle Chabauty and Marie-Christene Maurel are from the UPMC. References [1] Ananvoranich S, Perreault JP. Substrate Specificity of d Ribozyme Cleavage. The Journal of Biological chemistry 1998; 273: 13182–13188. [2] Andersen AA, Collins RA. Rearrangement of a stable RNA secondary structure during VS ribozyme catalysis. Mol Cell 2000; 5: 469–478. [3] Avner P, Heard E. X-chromosome inactivation: counting, choice and initiation. Nature reviews Genetics 2001; 2: 59–67. [4] Beattie TL, Collins RA. Identification of functional domains in the self-cleaving Neurospora VS ribozyme using damage selection. J Mol Biol 1997; 267: 830–840.

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CH AP TE R

Plastid Origin: A Driving Force for the Evolution of Algae ALEXIS MIGUEL BELLORIN 1 and MARIANA CABRAL DE OLIVEIRA2 1

Dept. Biologia, Escuela de Ciencias, University of Oriente (UDO), Av. Universidad, Cumana, Venezuela. AP 245 2 Dept. Botanica, Inst. de Biociencias, University of São Paulo, São Paulo, Brazil

ABSTRACT Chlorophyll a is present in all organisms that release oxygen during photosynthesis. The development of this molecule altered the earth atmosphere causing a great impact in the history of the planet, redirecting the evolution of life. The oxygenic photosynthesis was developed early in the history of life by a group of prokaryotes, the cyanobacteria. This ability was later laterally transferred to the eukaryotic realm by the acquisition of photosynthetic organelles (plastids) through the engulfment and retention of formerly free living cyanobacteria by an ancient eukaryotic cell. Such kind of cellular merging, called primary endosymbiosis, probably occurred only once. Three lineages of extant algae (including the ancestors of land plants) vertically evolved from the cells which first acquired plastids. Later on, the photosynthetic apparatus was laterally transferred to other unrelated eukaryotic lineages through independent secondary endosymbiosis, i.e., the acquisition of plastids by engulfing a eukaryote, already equipped with plastids. This gave rise to the ‘second-hand’ or secondary plastid-containing algae, a diverse assemblage including disparate organisms such as euglenoids,

Address for correspondence: Mariana Cabral de Oliveira, Dept. Botânica, Inst. de Biociências, University of São Paulo (USP), R. do Matão, travessa 14 n. 321, São Paulo, SP, cep 05508900, Brazil. E-mail: [email protected]

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dinoflagellates and kelps. Thus, the history of photosynthesis acquisition by eukaryotes is extremely complex, making it very difficult to briefly define algae and to discriminate them from other organisms. Current data support the view that plastid acquisition is a rather rare event and, accordingly, a single primary endosymbiosis followed by a few very ancient secondary endosymbiosis probably account for all extant plastid diversity. The following major issues are discussed in this chapter: (1) plastid acquisition and losses by different eukaryotic lineages; (2) some recurrent themes on photosynthetic endosymbionts (genome reduction and genomic features, lateral gene transfer from plastids to nucleus and properties of protein targeting systems to plastids); and (3) the placing of the plastid-containing eukaryotes into a probably global phylogeny for eukaryotes. Key Words: Algae, protein-targeting, endosymbiosis, evolution, plastids

Abbreviations: atpB: (ATP synthase > subunit); COXII: (mitochondrial cytochrome oxidase subunit 2); EF-2: (elongation factor 2); FBA: (fructose-1,6-bisphosphate aldolase); GAPDH: (glyceraldehyde -3-phosphate dehydrogenase); LHCs: (light harvesting chlorophyll binding proteins); LSU rDNA: (large subunit ribosomal RNA gene); PCR: (Polymerase Chain Reaction); psbA: (photosystem II polypeptide D1 gene); PsbO: (oxygen-evolving enhancer 1); rbcL: (RuBisCO large subunit gene); rbcS: (RuBisCO small subunit gene); RPB1: (RNA polymerase II large subunit); rpoC1: (RNA polymerase >’ subunit gene); rRNA: (ribosomal RNA); RuBisCO: (ribulose 1,5-biphosphate carboxylase/oxygenase); SSU rDNA: (small subunit ribosomal RNA gene); tufA: (plastid-encoded elongation factor Tu gene) INTRODUCTION

The algae were traditionally conceived as the simplest or ‘lower’ photosynthetic forms of life with chlorophyll a from which evolved the ‘higher’ plants (bryophytes and tracheophytes, collectively known as land plants). This view of algae was sufficiently wide to permit that within them were included organisms so distinct as prokaryotes and eukaryotes, and within the last, some are closely related to land plants, whereas others are clearly related to non-photosynthetic protozoans. Altogether, algae were supposed to be unified on the basis that all perform oxygenic photosynthesis, although it now is clear that the photosynthetic ability is not an unequivocal marker for vertical evolutionary relatedness. The oxygenic photosynthesis was developed early in the history of life by a group of prokaryotes, the cyanobacteria. This ability was later laterally transferred to the eukaryotic realm by the acquisition of photosynthetic

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organelles (plastids) through the engulfment and retention of formerly free living cyanobacteria by an ancient eukaryotic cell. Such kind of cellular merging, called primary endosymbiosis, probably occurred only once. Three lineages of extant algae (including the ancestors of land plants) vertically evolved from the cells which first acquired plastids [174]. Later on, the photosynthetic apparatus was laterally transferred several times to other unrelated eukaryotic lineages through independent secondary endosymbiosis, i.e. the acquisition of plastids by engulfing a eukaryote, already equipped with plastids [6]. This gave rise to the ‘second-hand’ or secondary plastid-containing algae, a diverse assemblage including disparate organisms such as euglenoids, dinoflagellates and kelps. Thus, the history of photosynthesis acquisition by eukaryotes is extremely complex, making it very difficult to briefly define algae and to discriminate them from other organisms [178]. In fact, the heterogeneity of organisms traditionally designated as algae make the definition of this assemblage of organisms a complete biological nonsense in view of the new evidences about the phylogeny of these groups. This chapter does not cover the evolutionary relationships of the cyanobacteria, a well defined phototrophic clade of Gram-negative eubacteria [for some recent reviews see 94, 95, 117, 229]. The discussion will be centered on plastid-containing eukaryotes (Table 1), including the land plants (ultimately, a specific lineage of green algae fully adapted to live in terrestrial habitats [139]) and some heterotrophic protozoans that have non-functional plastids and were never considered to be algae. The most striking of such protozoans are the apicomplexans, a phylum of obligate intracellular parasites, including the causative agents of malaria and toxoplasmosis, which have a highly reduced and peculiar plastid called apicoplast [71]. Recent literature suggests that many heterotrophic protozoans lacking plastid-like organelles probably evolved from plastidcontaining ancestors [3, 241], significantly extending the scope of the algal evolution and making it indissoluble from the resolution of the global phylogeny of eukaryotes. Obviously, a central subject for evolution of photosynthetic eukaryotes is the plastid. This organelle, besides performing the photosynthesis, has other important functions in the cells, participating in the biosynthesis of fatty acids, isoprenoids, carotenoids, haeme and a number of amino acids, as well as the reduction of nitrite to ammonia, among others [152, 241]. There are different types of pigmented plastids, including the green ‘chloroplast’ of green algae and land plants (the most

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widely studied plastid), the red ‘rhodoplast’ of red algae, and the brown or golden ‘chromoplast’ of a diverse assemblage of organisms collectively known as chromophytes. Besides, there are colorless plastids as the leucoplast and amyloplast of land plants, or the very reduced plastids contained in heterotrophic algae and protozoans (as in the euglenoid Astasia or in the apicomplexans). In addition to biochemical and functional differences, plastids from different groups have variations in the ultrastructure, as the number of the membranes surrounding them, which are believed to be the result of the type of endosymbiosis from which they evolved, and the disposition and grouping of the photosynthesizing lamellae, the thylakoids. The idea that plastids are ultimately highly reduced endosymbiotic cyanobacteria is not new. It was outlined over a century ago by Schimper [195] and Mereschkowsky [157] and since 1970, gained compelling support on biochemical, ultrastructural and molecular biology grounds [for historical reviews see 80, 152, 153]. Likewise, the origin of mitochondria from an endosymbiont heterotrophic =-proteobacteria was advanced by Altmann [1] and is now sufficiently supported [88]. In both cases, as well as in second-hand plastids, the genome of endosymbionts suffered a dramatic reduction in size and gene content leading to its dependence from the host cells (Fig. 1). Most of the proteins necessary for plastids functioning are encoded in the nucleus of the host cell, what was achieved mainly through a massive lateral gene transfer from the endosymbiont genome to the nucleus [150]. The product of each plastidrelated gene is expressed in the cytosol and targeted back to plastids by adding post-translationally transit peptides at the N-terminal [35, 151, 235], which are recognized by specific translocator proteins (TICs and TOCs complexes) in the plastid membranes [119]. Current data support the view that plastid acquisition is a rather rare event and, accordingly, a single primary endosymbiosis followed by a few very ancient secondary endosymbiosis probably account for all extant plastid diversity [18, 33]. A consequence of early acquisition of plastids in the main lineages of eukaryotes is that multiple plastid losses occurred in a number of late diverging lineages, which may still contain plastid-derived genes in its nuclear genomes. Currently, the evolution of photosynthetic eukaryotes and related organisms is a field of intense research, significantly fueled by the advent of molecular methods, and the existing literature is vast, precluding any intent to summarize all the available data here. The following major

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Fig. 1. Diagrammatic views of a primary (A-C) and a secondary endosymbiosis (D-F). The lateral gene transfer from endosymbiont to host place many genes in the nuclear genome, and the products of these genes are later targeted to the endosymbiont by adding targeting sequences (transit and signal peptides). The lateral gene transfer and the loss of many genes, lead to a reduction of the endosymbiont genome.

issues are discussed in this chapter: (1) plastid acquisitions and losses by different eukaryotic lineages; (2) some recurrent themes on photosynthetic endosymbionts (genome reduction and genomic features, lateral gene transfer from plastids to nucleus and properties of protein targeting systems to plastids); and (3) the placing of the plastidcontaining eukaryotes into a probably global phylogeny for eukaryotes. Given below is a historical review of the approaches applied to ascertain the evolutionary relationships in the protists.

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ASSESSING THE ALGAL AND PROTISTS EVOLUTION

Due to their delicate nature, the paleontological record for most of algal phyla is very incomplete and difficult to interpret. Therefore, evolutionary hypothesis should be inferred by comparing extant algae. The advent of the electron microscopy and effective preparation techniques in the second half of the 20th century and, later, the development of molecular methods, revolutionized the view of life evolution and diversity [222]. Nevertheless, some noteworthy visionary insights were gained using light microscopy, as the original proposition of endosymbiosis reported by Schimper [195], Altmann [1] and Mereschkowsky [157]. Biochemical studies also revealed important information, as the photosynthetic pigments, the composition of the cell wall and other cell coverings, the nature of storage product, among others, were used to refine the accepted algal groups and to establish some relationships among them. The basic ultrastructural characters of photosynthetic eukaryotes are: (1) the number of membranes around the plastids and its relationships with other cytoplasmic membrane systems; (2) the number of thylakoids, and its arrangement (isolated, in small groups or in stacks); (3) the nature of the eye-spot or stigma, if present; (4) the presence and nature of flagellar hairs and other flagellar structures, as paraxonemal rods or flagellar swellings; (5) the structure of the transition region between the flagellar axoneme and basal bodies; (6) the arrangement of microtubular roots or other associated structures, if present, that anchor the flagella; (7) the shape of mitochondrial cristae; (8) the behavior of the nuclear envelope and mitotic spindle during mitosis and the cytokynesis patterns, etc. [87, 110, 178] (Table 1). All this information opened a window for unsuspected relationships among photosynthetic and non-photosynthetic eukaryotes, and also confirmed some former evolutionary hypothesis. For example, the evolutionary relationship between green algae and land plants was assessed on the basis of mitosis, cytokynesis and flagellated cells architecture [181]. A sister group relationship between euglenoids and kinetoplastids (forming the ‘euglenozoa’) was also proposed on the basis of shared features of cell architecture, flagella, nuclei and mitochondria [46]. Moreover, it was recognized that many algae with chlorophylls a and c generally classified as separate groups (for example the brown algae, diatoms and chrysophytes sensu lato), as well as protozoans (for example opalinids, bicosoecids, proteromonads) and

4

4

Haptophytes (Haptophyceae)

4

2

2 2

Heterokont algae (Stramenopiles)

Secondary plastids Cryptomonads (Cryptophyta)

Green algae (Chlorophyta)

Primary plastids Glaucophytes (Glaucocystophyceae) [1] Red algae (Rhodophyta)

Plastid membranes

Absent

Absent

Present

Absent

Absent Absent

Nucleomorph

chl a, chl c, Fu

chl a, chl c, Fu

chl a, chl c, PB

chl a, chl b

chl a, PB chl a, PB

Main pigments

Table 1 Contd.

Unicells with a lateral flagellar groove lined by ejectisomes, both flagella with tubular hairs, pRER around the plastids One flagella with distinctive tubular tripartite hairs, pRER; it is a highly diversified and successful group including diatoms, kelps, crysophyceans sensu lato, etc. Unicells with haptonema, an appendage involved in prey capture and locomotion, pRER, most marine bloom forming algae

‘Cyanelles’: plastids with a peptidoglycan layer Lack of flagellate stages, most are multicellular benthic marine algae with ‘pit-connections’ among cells Stellate structures at flagellar transition region, starch stored into the plastids; it is a very diverse and successful group of algae, including the ancestor of land plants

Distinctive features

Table 1. The plastid-containing eukaryotes, including ‘canonical’ algae (and land plants) as well as some protozoans with plastids (apicomplexans)

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Absent Absent

3 4

nonphotosynthetic

chl a, chl b

chl a, chl b

chl a, chl c, Pe [2]

‘Dinokaryon’ nucleus, two unequal flagella (one ribbon-like that encircles the cell in a transversal groove; the other directed posteriorly along a longitudinal groove); many form toxic algal blooms Marine reticulofilose amoebae, flagellate or coccoid cells with green secondary plastids Green flagellates with an anterior flagellar pocket, discoid mitochondrial cristae Obligate intracellular parasites with a very specialized cell invasion apparatus

chl a: chlorophyll a; chl b: chlorophyll b; chl c: chlorophyll c; Fu: fucoxanthin; PB: phycobilins; Pe: peridinin; pRER: plastidial rough endoplasmic reticulum. [1] The taxonomic classification of each algal group follows NCBI (http://www.ncbi.nlm.nih.gov/). [2] Several dinoflagellates have plastids with two or four membranes and with a pigmentation different from those indicated in this Table.

Present

4

Chlorarachneans (Chlorarachniophyceae) Euglenoids (Euglenida) Apicomplexans (Apicomplexa)

Absent

3[2]

Dinoflagellates (Dinophyceae)

Table 1 Contd.

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fungus-like organisms (e.g. oomycetes, hyphochytrids, labyrinthulids), are all closely related in having heterokont flagellate stages (cells with two dissimilar flagella, one smooth and the other carrying tubular tripartite flagellar hairs). This large and very diverse assemblage of protists, presumably monophyletic, was accordingly dubbed as Heterokonta [30] or Stramenopiles [177], and was established as one of the main lineages of eukaryotes. Concurrent with the development of the ultra-microscopy was the development of the molecular biology techniques, which allowed the direct manipulation and analysis of nucleic acids (DNA and RNA) and proteins. The hallmarks to study phylogenetic relationships were the development of the DNA sequencing method in the 1970s [193] and the PCR (Polymerase Chain Reaction) in the 1980s [191]. These techniques opened the way for the use of molecular markers to construct phylogenies [8]. Much progress was achieved by comparing homologous nucleotide sequences among different organisms. Initial studies were based on comparisons of a single molecule, i.e. the small subunit ribosomal RNA gene (SSU rDNA [see 182 for a review]), which established a ‘standard model’ to study global evolution of prokaryotes [245] and eukaryotes [203] for nearly ten years. In this model, the eukaryotic evolution was outlined as a basal ‘stem’ of deep-branching separated lineages of protists, followed by the radiation of the ‘crown group’, including fungi, animals and plants together with their protistan allies, as well as several separated lineages purely composed of protists, as the heterokonts or the group called ‘alveolates’, including ciliates, apicomplexans and dinoflagellates. Several amitochondriate protists (diplomonads, microsporidia and parabasalids) were recovered as the first diverging eukaryotes, supporting the ‘archezoa’ hypothesis [29], which claims that these and other amitochondriate microbes evolved prior to the acquisition of mitochondria by endosymbiosis. However, the deep-branching position of most of these organisms in SSU rDNA trees was shown as artifactual when other sources of information were incorporated, especially protein sequences [for example, 23, 67, 127, 189]. This and other discrepancies stimulated a period of significant revisionism, leading to the establishment of new models of eukaryotic evolution based on: larger data sets including broader taxon sampling and/or concatenated multigene analyses [11, 12, 133, 161, 166, 248]; some rare and putatively irreversible genomics events, as ancient gene duplications, gene replacements or gene fusions that can be used as markers [70, 103, 176,

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204]; comparative genomics [47, 149]; as well as on the synthesis of many different trees into a unique and plausible scenario [202]. These new phylogenies present most of the phylogenetic groups supported by the ultrastructure and SSU rDNA sequences, as the heterokonts, alveolates, euglenozoa, opisthokonts (fungi, animals and related protists), viridiplantae (green algae plus land plants). Nevertheless, the emerging picture [for example 10, 34] is striking in revealing a major evolutionary diversification of eukaryotes: the nearly simultaneous radiation of five to eight ‘supergroups’ from an unknown common ancestor (Fig. 2). These supergroups are primarily composed by unicellular protists and do not have formal names or rank. Obviously, some of them are better supported than others and many important issues are still not settled, in fact, conflicting views are often proposed [for example, 167], but it is currently accepted that eukaryotic early evolution was a rather explosive radiation [47]. An overall probable history of plastid acquisitions and transfers is also depicted in Fig. 2, showing that plastid spreading through endosymbiosis was very effective, in the sense that most of the major eukaryotic supergroups contain at least one member with plastids. PRIMARY ENDOSYMBIOSIS: THE BIRTH OF PLASTIDS

Chlorophyll a is present in all organisms that release oxygen during photosynthesis. The development of this molecule altered the earth atmosphere causing a great impact in the history of the planet, redirecting the evolution of life. Chlorophyll a molecule and its associated chemical and photochemical systems are extremely complex to have been developed independently more than once. It is a wellestablished fact that plastids bound by two membranes are the vertical descendant of cyanobacteria, which became fully adapted to live into eukaryotic host cells through primary endosymbiosis (Fig. 1 A-C). The two membranes of such plastids, called primary (or simple) plastids, have been explained in two different ways [80]. The most common is that the inner membrane was the plasmalemma of the endosymbiont, while the outer membrane represents the phagosome (food vacuole) of the host. The other explanation is that the inner and outer membranes are the original cyanobacterial Gram-negative envelope, and assumes that the phagosomal membrane has been lost. This is supported by the peculiar carotenoid composition of the outer plastid membrane, which is similar to that of the cyanobacterial outer membrane [111] and, recently, by the

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Fig. 2. An unrooted hypothetical global phylogeny of eukaryotes based on the synthesis of many molecular data [based on Refs. 10, 34, 125]. A plausible history of plastids (in colored lines and arrows) is also illustrated. Dotted lines represent putative relationships, but weakly supported. The groups that contain plastid are in bold typeface. CB indicates the first gain of plastids by one ancient plantae, which enslaved a cyanobacteria through a primary endosymbiosis. The red arrow indicates a secondary endosymbiosis implicating a red alga, which occurred basally in the supergroup called chromalveolates. The green arrows indicate two independent secondary endosymbiosis implicating green algae, transferring green plastid to a cercozoan lineage (chlorarachneans) and to a discicristate lineage (euglenids).

discovery of some cyanobacterial proteins that are putatively homologues to translocator proteins located on the outer (Toc75 and Toc34) and inner (Tic55) plastid membranes [21, 186]. There are evidences that the cyanobacterial homologue of Toc75 may act as a voltage-gated channel,

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which is related to secretory proteins of cyanobacteria [104, 186], bolstering the hypothesis that the protein import machinery of plastid could be derived from the secretory system of these prokaryotes. Once the endosymbiosis occurs, there is a close contact between different genomes, opening the door for lateral gene transfer. The mechanism by which this transference occurs is not established, but transposable elements could be involved. Lateral gene transfer, as well as gene loss and replacement are probably random events, but there seems to be some kind of directionality once the host nucleus has a tendency to acquire genes from the endosymbionts, while the genome of those have a tendency of losing redundant genes [20]. One possible explanation for that apparent opposite tendency of host and endosymbiont genomes is that the nucleus would be a more geneticallystable ‘environment’ and, therefore, less prone to gene loss. Interestingly, this process of transference of plastid genes leads to a partial dependence of the endosymbiont to the host, and is probably essential for the maintenance of a stable endosymbiotic association. The plastid genomes are greatly reduced (~34 to 200 thousand base pairs) if compared to extant cyanobacteria (~1.7 to 9 million base pairs) [http:// www.ncbi.nlm.nih.gov/]. The number and composition of genes will vary from lineage to lineage, as well as the presence and abundance of introns, repeats of rRNA genes and other characteristics [99]. Up to now, 40 plastid genomes have been fully sequenced, of those 29 are from the viridiplantae (green algae plus land plants [http:// www.ncbi.nlm.nih.gov/]). The primary plastids are contained in three well-defined lineages of extant eukaryotes: the glaucophytes, the red algae and the green algae (including land plants) (Table 1), which probably descend from the first eukaryote that performed photosynthesis. All the other photosynthetic eukaryotes acquired plastids by engulfing not cyanobacteria, but one of those primary photosynthetic eukaryotes (Fig. 3). The plastids of glaucophytes or glaucocystophytes, a small group of freshwater unicellular algae, provide compelling evidence for their cyanobacterial origin. These plastids, termed traditionally ‘cyanelles’, have retained a thin layer of peptidoglycan (the main component of the cyanobacterial cell envelope) between the plastid membranes, and a large central body resembling a cyanobacterial carboxisome [143]. In addition, these plastids, like those of the red algae, have cyanobacterial-like pigmentation (chlorophyll a plus phycobilins) and ultrastructure (unstacked equidistant thylakoids

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Fig. 3. Schematic representation of plastid evolution through primary and secondary endosymbiosis. chl a: chlorophyll a, chl b: chlorophyll b, chl c: chlorophyll c, Fu: fucoxanthin, PB: phycobilins, Pe: peridinin, pRER: plastidial rough endoplasmic reticulum. Plastid losses that occurred in other lineages than ciliates are not indicated.

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with attached granules of phycobiliproteins called phycobilisomes). In both groups, glaucophytes and red alga, the plastid genomes, though significantly reduced if compared with free-living cyanobacteria [62, 121, 122, 163, 164], contain the largest known gene repertoire for these organelles, including many fundamental genes for photosynthesis, as well as genes necessary for ‘housekeeping’ and other more specialized tasks [84, 169, 185, 212, see Table 1 in Ref. 99]. The green algae, a very diverse assemblage of photosynthetic eukaryotes ranging from unicellular planktonic forms to macroscopic seaweeds, have plastids that differ somewhat from the cyanobacteria (for example, lacking phycobilins, containing instead, chlorophylls a and b, and having stacked thylakoids). Nevertheless, these plastids have retained cyanobacterial ancestral features, as a gene involved in the growth of the peptidoglycan layer during septum formation [228] and many other features that suggest, without doubt, that these plastids also evolved from cyanobacteria [175]. Since 1980, the primary endosymbiotic origin of plastids rarely has been doubted. The main questions have been: how many times did such endosymbiosis occur and who were the partners? [reviewed in Refs. 51, 173, 174, 175]. While the presence of different photosynthetic pigments suggests separate origins for green algae plastids and glaucophytes/red algae plastids, involving independent endosymbiosis with differently pigmented prokaryotes (a polyphyletic scenario) [184], some conserved complex features of plastids, thought to be ‘mutationally onerous’, as the protein targeting and importing machinery, suggest a single origin. It is more parsimonious to think that the basics of this machinery evolved only once in a putative common ancestor and were inherited by the green algae, the glaucophytes and the red algae plastids (a monophyletic scenario), rather than by independent origins followed by convergent evolution [28] (Fig. 3). Despite the considerable debate among these conflicting hypotheses, the bulk of molecular data, especially from plastid genomes, favors the monophyletic origin of plastids. Almost all individual plastid sequences compared, such as plastid encoded SSU rDNA [for example 16, 229] and other four plastid genes (tufA, atpB, rpoC1 and psbA [175]), as well as light-harvesting antenna proteins [64, 246], consistently indicate that the primary plastids, mainly the well sampled red and green algae plastids form a single robust clade within the cyanobacteria. The only plastid genes that depart from this pattern are the ribulose 1, 5-biphosphate carboxylase/oxygenase (RuBisCO) genes rbcL and rbcS, which are misleading in this regard, since those were the

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subject of multiple ancient gene duplications and lateral gene transfers [53]. Unfortunately, the published concatenated multigene analyses of plastids [149, 150, 228] include only one representative of cyanobacteria and do not, therefore, indicate the number of times that cyanobacteria were ‘enslaved’ by eukaryotes. Other plastid traits, as the rRNAencoding inverted repeats and two distinctive gene clusters (namely psbB/N/H and atp/rps/rpo) shared among most primary plastids, but absent in cyanobacteria, are often interpreted as derived postendosymbiotic innovations, suggesting a single plastid origin [175]. Additionally, the protein import machinery, which has the same general mechanism in these plastids (by post-translationally adding a transit peptide at the N-terminal to each plastid-targeted protein [151]), also strongly suggests a monophyletic origin of plastids, as transit peptides of glaucophytes, red and green algae are interchangeable among these plastids with no significant loss of function [206]. On the other hand, a common genomic evidence assumed to favor plastid monophyly, the similarities in gene content among primary plastids, was recently shown as most consistent with a convergent evolution scenario and not as a direct support for single origin of plastids [210]. The primary endosymbiosis is a very ancient event (late paleoproterozoic, ~ 1,600 MYA [249]) which makes it extremely difficult to recover its evolutionary history. Overall, plastid data strongly favor the single origin of plastid, although the alternative and almost untestable hypothesis of multiple origins involving closely related cyanobacteria cannot yet be fully discarded. The monophyletic origin of primary plastids implies that host cells containing such plastids are vertically related as shown by comparisons of nuclear or mitochondrial genes. If primary plastid-containing eukaryotes form a well supported clade to the exclusion of all other eukaryotes in nuclear or mitochondrial gene phylogenies, the most plausible scenario is that a putative common ancestor that first acquired plastid gave rise to the extant glaucophytes, red and green algae (and perhaps some unknown extinct lineages). This rather holophyletic clade is consistent with the proposal of revised Kingdom Plantae, including only these primary photosynthetic eukaryotes [27] and named the ‘Plantae’ hypothesis. Contrarily, if there is no congruence between plastid and nuclear/mitochondrial genes, it is still possible that primary plastids were acquired only once (as is widely supported by plastid genes). If this is the case, from the cell that first acquired plastids, evolved not only the

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glaucophytes, the red and the green algae, but also other extant eukaryotes that secondarily lost their plastids. In such a scenario, it is possible that the closest relatives of one of the primary plastid-containing group might be plastid-lacking eukaryotes instead of the plastidcontaining ones. Mitochondrial individual genes [for example, 22] as well as multiple concatenated mitochondrial genes [24, 171] strongly favor the first scenario, suggesting that at least red and green algae are sister groups to the exclusion of other eukaryotes. The split of red and green algae was calculated to have occurred about 1,500 MYA [249]. The statistical supports for the red and green algae clade are usually high, although these results have been disputed on the basis that other mitochondrial genes, not included in such analyses, favored the non sisterhood of red and green algae [211]. The nuclear genes, on the other hand, have yet not provided an unequivocal answer. Most of individual nuclear genes compared are uninformative [57]. Weak to moderate support for the monophyly of the primary-plastid lineages has been obtained only from the actin gene [17, 207] or some SSU rRNA phylogenies based on corrected models for sequence evolution [232]. A further strong support, at least for red and green algae sisterhood [183], was found in the nucleus-encoded protein EF-2 [161]. Most authors welcomed it as an important piece of evidence towards the general acceptance of a monophyletic origin of primary plastid-containing eukaryotes. The subject is not resolved yet, as it was argued that the phylogenetic signal of this gene is highly compartmentalized and only half of the EF-2 sequence favors the red and green algae sisterhood [211]. However, the concatenated nuclear genes analyses favored the Plantae hypothesis, first with weak statistical supports [11, 161], but recently with a strong support based on an impressive larger data set (more than 100 nuclear genes) [12]. All these results seem to show that there is a tenuous, though meaningful, phylogenetic signal for the relatedness of primary plastid-containing eukaryotes in the nuclear genome. The only nuclear sequence that appears to tell a different history, and that has in fact fueled most of debates about primary plastid origins, is the RNA polymerase II large subunit (RPB1). This, in some analyses, regarded the red algae as a deeply separate lineage from the green algae [208, 209]. Other studies of this protein [48, 144, 161], however, did not support this separation. Likewise, a recent multigene study has challenged the view that primary photosynthetic eukaryotes are monophyletic [167]. This study was largely based on previous multigene datasets [11, 161]

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updated with new sequences and excluding fast-evolving sequences (for example, those from amitochondriate protists and cryptomonads’ nucleomorph) as well as bacterial outgroups. The red algae and glaucophytes were retrieved as basal separate lineages, whereas the green algae were late diverging and related to euglenozoa and some amoebae. The statistical supports for these results, however, were rather few. When the EF-2 sequence was considered, the sisterhood of red and green algae was recovered with a higher statistical support. In conclusion, if all the available evidences favoring primary plastid monophyly are compared to evidences unequivocally refuting it, the most plausible scenario is the single origin of plastid, although more data are needed to establish it as a fact. Another important issue on the origin of primary plastids is who were the two partners of such primary endosymbiosis? There is no specific candidate at the cyanobacterial side, mainly because the evolution of cyanobacteria is still poorly understood. The most well sampled gene among cyanobacteria (SSU rDNA) supports the view that these organisms suffered an explosive radiation after the splitting of the structurally simplest cyanobacteria [229], the Gloeobacter group. In this group, which lacks thylakoids, photosynthesis takes place in the plasmalemma, as in other photosynthetic bacteria [187]. The recent published complete sequence of Gloeobacter genome [164] supports this separation. Based on SSU rDNA sequences, all the other extant cyanobacteria, including plastids, fall into at least six main lineages that radiate nearly simultaneously from a single node and there is no specific group consistently related to the plastid lineage [229]. Other SSU rDNA comparisons retrieve the plastids as a sister group of the whole cyanobacteria radiation [for example, 107], or as a basal lineage, but none of the published phylogenetic analyses has revealed the extant cyanobacteria lineages that are more related to plastids. On the other hand, plastid relationships with extant cyanobacteria were traditionally inferred by comparing biochemical features, especially pigment composition, but this approach has also significant uncertainties. This approach was initially bolstered with the discovery of the ‘prochlorophytes’ (oxygenic photosynthetic prokaryotes having chlorophylls a and b but no phycobilins) [25, 42, 138], which were supposed to be the link between green plastids and prokaryotes. Further phylogenetic studies, however, rejected this view; the three described species of prochlorophytes correspond to separate lineages within the

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cyanobacteria, and none of them are related to the plastid clade [172, 230]. Moreover, though prochlorophytes have chlorophylls a and b as main pigments, their core proteins of light harvesting antenna systems (called PCBs or Prochlorophyte Chlorophyll-Binding proteins) [135] are not related to eukaryotic Light Harvesting Chlorophyll-Binding proteins (LHCs), which are the core membrane plastid proteins to which the chlorophyll molecules (only chlorophyll a in ‘red plastids’, chlorophylls a/b in ‘green plastids’ and chlorophyll a/c in ‘brown plastids’) are coupled [91]. These results, together with the discovery of phycobilin genes in prochlorophytes [106, 179, 224] as well as the likely ancient presence of chlorophyll b in cyanobacteria [227], suggest that cyanobacteria were probably ancestrally more diverse in their light harvesting antenna complexes, probably having phycobilisomes with cyanobacterial types of chlorophyll a binding proteins (not LHCs, which are a plastid invention) and probably chlorophyll b [89]. In this scenario, it is likely that the pigments were differentially lost in the plastids: in glaucophytes and red algae the chlorophyll b of ancestral cyanobacteria was secondarily lost, whereas in green algae the phycobilisomes were lost (Fig. 3). An alternative scenario is the independent acquisition of chlorophyll b more than once in the evolution of those lineages [172, 230]. Chlorophyll b differs from chlorophyll a by presenting a formyl group (CHO) in place of a methyl group (CH3) on one of the pyrroles. This conversion reaction from chlorophyll a to b uses molecular oxygen and is catalyzed by chlorophyllide a oxygenase (CAO) [66,219]. At the host side, if the Plantae are holophyletic, the features of a putative common ancestor can be inferred by comparing extant glaucophytes, red algae and green algae [34]. As such, it is likely that the Plantae ancestor was a biflagellate anterokont with a cruciate system flagellar rootlets and probably associated multilayered structures, flattened mitochondrial cristae, =-1,4 linked glucans as main storage products, among other general features [34]. From this putative ancestor that acquired plastid, probably first evolved the glaucophytes, as the cyanobacterial-like cyanelles and most of multigene studies suggest, followed by the sister lineage that gave rise to the red and green algae. The red algae lost the flagella secondarily and evolved mainly as a multicellular group of marine benthic phototrophs [see 194 for a review]; on the other hand, many green algae maintained the unicellular or colonial flagellate way of life, although others evolved independently different degrees of multicellularity: filamentous, parenchymatous,

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coenocytic, etc. but, whatever the case, almost always producing flagellate stages for reproduction (zoospores and gametes) [see review in 139]. Early in its history, the green algae split into two main groups: the Chlorophyta, most of them with cruciate flagellar roots, and the Streptophyta, with a unique large flagellar root disposed laterally and open mitosis. This last group includes the land plants and its closest green algal relatives, the charophytes [86, 123]. SECONDARY ENDOSYMBIOSIS: SPREADING THE PLASTIDS

The gain of photosynthesis was not limited to the primary plastidcontaining eukaryotes. Other extant eukaryotes perform photosynthesis, using plastids ultimately acquired laterally from other plastid-containing eukaryotes, which were engulfed and retained by secondary hosts. Such secondary symbiogenesis accounts for plastids of a bewildering diversity of photosynthetic eukaryotes, most of them unicellular, as the euglenoids, the heterokont algae, the dinoflagellates, the haptophytes, cryptomonads, dinoflagellates and chlorarachneans (Fig. 3). Moreover, there are nonphotosynthetic second-hand plastids in the parasites of the phylum Apicomplexa, a sister group of the dinoflagellates (Table 1). The first clue suggesting that all these plastids have eukaryotic origin was the presence of more than two membranes surrounding them [220], i.e. three membranes in euglenoids and most of dinoflagellates, and four membranes in heterokont algae, haptophytes, cryptomonads, apicomplexans and chlorarachneans. To explain the origin of such additional membranes different processes were invoked [28, 78, 79, 137, 196, 240], but the most likely explanation is that all secondary plastids were formed ultimately by the same general way: the phagotrophic ingestion and further retention and reduction of an entire eukaryotic unicellular algae into a secondary host cell, leading to the establishment of a very reduced cell within another cell [78, 79] (Figure 1D-F). This is more apparent in cryptomonads and chlorarachneans, where the endosymbiont reduction is not fully accomplished and the plastids still keep several eukaryotic cellular features. The uptake of a complete eukaryotic cell by another cell implies that secondary plastids first arose bound by four membranes; the two innermost membranes corresponding to the primary plastid envelope; the third membrane corresponding to the former plasma membrane of the primary host, now called periplastid membrane; and the fourth outermost membrane probably corresponding

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to a phagosomal membrane produced by the secondary host (as is likely in chlorarachneans and apicomplexans) or to a rough endoplasmic reticulum membrane of the secondary host (in cryptomonads, heterokont algae and haptophytes) [32, 35]. The three membrane plastids of euglenoids and most dinoflagellates could have evolved later through the loss of one membrane, probably the periplastid membrane, from a previous four-membrane plastid [32, 35, 54]. The uptake of eukaryotic endosymbionts implies the acquisition of three distinct genomes by a host cell (the endosymbiont’s nucleus, mitochondria and plastid genomes), which were dramatically reduced. This is especially true in the case of mitochondria, which were completely lost in all secondary plastids, and the endosymbiont nucleus, which was completely lost in most of the secondary plastids, but still retained, though highly reduced, in plastids of cryptomonads and chlorarachneans. In other words, a massive gene loss and transfer from the nucleus and plastid of endosymbiont to the secondary host nucleus characterized the secondary symbiogenesis. This also implies the evolution of effective protein trafficking machinery through the three or four membranes surrounding such plastids, and the addition of specific targeting sequences to all transferred proteins [39]. All that seemed to have been achieved by utilizing the cell secretory and plastid importing systems in a two-step process that still is poorly understood in most of the secondary plastid-containing eukaryotes: first, the nucleus-encoded, plastid-targeted proteins are placed in the lumen of the endomembrane system using standard signal peptides added at the N-terminal; then the proteins are imported through the plastid membranes by Toc and Tic translocons, which recognize the standard transit peptides also added at the N-terminal of presequences [4, 50, 113, 114, 129, 131, 134, 165, 198, 214, 236, 237]. Thus, in plastids bound by four membranes, the former plasma membrane would have to acquire some kind of protein trafficking machinery [35], though this is still not well characterized in most of secondary plastids. The eukaryotes that contain secondary plastids are so diverse, and the secondary plastids are so distinct, that the separate origins of such plastids have rarely been doubted. Currently, it is well established that there are two main kinds of secondary plastids: (1) those derived from green algae, which contain chlorophylls a and b as main pigments, and are present in euglenoids and chlorarachneans; and (2) those derived from red algae, some of which (the cryptomonad plastids) have retained

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the typical red algal pigmentation (chlorophyll a and phycobilins) plus one postendosymbiotic and purely eukaryotic pigment, the chlorophyll c, whereas others (heterokont algae, haptophytes and dinoflagellate plastids) contain chlorophylls a and c together with carotenoids such as fucoxanthin or peridinin, but have lost the phycobilins (Table 1, Fig. 3). Altogether, the plastids with chlorophylls a and c make up the ‘chromophyte’ plastids. The origin of unpigmented apicomplexan plastids was largely controversial, but the most accepted hypothesis is that such non-photosynthetic plastids also evolved from red algae [174, 241]. Although we are almost certain about the eukaryotic ancestors of all known secondary plastids (green or red algae), we do not have confidence in the number of times that these algae were enslaved and converted into plastids. It has been argued [31, 32, 35] that such a complex process as secondary symbiogenesis is too ‘evolutionarily onerous’, requiring first a massive and successful gene transfer from endosymbiont to host nucleus, and second the evolution of an effective protein trafficking system across three or four distinct membranes; accordingly, such a process may not be as frequent as it has been often claimed (for example, seven separate endosymbiosis in the separate ancestors of each algal group containing secondary plastids, plus apicomplexans [175]). The most parsimonious view suggests only two secondary endosymbiosis, one implicating a green alga and other implicating a red alga [32]. However, the most supported current scenario is that three separate events account for all secondary plastids diversity: two separate endosymbiosis implicating distinct green algae (one leading to the plastids of euglenoids, and another leading to plastids of chlorarachneans) and an independent endosymbiosis implicating a red algae (giving rise to plastids of cryptomonads, heterokonts, haptophytes, dinoflagellates and apicomplexans) [6, 18, 125, 241]. This last event was calculated to have occurred about 1,300 MYA [249]. The Green Secondary Plastids

The chlorarachneans or chlorarachniophytes are perhaps the best characterized case of a secondary endosymbiosis. These organisms are chimaeras of reticulofilose amoebae that belong to the eukaryotic lineage called Cercozoa [7, 15, 36, 37, 108, 124, 126] with unicellular green algae functioning as plastids [81, 116, 155, 234]. There are a few recognized species of chlorarachneans, most of them are benthic marine phagotrophic amoebae that can form naked flagellate dispersal stages or

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even walled unicellular non-flagellated coccoid stages. There are also purely flagellate planktonic species, as the most studied chlorarachnean, Bigelowiella natans Moestrup [158]. The endosymbiont green alga has lost most of its cell components, although it still keeps a vestigial cytoplasm with functional ribosomes (called periplastid space), a reduced nucleus (the nucleomorph) and, of course, its plastid. In total, the plastid stroma is separated from the secondary host cytosol by four distinct smooth membranes, none connected to the endoplasmic reticulum or to the nuclear envelope. The nucleomorph of chlorarachneans has a much reduced, compacted genome (380-455 kb) contained in three chromosomes [81, 82, 154]. This genome contains rRNA operons and a minimal set of genes for proteins implicated in expression and maintenance of the nucleomorph genetic system (housekeeping) as well as other tasks, including some plastid proteins with putative transit peptides; however, the bulk of plastid proteins were retransferred from the nucleomorph to the secondary host nucleus. Remarkably, all the genes for enzymes involved in the primary metabolism were lost in the nucleomorph. On the other hand, there are many introns but they are much reduced and in fact correspond to the smallest known spliceosomal introns [81]. Although very divergent and AT biased, the SSU rRNA and other nucleomorph genes have widely confirmed that this reduced nucleus belongs to the green algae clade [81, 155, 234], probably an ulvophycean green alga [116]. For the euglenoids, there are convincing data suggesting that the three membrane-bound green plastid is also a reduced green alga [16, 229]. The euglenoids, together with kinetoplastids and diplonemids, belong to the euglenozoa [178, 200], which in turn probably belongs to a very diverse supergroup of protists called excavates [34, 201]. These peculiar unicellular flagellate algae have an anterior pocket into which are inserted the flagella and a characteristic rodlike structure that lies parallel to the flagellar axoneme. This group also presents a nucleus with permanently-condensed chromosomes, with a special type of closed mitosis, and a complex proteinaceous, generally flexible, pellicle (or periplast) beneath the plasma membrane, among other interesting features. Most of euglenoids are mixotrophs, having plastids and performing photosynthesis together with osmotrophy, while others have non-photosynthetic plastids and are purely osmotrophic, and still others lack plastids altogether and are phagotrophic. Although it was initially proposed that euglenoid plastids originated by a primary endosymbiosis [28], the green algal secondary origin for such plastids

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[78] has been widely supported both by individual plastid gene sequences [16, 53, 146, 175] as well as multiple concatenated plastid genes [149, 150, 229]. The ancient green alga endosymbiont was probably a prasinophycean (scaly green flagellates, most of them marine) [146]. This endosymbiont was almost completely reduced, only its plastid remains enclosed by a phagosomal membrane produced by the host. An intriguing feature of the plastid genome of Euglena is that it is characterized by the proliferation of numerous introns: both group II introns and very small group III introns, as well as twintrons (introns-within-introns) [100]. There are also three tandem-like organized rRNA repeats instead of two, as in most plastids [100]. The plastid genome of the colorless euglenoid Astasia recently has been sequenced [85]; this very small genome (73 kb) is roughly similar to that of Euglena, except that it lacks nearly all photosynthetic genes. It is not clear when the plastids were acquired by euglenoids. Most of nucleus-encoded SSU rDNA phylogenies show that plastids-lacking phagotrophic euglenoids are the basal lineages and, accordingly, support the view that plastids were acquired later in this group by a phagotroph that gave rise to phototroph lineages. Thereafter, from some of these phototrophs evolved secondary heterotrophic colourless species [142, 147, 159, 168]. Comparisons of the complete plastid-encoded rRNA operon also supported this view [146]. However, a recent study based on a broader sample of phagotrophic species has shown that they are not necessarily basally diverging [26]. On the other hand, the surprising discovery of putative plastid-derived genes in the parasitic kinetoplastids [101, 199] bolstered the view of a more ancient plastid origin in the euglenoids, suggesting as a most parsimonious scenario that both kinetoplastids and euglenoids evolved from a common ancestor with plastids, which were secondarily lost in the kinetoplastids as well as in the heterotrophic euglenoids. Many of these plastid-like genes in kinetoplastids have been disputed as methodological artifacts [190], but a striking organelle unique of kinetoplastids seems to support the idea that this group ancestrally contained plastids. This organelle, known as glycosome, is not a relic of a plastid, but a kind of peroxisome that has sequestered most of the plastid-like enzymes of the putative former endosymbiont, including those of the glycolitic pathway, which are then topologically separated from cytoplasm [101]. Considering all the available data discussed above, especially the putative ulvophycean nature of the chlorarachniophyte plastids and the

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prasinophycean nature of euglenoid plastid, the ‘cabozoan’ hypothesis [32] suggesting a single secondary endosymbiosis origin for chlorarachneans and euglenoid plastids, is not supported. Likewise, there are no molecular data supporting that cercozoans and discicristates (the lineages to which chlorarachneans and euglenoids respectively belong) are related, although this should not be definitively discarded as the understanding of eukaryotic mega-evolution is yet not complete. The ‘Red’ Secondary Plastids

A red algal origin was first proposed for plastids of cryptomonads [79, 240], which are the unique plastids that contain phycobilins, bound by more than two membranes. These plastids also contain chlorophyll c, a pigment found in plastids of the heterokont algae, haptophytes and dinoflagellates (Table 1). On this ground, in 1962 it was proposed that all these algal groups are related and should be classified together in a single phylum, the Chromophyta [43, 44], a revolutionary idea that later was refined by proposing a specific kingdom (Chromista) including cryptomonads, heterokont algae, haptophytes and non-photosynthetic heterokonts, all of which have very similar plastid ultrastructure, but excluding dinoflagellates [30]. Since a red algal origin of cryptomonad plastids became widely adopted in the light of convincing ultrastructural and biochemical evidences, a red algal origin was also extended to the other chromists, as well as dinoflagellate plastids [79, 240], a hypothesis that has been further supported by molecular data [59]. The origin of the plastids of Apicomplexans, on the other hand, has been more difficult to access and is still debated, although most evidences favor the red algal origin. The cryptomonads are unicellular marine or freshwater flagellates with dorsiventrally constructed cells that have a distinctive ventral flagellar pocket lined by ejectisomes [178]. The secondary plastids of cryptomonads, like those of chlorarachneans, are cells within cells: they have a vestigial cytoplasm (the periplastid space) with ribosomes, and even stored granules of the main food reserve (starch). In the periplastid space are also located the endosymbiont reduced nucleus (the nucleomorph) and its plastid, which contains peculiar broad thylakoids, disposed in pairs, containing phycobilins in its lumen, not aggregated into phycobilisomes. There are four membranes separating the plastid stroma and host cytosol, but contrarily to chlorarachneans, the outermost

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membrane is continuous with the rough endoplasmic reticulum and with the outer membrane of the nuclear envelope. This membrane is called here as plastidial rough endoplasmic reticulum (pRER). Thus, the eukaryotic endosymbiont lies into the lumen of the host endoplasmic reticulum, and not into a phagosomal vesicle. The nucleomorph was first described in the cryptomonad plastids [93] and later it was verified that this compartment contained DNA [145]. The complete sequencing of the nucleomorph genome of the cryptomonad Guillardia theta D.R.A.Hill and R.Waterbe [61] showed it to be a very reduced (ca. 551 kb with a total of 511 identified genes) eukaryotic genome contained in three linear chromosomes. The nucleomorph genes are very divergent and AT biased, but contrarily to chlorarachnean nucleomorph, they contain very few introns [61]. Phylogenies of many of these genes show that the nucleomorph belongs to the red algae [5, 16, 38, 48, 58, 61, 109, 207, 231], representing a monophyletic early diverging lineage radiating in many trees before the Bangiophycidae [162, 170]. On the other hand, the sequencing of the plastid genome of cryptomonad endosymbiont [60] also confirms the red algal origin of cryptomonad endosymbiont, as this genome contains basically the same gene content and arrangement as found in the red algae Porphyra [185] and Gracilaria [99]. The heterokont algae and the haptophytes have almost identical four-membrane plastids, which are also located in the lumen of the endoplasmic reticulum, but lack nucleomorphs. Besides chlorophylls a and c, these plastids have the brown carotenoid fucoxanthin or fucoxanthin-derived pigments, and lack phycobilins. The periplastid space is almost completely reduced, lacking ribosomes and the food reserve (a soluble >-1,3 glucan) is stored in the host cytoplasm. An intriguing feature of such plastids, shared with cryptomonads, which may be implicated in protein trafficking [35], is a system of small vesicles located in the periplastid space between the nucleus and plastid envelope. The heterokont algae constitute the most diverse algal group, ranging from tiny picoplanktonic unicells to the large and complex multicellular kelps, including all the levels of organization present in algae and thousands of described species [177]. There are many classes of heterokont algae, the most familiar being the brown algae, the diatoms, the chrysophytes and the xanthophytes. On the other hand, the haptophytes (also known as prymnesiophytes or coccolithophorids) are almost always planktonic marine unicellular flagellates, each cell bearing two smooth flagella and one additional locomotory/feeding filamentous

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appendage, the haptonema [92, for a review see Ref. 2]. The genes contained in the plastid DNA of both heterokont algae and haptophytes have confirmed the red algal origin of such plastids [16, 52, 160]. The arrangement and nature of RuBisCO genes also strongly support this origin. In the red algae the RuBisCO-encoding genes have a proteobacterial origin rather than a cyanobacterial one, as is the case of green algae, land plants and glaucophytes [176]. The plastids of heterokont algae and haptophytes (and also cryptomonads), as we should expect if they have originated from red algae, also have proteobacterial genes for RuBisCO. Moreover, both in red algae as in heterokont algae, haptophytes and cryptomonads, the two RuBisCO genes are encoded in the plastids, whereas in all other plastids the rbcS was laterally transferred to the nuclear genome [72]. Most confusing about its plastid origin are the dinoflagellates, mainly because of the unique features of its plastid genomes, which largely preclude gene amplifying and sequencing. As was already noted, these organisms encompass the largest known plastid diversity, which was soon assumed to be the result of multiple endosymbiosis [56]. About half of the dinoflagellates have plastids, most of which are surrounded by three membranes and contain chlorophylls a and c plus the brown carotenoid peridinin [for review see Ref. 96]. A compelling body of evidences supports the view that this typical dinoflagellate plastid evolved from an endosymbiont red alga. In these dinoflagellates, termed ‘peridinin’ dinoflagellates, the plastid genome is radically distinct from all the other known plastids, consisting of several small (2-6 kb) circular chromosomes, each one encoding a single gene plus a non-coding region [13, 250, 251, 252]. Two separate minicircles encode the SSU and LSU rRNA and, at least in one species, there are seven other minicircles encoding different typical plastid proteins. Phylogenetic analyses of these protein genes [252], as well as psbA [218], tufA and SSU and LSU rDNA [253] retrieved the peridinin dinoflagellate plastids as a monophyletic lineage into the red algal clade of plastids. Moreover, it was supported that a plastid was present in the latest common ancestor of all peridinin dinoflagellates, a group that encompasses most of the extant dinoflagellates; hence the extant numerous heterotrophic dinoflagellates have secondarily lost their plastids [192]. Further, strong support for a red algal origin of peridinin plastid was obtained from the nucleusencoded, plastid-targeted genes, as the ones coding for light-harvesting chlorophyll-peridinin binding proteins [64] and oxygen-evolving

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enhancer 1 (PsbO) protein [115]). However, the history of plastid evolution in dinoflagellates is very complex and some of the dinoflagellates that lost peridinin plastids, seemed to have replaced them later with distinct plastids acquired from other non-red algal eukaryotes [97, 115, 192, 223]. On the other hand, in several species that retain the peridinin plastids most of the typical plastid genes contained in minicircles have been transferred to the nucleus [9, 98, 132], so the plastid genome achieved a maximum of reduction compared with other algae [90]. Perhaps the most surprising finding in plastid evolution was the discovery of the apicoplast, the reduced and non-photosynthetic plastid that has been retained in many apicomplexans, as the important animal pathogens Plasmodium, Eimeria and Toxoplasma [71, 243]. These organelles were first noticed in 1965, through electron microscopy, as an enigmatic membrane-bound compartment of unknown function [see historical review in Ref. 83]. Later it was observed also by electron microscopy, an extrachromosomal circular DNA [128], first assumed to be mitochondrial DNA, although further sequencing showed that this DNA had homology with plastid DNA instead of mitochondrial DNA. Striking plastidial features were discovered in this organelle, as rDNA inverted repeats and plastidial gene clusters [77, 130, 244]. The exact localization of this DNA into the enigmatic membrane bound compartment [130, 156] definitively supported the idea that apicomplexan parasites have secondary plastids [77], bound by four membranes [130]. The apicoplasts have a very reduced and divergent genome (27-35 kb), lacking genes directly related to photosynthesis, but presenting other important functional genes [130, 156], especially the plastid rRNA genes and those for fatty acid and isoprenoid biosynthesis, as well as other important biochemical tasks [76, 237, 243]. Most of proteins functioning in the apicoplast are nucleus-encoded and such genes contain plastid-like targeting sequences [237], confirming the plastid nature of this organelle. Studies in malaria and toxoplasmosis have shown that both the organelle and its genome are essential for those pathogens, which consequently led to an intense research about the nature of this plastid and its use as a drug target [243]. However, to discern the origin of such plastids has been a very difficult task, as has been discussed in the light of new evidences supporting a green algal origin [73, 74] instead of the red algal origin that is commonly assumed [6, 238]. The apicoplast genes are so divergent and AT biased that a

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clear answer about its origin is not possible so far. Some single gene comparisons supported, albeit weakly, that apicoplasts have a red algal origin (for example, the ORF 470 [242] or plastid-encoded SSU rRNA [252]), whereas others favored a green algal (tufA [130]) and even a euglenoid origin (rpoB [75]) and plastid-encoded SSU rRNA [65]). Similarities in gene content and order between apicoplast and red algae plastids [213], as well as non-common genomic structural features between green plastid and apicoplasts [19], were interpreted as supporting a red algal origin, though this interpretation may be unreliable as the apicoplast genome is very reduced and these similarities can be an effect of convergent evolution, as was argued for primary plastids [210]. The strongest evidences in favour of red algal origin of apicoplast do not lie in this peculiar plastid. It is the evolutionary history of some nuclearencoded, plastid-targeted enzymes, namely glyceraldehyde-3-phosphate dehydrogenase (GAPDH [70]), together with analyses of multiple nuclear genes [11] that have strongly supported the view that apicoplasts are ‘red’ in origin. This is better explained if the emerging whole evolutionary framework for the apicomplexan evolution is reviewed in the light of chromalveolate hypothesis. The ‘chromalveolate’ hypothesis

Suggests that from a single ancestor that acquired a red alga as endosymbiont evolved all the extant algae containing chlorophyll c plus numerous related heterotrophic eukaryotes that presumably secondarily lost photosynthesis [32] (Fig. 3). This hypothesis stipulates that chromists and alveolates share a common ancestor and were ancestrally phototrophic. The chromists encompass the cryptomonads, heterokonts and haptophytes, which were supposed to be related on the basis of some shared plastid features, especially the plastidial rough endoplasmic reticulum surrounding the endosymbiont [30], a suspicion that has been strongly supported by analyses of chlorophyll-fucoxanthin binding proteins [63] and recently by combined plastid genes phylogenies [99, 248]. Besides algae, in the chromists are included colorless heterotrophic eukaryotes, as the oomycetes (for example, Phytophthora and other plant pathogens that historically were treated as fungi), opalinids, labyrinthulids, bicosoecids and thraustochytrids, all of which form heterokont flagellate stages. According to the chromalveolate hypothesis, the plastids were lost in these heterotrophic chromists. Recently a

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%$cyanobacterial gene for the enzyme 6-phosphogluconate dehydrogenase in oomycetes was discovered [3], which could be a remnant of the plastid supposed to be originally present in the ancestor of these organisms. The alveolates, on the other hand, include the ciliates, apicomplexans and dinoflagellates, a very diverse group of eukaryotes that share a distinctive system of vesicles beneath the plasma membrane subtended by microtubules (the cortical alveoli), plus tubular mitochondrial cristae and presumptive picnocytotic structures, called micropores [45, 136, 178, 221]. The monophyly of alveolates has been widely supported since first molecular data were available [203] and currently it is not doubted that this assemblage represents one of the major diversification of eukaryotes. If alveolates were ancestrally phototrophic, most of these organisms adopted modes of life radically distinct. For example, the ciliates are mainly predators that presumably lost their plastids, and the apicomplexans are highly derived and specialized obligate parasites with very complex life cycles that maintained a colorless plastid. Finally, half of the dinoflagellates are phototrophs or mixotrophs, whereas the others presumably lost their plastids (or at least the photosynthesis ability) and have adopted a predatory or parasitic way of life [192]. While the chromalveolate hypothesis was proposed in order to minimize the number of secondary endosymbiosis involving a red alga [32, 35], the monophyly of this group had become supported both by plastid genes [248] and nucleus-encoded, plastid-targeted genes [70, 103, 176]. The first hard evidence for chromalveolates was the complex evolutionary history of the GAPDH enzyme in plastid-containing eukaryotes. In photosynthetic eukaryotes there are two distinct isoforms of GAPDH, both nuclear-encoded: one eukaryotic functioning in the cytosol (GapC) and another in the plastid (GapA), which must be targeted to the plastid by adding specific signal sequences [141, 148]. The phylogenies of both GAPDHs have shown that in plants, green algae and red algae the primary plastid-targeted GAPDH is related, as expected, to cyanobacterial and eubacterial GapA isoforms [68, 141] and, accordingly, its gene probably was transferred from the ancient endosymbiotic cyanobacteria to the nucleus of the host cell in the course of the primary symbiogenesis, as occurred with most of the proteins functioning in the plastids. In the secondary plastid of euglenoids, the plastid-targeted GAPDH is also homologous to the cyanobacterial form [105]. However, in the chromists and alveolates with plastids (dinoflagellates and apicomplexans) the plastid-targeted GAPDHs are

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related to eukaryotic cytosolic (GapC) instead cyanobacterial forms, and form a robust monophyletic clade in phylogenies of this enzyme [70, 103, 216]. Such result has been explained by invoking a probable single gene duplication of the GapC of the putative common ancestor of chromists and alveolates, which was already equipped with an endosymbiont red alga [70, 103]. Hence, one of the host’s cytosolic GapC copies, by incorporating targeting sequences, was successfully targeted to the endosymbiont, where it replaced in function the cyanobacterial isoform [70], and this endosymbiotic gene replacement occurred prior to the divergence of the chromists and alveolates, which inherited the shared GapC isoform. Some dinoflagellates, however, severely complicate this history. For example, one dinoflagellate departs from the typical chromalveolate plastid-targeted cytosolic eukaryotic GAPDH isoform, having instead a cyanobacterial like plastid-targeted GAPDH closely related to the Euglena plastid-targeted GAPDH [69, 215]. Likewise, a very rare GAPDH has also been isolated from dinoflagellates, though it is apparently not targeted to plastids [215]. Further support for chromalveolates monophyly has been inferred from the nucleus-encoded, plastid-targeted gene for fructose-1,6bisphosphate aldolase (FBA [176]), whose evolutionary history is also characterized by a unique gene replacement in the putative common ancestor of chromalveolates. Roughly, there are two distinct and unrelated classes of FBA: a eukaryotic isoform (class I) and a primarily prokaryote isoform (class II), which is present in cyanobacteria. In glaucophytes, the nucleus-encoded, plastid-targeted FBA is class II related to cyanobacteria. However, in the red algae the nucleus-encoded, plastid-targeted FBA is a class I isoform, which is supposed to have arisen from a duplication of the cytosolic red algal homologue and an endosymbiotic gene replacement of the cyanobacterial version [176]. Surprisingly, in photosynthetic chromalveolates (there are no plastid targeted FBA in apicomplexans), the nucleus-encoded, plastid targeted FBA is a class II isoform not directly related to the cyanobacterial class II isoforms. Moreover, the phylogenies reveal that the class II isoforms of chromalveolates form a monophyletic clade in the phylogenies suggesting a single origin of all chromalveolate plastids. As such, probably the photosynthetic chromalveolates all acquired their plastid-targeted FBA from some source other than that of all other plastids prior to their diversification in the extant lineages [176].

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Overall, the results from GAPDH and FBA are congruent with those of plastid SSU rDNA [252] in retrieving the apicoplast as a sister group of dinoflagellate plastids, as well as with previously corrected nuclear SSU rDNA phylogenies [233], LSU rDNA [14] and RPB1 [48], that retrieve the heterokonts as related to alveolates to the exclusion of other eukaryotes. Multigene analyses of nuclear genes [11, 12, 166] also support this hypothesis, albeit weakly. However, it is still not easy to put the apicomplexan plastids into a simplified evolutionary picture. A recent study of the nuclear gene for COXII (mitochondrial cytochrome oxidase subunit 2) in apicomplexans did strongly support a green algal origin of apicoplast [73, 74], instead of a red algal origin, as is postulated in the chromalveolate hypothesis. Although these results have been disputed on the basis that no ciliates were included in the phylogenies of COXII genes [238], some striking features of this protein strongly call for a green algal origin of apicoplast. For example, in apicomplexans as in some green algae, the cox2 gene is nucleus-encoded and is present as two chromosomally separated fragments (cox2a, cox2b), whereas in almost all other respiring eukaryotes (including other chromalveolates and red algae) the cox2 gene is mitochondrial and not fragmented [73, 180]. Accordingly, it was proposed that the split cox2 genes were laterally transferred from an endosymbiotic green alga (the ancestor of apicoplast) to the apicomplexans ancestor, as is supported by cox2 genes phylogenies [73]. Besides, the cox2b gene contains an extension in the N-terminal probably related with mitochondria targeting that is shared with green algae, but not with ciliates [73, 74, 180]. This suggests that the apicomplexans acquired the cox2 genes from a distinct source than other alveolates. These evidences strongly support a green algal origin of apicoplast contrary to the GAPDH enzyme claim for a red algal origin. It is still possible that ancestrally apicomplexans harbored plastid derived from red algae (as is predicted by the chromalveolate hypothesis), but these plastids were lost and apicomplexans later reacquired plastids by capturing a green alga, from which they got the cox2 genes [174]. While this controversy remains open, there are convincing evidences that plastid replacement occurred in several dinoflagellates. PLASTID REPLACEMENTS IN DINOFLAGELLATES

Recent molecular data have supported the view that dinoflagellates were ancestrally photosynthetic and that about a half of the extant

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dinoflagellate species have independently lost the photosynthetic ability and live as phagotrophs, although we do not know whether they retain a relict plastid [192]. The most widely distributed plastid in dinoflagellates is the three membrane-bound peridinin-containing plastid, which was probably inherited vertically from an alveolate ancestor and is red algal in origin. However, an irrefutable haptophytic origin is known for plastids of three related species, Karenia brevis (C.C.Davis) G.Hansen & A.F.Moestrup, Karenia mikimotoi (Miyake & Kominami ex Oda) G.Hansen & A.F.Moestrup and Karlodinium micrum (B.Leadbeater & J.D.Dodge) J.Larsen, which contain acylfucoxanthins (19’-hexanolyloxy- and/or 19’-butanoyloxy-fucoxanthin), a kind of carotenoid found only in some haptophytes and in one heterokont [49, 102, 120], instead of in peridinin. Additionally, these plastids are bound by four membranes instead the typical three [205, 102]. Molecular comparisons of plastid-encoded genes (SSU rDNA [223]), psaA, psbA and rbcL [247]) as well as nucleus-encoded, plastid-targeted protein (PsbO [115]) have quite confirmed that these fucoxanthin-containing plastids belong to the haptophyte clade of plastids. Host nuclear SSU rDNA phylogenies retrieve these species as a monophyletic cluster [192], suggesting a single haptophyte acquisition by their latest common ancestor. However, it is not clear from host phylogenies, when such plastid replacement did occur. Recently, a study of three plastid-encoded protein genes, psaA, psbA and rbcL, individually or combined, recovered both the typical peridinin plastids and the fucoxanthin-containing plastids as sister groups forming a monophyletic clade radiating together the haptophytes, also suggesting a haptophyte origin for peridinin-plastids [247]. Nevertheless, the nucleus-encoded, plastid targeted GAPDH clearly support separate origins for the fucoxanthin-containing plastids of Karenia brevis and related species, and the typical peridinin plastids of most dinoflagellates [216]. The most parsimonious explanation for fucoxanthin-containing plastids in dinoflagellates is that these plastids were acquired by a tertiary endosymbiosis between a haptophyte and an ancient heterotrophic dinoflagellate that lost the typical peridinin plastids [192] (Fig. 4). Independent plastid loss and replacement occurred in some members of Dinophysis, which harbor plastids very similar to those of cryptomonads in relation to the main pigments, chlorophylls a and c plus phycobilins, though these plastids are bound by only two membranes and do not

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Fig. 4. Diagrammatic phylogeny of chromists and alveolates with emphasis in dinoflagellates, showing independent losses of peridinin plastids. Several of these secondarily heterotrophic dinoflagellates later acquired distinct plastids from cryptophytes, heterokonts or haptophytes by tertiary endosymbiosis with chromists or green plastids from a green alga by serial secondary endosymbiosis [based on Refs. 41, 68, 69, 70, 90, 97, 98, 112, 192, 217, 223, 239, 247].

contain nucleomorph [197]. It is not clear, however, if these plastids are true integrated organelles or are recent acquisitions that remain functional but which have still not transferred the bulk of their genes to the host [39]. Several predatory dinoflagellates, in fact, retain temporarily functional plastids from its photosynthetic preys (‘kleptoplastidy’ [for example 140]), but these plastids are not capable of reproduction into the host. Host phylogenies based on nuclear SSU rDNA retrieve the Dinophysis species as a monophyletic cluster into the peridinin containing dinoflagellates, supporting the view that these species ancestrally harbored peridinin plastids that were lost. Comparisons of plastid-encoded SSU rDNA [97, 217] and psbA [97, 118] confirm the cryptomonad origin of such plastids, but, as the sequences show a very limited polymorphism among species, it is believed

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that these plastids are recently acquired kleptoplastids and not true organelles [97, 217]. There is also a dinoflagellate with green plastids, Lepidodinium viride M.Watabane, S. Suda, I. Inouye, T. Sawaguchi and M. Chihara [239], which contains chlorophyll a and b as well as carotenoids similar to those of prasinophycean green algae and lacks peridinin and fucoxanthin. Two membranes bound the green plastids of L. viride and there are no other identifiable endosymbiont components. The only molecular data available for these species is the nuclear SSU rDNA of L. viride [192], which confirm that this species ancestrally contained peridinin plastids that were replaced by green two membranebound plastids, though in the absence of plastid molecular data the status of such aberrant plastids is not clear. A very peculiar feature of L. viride is that it produces scales on its cell surface, very similar to those of prasinophycean algae, suggesting that presumably this is host acquired and expresses non-photosynthetic genes from its endosymbiont [239]. The most bizarre endosymbionts are found in two related species of dinoflagellates, Peridinium foliaceum Biecheler and Peridinium balticum (Levander) Lemmermann, which contain an endosymbiotic pennate diatom [41, 112] that still is delimited by its own plasma membrane and contains its own heterokont four-membrane plastids with fucoxanthin, together with its own non-dinokaryotic nucleus, mitochondria, endoplasmic reticulum and other organelles [226]. These dinoflagellates were first recognised by having two nuclei, the typical dinoflagellate (‘dinokaryotic’) nucleus, in which the chromosomes are permanently condensed and the DNA is not linked to nucleosomes [188], plus an endosymbiont typical eukaryotic nucleus [55]. During cell division, these nuclei accomplish a very rare nuclear division [225]. Moreover, it was observed that during syngamy of two host cells, the endosymbiont nucleus of one cell fuses with its compatible of the other cell [40], which strongly suggests that these endosymbionts represent a transitional stage of an ongoing process (the tertiary plastid evolution) and are still not integrated into stable and fully enslaved organelles. Nucleus-encoded SSU rDNA of P. foliaceum [192] showed that this species ancestrally contained peridinin plastids. Compared with other photosynthetic eukaryotes, the plastid genome of dinoflagellates is severely reduced and probably most of original red algal endosymbiont genes for photosynthesis were transferred to the dinoflagellate nucleus [90]. It has been speculated that dinoflagellates are

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particularly susceptible to plastid replacement because the residing nucleus-encoded, plastid-targeted genes obtained from the ancestral red algal plastid can be recycled in the tertiary endosymbiosis by assuming the functions of homologous genes of the new endosymbiont [223]. For PsbO, however, this is not the case, as this nucleus-encoded, plastid targeted gene has a clear haptophyte origin [115]. In any case, it is clear that dinoflagellates are the lineage that have developed more diverse means for endosymbiosis [96]. CONCLUSION

Most of this chapter has been devoted to the chloroplast origin and evolution, which is the basic driving force for the evolution of algae. To talk more about the host origin and evolution is to talk about almost the whole spectrum of eukaryotes, a subject that is growing very fast in the light of current molecular techniques. If we consider the eukaryotic diversification as depicted in the Fig. 2, only one eukaryotic metalineage (Unikonts) does not contain algal groups. It is not surprising, however, that many fungi, which belong to this metalineage, have established very elaborate symbiosis with algae called lichens, as well as many metazoa as corals and hydra. Hence, the symbioses (including endosymbiosis) have really shaped the tree of life. In terms of eukaryotic genome sequencing, of the available list in GenBank [http://www.ncbi.nlm.nih.gov], only three of the nine lineages are represented by at least one complete genome sequence, one alveolate and all the others are in the fungi/ metazoa group or angiosperms. It is clear that the diversity existing in the eukaryotic lineages is still understudied. Investigations of the host genomes already are, and will be even more crucial for the understanding of the origin and evolution of the algal groups. Acknowledgments

We would like to thank Eurico C. de Oliveira for critical review of the manuscript. Alexis Bellorin is thankful for support from the Consejo de Investigación–UDO, Venezuela (CI-N 258/2004) and Mariana Oliveira is thankful for support from CNPq, Brazil.

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Evolution and Diversity of Dinoflagellates: Molecular Perspectives MICHAEL J. BENNETT 1 and JOSEPH T.Y. WONG 2 Department of Biology, Hong Kong University of Science and Technology, Kowloon, Hong Kong SAR, People’s Republic of China

ABSTRACT The dinoflagellates demonstrate unique roles in the marine environment as symbionts in corals and as major causative agents of Harmful Algal Blooms. Molecular phylogenetic investigations of dinoflagellates have centred mainly on the use of ribosomal RNA sequences. However, as a higher level of resolution between members of this order is required more sensitive comparisons are demanded. These methods have employed phylogenies based on concatenated data sets of various proteins, proteins of either nuclear or plastid origin, and the 5.8S rRNA and ITS sequences. Despite the array of markers available there still remains a great deal of ambiguity between the various resultant trees. This problem is exacerbated by the existence of “species complexes” which require even finer analysis using techniques such as RAPD (restriction length fragment polymorphism), lysozyme or toxin profiles. Another complicating factor for the elucidation of an accurate phylogeny is that many dinoflagellate genes come from diverse origins that have been acquired at different times during the course of their evolution. These include genes of bacterial origin as well as a collection of plastid derived genes from different donor endosymbionts. The

Address for correspondence: Dr. Joseph T.Y. Wong, Department of Biology, Hong Kong University of Science and Technology, Clear Water Bay, Kowloon, Hong Kong SAR, People’s Republic of China. Tel: 852-23587343, Fax: 852-23581559, E-mail: [email protected].

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dinoflagellates as a group were once thought of as primitive “Mesokaryotes” but as this review will show, dinoflagellates are in fact highly derived organisms that have followed a different eukaryotic strategy to those taken by the metazoans. Much has yet to be learnt about these amazing organisms, many thousands of species are yet to be described, but over time as more data is gathered the story of their fascinating trek through time may yield a beautiful tapestry of dinoflagellate life. Key Words: Dinoflagellates, evolution, diversity, mesokaryotes

Abbreviations: EST: Expressed Sequence Tags; HAB: Harmful Algal Blooms; ITS: Internal Transcribes Spacer; LSU: Large Subunit; RAPD: Random Amplification of Polymorphic DNA; rRNA: Ribosomal RNA; EFLP: Restriction Fragment Length Polymorphism; SSU: Small Subunit INTRODUCTION

Dinoflagellates are a diversified group, with over 4,000 living and extant members. The group consists of unicellular organisms with almost all known nutritional modes, from free-living photosynthetic, heterotrophic, mixotrophic, phagotrophic, symbiotic to parasitic forms. They are important members of the aquatic environment, and can be found in most marine and freshwater ecosystems. They are the symbiotic zooxanthellae in corals, enabling biodiversity in an otherwise oligotrophic environment. They are also the major causative agents of Harmful Algal Blooms (HABs) and the producers of a wide range of toxins. Many species also produce useful bioactive compounds, most notably the Polyunsaturated Fatty Acids, which are now produced industrially by a species of heterotrophic dinoflagellate. The group also has a distinct nuclear cytology, representing exceptions to many typical eukaryotic characters; including quasi-condensed chromosomes, extra-nuclear spindles, complete lack of nucleosomes and enormous genomes. Molecular studies in the last two decades have generated very interesting data on the group. This chapter focuses on the molecular data in the evolution and diversity of dinoflagellates. EARLY EVOLUTION OF DINOFLAGELLATES

Phylogenetic studies using rDNA (ribosomal DNA) sequences and other genes (actin, beta-tubulin, hsp70, BiP, hsp90, and mitochondrial hsp10) suggest that the dinoflagellates are members of the Alveolates (ciliates,

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apicomplexans and dinoflagellates) and form a clade with the apicomplexans [21, 94]. In a separate phylogenetic analysis with both the actin and hsp90 genes, apicomplexans and dinoflagellates again formed sister groups to the exclusion of ciliates. Oxyrrhis marina, a species with a more typical eukaryotic-like nucleus, formed the earliest diverging sister lineage to the “core” dinoflagellates, and Perkinsus formed the earliest diverging sister lineage to the Oxyrrhis-dinoflagellate clade [44]. The early branching lineage of Oxyrrhis was also confirmed by a study using actin, beta-tubulin and alpha-tubulin [69]. The phylogenetic topology was further supported in morphological analyses of Oxyrrhis and dinoflagellates [76], but not with molecular phylogenetic analysis using SSU (small sub-unit) rRNA [88]. Based on the fossil record of cyst-forming dinoflagellates, the origins of the Peridiniales, Gonyaulaceae, and Ceratiaceae were estimated to be 190, 180 and 145 million years ago (MYA) respectively [39]. However, biochemical examination of dinoflagellate specific biological markers (dinosteranes and 4 alpha-methyl-2 4-ethylcholestane) in concentrated microfossils identified ancient dinoflagellate ancestors from the Early Cambrian, estimated at about 520 MYA [51]. Several other extant groups of alveolates may have close ancestors with the dinoflagellates. Analysis of SSU rRNA sequences of ellobiopsids, multinucleate protist parasites of aquatic crustaceans, place this group among the marine alveolata of dinoflagellates-apicomplexans-ciliates, with close affinity to the dinoflagellates [83]. Furthermore, analysis of ribosomal DNA sequences of microbial communities from Antarctic deep-sea samples identifies two diverse and abundant groups of alveolate sequences, both related to the dinoflagellates. The phylogenetic position of these important deep-ocean microbes suggests a radiation early in the evolution of alveolata [47]. THE DINOFLAGELLATE “MESOKARYOTIC” NATURE, HISTONE-LIKE PROTEINS AND EVOLUTION OF THE LIQUID CRYSTAL GENOME

Dinoflagellates are characterised with many prokaryotic cytological features, including circular DNA, lack of nucleosomes (and major histones), aggregation sensitivity of DNA, etc. Previous observations of continuous DNA replication in the cell cycle have not been

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substantiated with more recent flow cytometric analysis. Similarly, a previous proposal [16] of the group being intermediate between the prokaryotes and the eukaryotes (i.e. mesokaryotes) cannot be sustained in the light of molecular phylogeny that shows the dinoflagellates as being members of the Alveolata [21, 94]. Despite this, recent molecular data does point to the presence of prokaryotic genes in the dinoflagellate genome. Genes which bear homology to their proteobacteria orthologs include RuBisCO from Gonyaulax polyhedra, as well as polyketide synthetase and a rhodopsinlike protein from Pyrocystis lunula [64]. TATA-box binding protein also has intermediate characters between eukaryotic and prokaryotic orthologs [28]. Sequence alignment identified significant similarities between the dinoflagellate chromosomal histone-like proteins of Crypthecodinium cohnii (HCCs) and the bacterial DNA-binding HU and the eukaryotic histone H1 proteins. Phylogenetic analysis also supports the origin of the HCCs from histone-like proteins of bacteria [95]. This suggests that the dinoflagellates have acquired significant parts of their genome from a prokaryotic source- a “mesokaryotic” nature. Both the ciliates and the apicomplexans, the two other major Alveolata, have nucleosomes and the major histones. The loss (or silencing) of the eukaryotic histones from the ancestral Alveolate is a major step in the evolution of the dinoflagellates. Curiously, there are dinoflagellates (for example, the Noctilucales [84]) that have the characteristic dinokaryon during only a part of their life cycle, indicating a possible silencing mechanism of the major histone genes. Pure DNA at high concentrations will spontaneously turn into a liquid crystal state. High resolution ultra structural reconstruction, freeze fracture analysis, as well as circular dichroism and birefringence measurements support the cholesteric liquid crystal state of the dinoflagellate chromosomes (also seen in bacterial nucleoids). The loss of the histones is both the requirement, and probably the cause, of this liquid crystal state. Interestingly, dinoflagellates can be subdivided into three different groups based on three different birefringence measurements [8]. The authors related these observations to the presence of different histone-like proteins for each group. It is possible that the loss of the nucleosomes in the ancestral alveolate was linked to the acquisition of a partial bacterial genome (including the bacterial

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histone-like proteins). This may then have triggered the amplification of the genomic DNA, resulting in the giant genomes (2-200 pg per haploid cell, compared to 3 pg per human diploid genome) observed today. Oxyrrhis marina represents an interesting and phylogenetically problematic dinoflagellate (as mentioned earlier). Nuclear division in O. marina is atypical of the dinoflagellates [25]. The mitotic spindles of O. marina are intra-nuclear [41] and not extra-nuclear as in other dinoflagellates. O. marina is generally quoted as having the major eukaryotic histones and is suggested as the prime evidence for a gradual loss of histones and nucleosomes during the early evolution of dinoflagellates. However, there is only one major chromosomal protein in O. marina with an apparent electrophoretic mobility similar to histone H4 [45]. More recent purification of the major basic chromosomal protein (NP23) of O. marina suggests an apparent molecular size of 23KD [40], which does not correspond to histone H4 (about 11.5KD). Although addition of purified NP23 can apparently induce the formation of “nucleosome”-like structures with DNA [45], this is also a property shared with HU/HLP of eubacteria. More research is needed to elucidate the sequence and functions of the basic chromosomal proteins of O. marina and of the Noctilucales. MOLECULAR PHYLOGENY OF THE DINOFLAGELLATES Overview

Recent multi-genera studies [14, 22, 70, 73, 88] have dealt with the phylogeny of dinoflagellates and this review will summarise and add on to their findings. Studies relating to individual taxa are summarised in Table 1. Common methods employ LSU (Large Sub-Unit) and SSU (Small Sub-Unit) rRNA molecules, with greater resolution being given to the LSU molecule, owing to its faster rate of evolution and its longer length. Other approaches include specific D regions of the partial LSU rRNA gene, or the two ITS (Internal Transcribed Space) regions combined with the intervening 5.8S rRNA molecule. A better result is obtained using concatenated data sets of any of the above molecules, but the major hindrance to these efforts lies in the lack of available data. Protein alignments have also been employed, both singly and in combination as concatenated data sets; hsp90, actin, a-tubulin, btubulin. A later approach is the use of plastid-encoded genes such as

ITS, morphology

ITS, 5.8S rDNA

LSU rDNA SSU rDNA LSU rDNA LSU rRNA ITS1, ITS2, 5.8S, 18S and 28S rRNA,

Calciodinelloideae Scrippsiella, Ensiculifera, “Pentapharsodinium” (species with calcareous cysts)

Calciodinelloideae (species with calcareous cysts)

Karenia species

Prorocentrum

Karenia species

Alexandrium catenella, A. tamarense, A. fraterculus, A. concavum, A. ostenfeldii, A. margalefi and A. pseudogonyaulax

3 novel species

9 species

Karenia umbella sp. nov.

11 monospecies recognized; the molecular data identified two groups of species within one of the genera (Scrippsiella) which can be separated on the basis of the morphology of their resting stages

Scrippsiella form a monophyletic clade; Ensiculifera & “Pentapharsodinium” form another clade

LSU rDNA (D8-D10) LSU rDNA useful markers isozymes

Gambierdiscus 3 species

Comments

Method

Species

Table 1. Phylogenetic studies within different genera of dinoflagellates

[92]

[32]

[27]

[15]

[59]

Table 1 Contd.

ITS may also be a good molecular marker at population level

[17]

[12]

Ref

94 Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

LSU rRNA

SSU rRNA SSU rDNA

Amphidinium species complex

Genus Lessardia and Roscoffia (Podolampaceae)

Halostylodinium arenarium stalked coccoid dinoflagellate

Table 1 Contd.

No clear affinities between this organism and any species included in the analysis

Lessardia and Roscoffia form sister groups

30 strains were analyzed; through morphological and molecular phylogenetic analyses, six distinct species were identified, including Amphidinium trulla sp. nov. and Amphidinium gibbosum comb. nov

[34]

[68]

LM and SEM [55] of some strains correspond with LSU rRNA data

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psaA, psbA and rbcL, or nuclear encoded/plastid targeted genes such as psbO. Most phylogenetic studies of the dinoflagellates to date have made use of the LSU and SSU rRNA molecules. While there may be a plethora of LSU sequences and an abundance of SSU sequences the two data sets do not strongly overlap for many of the important taxa [22]. The results of the various studies have certainly gone a long way towards resolving the relationships amongst the various members of the dinoflagellates, but there is still a great deal of molecular confirmation required to clarify the remaining ambiguities. Chief amongst these inconsistencies is the badly supported backbone of short-branched taxa. The backbone formed by all rRNA trees is usually very weakly supported, consistent with a rapid dinoflagellate radiation [14]. As such, it is difficult to state with absolute certainty using molecular techniques alone, which of the major taxa are basal to the dinoflagellates phylogeny and which are most derived. However, there still exist some very well supported short-branched sub-groups, as well as some longer branched taxa that consistently form clades emanating from the backbone. Quite often a genetic phylotype does not correspond to a taxonomical phenotype. Description of the Major Orders of Dinoflagellates

During the following discussion please refer to Fig. 1 for graphical classification. The Syndiniales, comprising of a potentially diverse group of species [49] is not well supported by molecular data, but the SSU sequences for Hematodinium and Amoebophyra generally form a clade, located near the base of the Dinoflagellate lineage [67], but branching after Perkinsus. The genus Oxyrrhis is problematical because the SSU sequence produces an aberrant derived position while protein data suggest a more basal placement [69]. It is uncertain whether the Syndiniales or Oxyrrhis should occupy the more derived position at this stage. The Noctilucales often branch basally in many phylogenetic trees [67, 74], but when out-groups are removed and more sites are used in the alignment, Noctiluca branches from within the GPP (Gymnodiniales, Peridiniales, Prorocentrales) complex. Certain morphological characters also support a relationship to some Gymnodinialeans.

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The Blastodiniales are represented by SSU sequences for Amyloodinium and Haplozoon only. These two taxa are never found to branch together in phylogenetic analyses and suggest that the Blastodiniales are polyphyletic [46]. In fact Amyloodinium often forms a sister clade to Pfiesteria in a highly derived position in phylogenetic trees and may belong to the Peridiniales instead [43]. Beyond the Syndiniales, the branching order becomes increasingly difficult to unravel using molecular techniques. Molecular data tends to favor the Heterocapsaceae (a Peridiniale), followed by the Suessiales clade or vice versa. But other top contenders for the bottom position may be of the orders Prorocentrales/Dinophysiales. The Suessiales may well represent an intermediate form, as phylogenetic studies rarely place them at the base of the tree, yet they still form a clade to the exclusion of the other more derived taxa. The Suessiales clades include the genus Symbiodinium [74], Polarella [52] and certain Gymnodiniales species such as G. beii and G. simplex. Molecular data have also demonstrated that the genus Symbiodinium forms several sub-clades based on Cp23S (Chloroplast 23S and SSU rRNA analysis [9, 71, 74,], making possible separation of this morphologically homogeneous taxa into individually identifiable species. The GPP complex is composed principally of members of the orders gymnodiniales, peridiniales and prorocentrales [74], but also includes members of the Dinophysiales, and could possibly also include the Noctilucales, as well as some members of the Blastodiniales. This large derived clade is characterised by a very weakly supported backbone containing many short branched, but often well supported sub-clades [14]. The taxonomically defined Gymnodiniales that branch from within the GPP complex form sub-groups, separated from one another by members of the other representative orders. By far the most irksome order is the Gymnodiniales [26], which includes species from the genera Gymnodinium, Amphidinium [55], Gyrodinium, Karenia [32], Karlodinium and Ashikawo, and they are often found scattered in most phylogenetic trees despite supposed morphological congruency. It is probable that many of the traditionally classified taxa within the Gymnodiniales may in fact belong to other orders, as in the case of Karenia and Karlodinium. Gymnodiniales SSU rRNA sequence data indicate that the trees have several major subgroups within the GPP complex, another sub-group that includes most of the genus Amphidinium [55] that groups near to the Gonyaulacales, and one group that forms a very strong relationship to the Suessiales group.

CMYK

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Fig. 1. Schematic illustration of the phylogeny of dinoflagellates based primarily on molecular data Note the deliberate ambiguous position for the common ancestor of the GPP complex and the Gonyaulacales. Also indicated are the likely placements for (a) the gain[67], and (b)loss of armor[88]. The plastids shown, include the following types: (c) the main peridinin chromalveolate derived plastid, (d) the fucoxanthin haptophyte plastid of Karenia and Karlodinium, (e) the cryptophyte kleptoplasts of the Dinophysiales, the prasinoxanthin plastid of Lepidodinium viride, and (f) the diatom derived fucoxanthin plastid of Peridinium foliaceum and P. balticum. (g) Independent losses of the original peridinin plastid have also been highlighted.

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''

The Peridiniales are also a complex paraphyletic group, as demonstrated by molecular trees that tend to place this order in an ancestral position to the holophyletic Gonyaulacales. This order is represented by molecular data for the genera Heterocapsa, Coolia, Ostreopsis, Kryptoperidinium, Peridinium, Scrippsiella [5, 17], Lessardia [68], Roscoffia [68] and Protoperidinium. Another possible relative might be the Thoracosphaerales, with molecular data again providing support for a relationship of this order to other Peridinilean genera, including Peridiniopsis and Scrippsiella. Branching is again unresolved for the Peridiniales, and it is doubtful whether this order has a closer relationship to the Dinophysiales/Prorocentrales or the Gonyaulacales. The Peridiniales, therefore, occupy an important place in the hierarchy of the molecular phylogeny of the dinoflagellates. An ancestral Peridinilean may have given rise to the Dinophysiales, Prorocentrales, Gonyaulacales, some Gymnodiniales, and perhaps Blastodiniales, Noctilucales and Thoracosphaerales [88]. The Dinophysiales are always found within the GPP complex, as are the Prorocentrales. Both orders have a close relationship with one another, and despite the Prorocentrales often producing two sub-clades [27], it is widely considered that these two orders are monophyletic. The Gonyaulacales are characterized in molecular trees as a monophyletic group [74] with longer branch lengths and better bootstrap support. This is due to an apparent faster rate of evolution for the SSU and LSU rRNA molecules for this order. Within the Gonyaulacales, the paraphyletic suborder Gonyaulacineae usually gives rise to the other two suborders, Goniodomineae and Ceratiineae, as well as to taxa such as Crypthecodinium, Thecadium and Halostylodinium [34]. The Phytodiniales are only represented by SSU rRNA data for the taxa Halostylodinium, Hemidinium and Gloeodinium [5]. These taxa do not group together in molecular trees, with Halostylodinium associating with the Gonyaulacales, and the other two taxa always falling within the GPP complex. MULTI-SPECIES COMPLEXES

Multi-species complexes are known in several genera of dinoflagellates, consisting of closely related taxa with highly similar morphotypes, or taxa with a high degree of morphological plasticity. Investigations with molecular markers have characterised some of the groups. Recent studies



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in some of the bloom-forming Gymnodinium species with morphological, ultrastructural and molecular characters have clarified the taxonomic positions of some of these athecate species into four genera [14, 66]. Some of the Karenia brevis (=Gymnodinium breve)-like taxa are now recognized as novel species. This represents a taxonomic platform for further characterisation of related species. Ultrastructural and LSU rDNA sequence analysis of 30 strains of the Amphidinium operculatum species complex identified three novel species. Partial LSU rDNA sequences among strains of A. carterae and A. massartii differed by as much as 4%, and three distinct genotypes could be identified. However, no morphological differences among strains could be observed using LM (Light Microscopy) or SEM (Scanning Electron Microscopy). Very little sequence differences among strains of other Amphidinium species were observed [55]. Previous studies using short partial sequences of the LSU rDNA identified a mean difference of 7 bp between 60 strains of the heterotrophic dinoflagellate Crypthecodinium cohnii [42]. C. cohnii consists of loosely defined and globally distributed taxa that are used in the industrial production of polyunsaturated fatty acids. More genetic markers are required to investigate this biotechnologically important group of dinoflagellates. The origin of the genus Alexandrium, a thecated PSP (Peraletic Shell Fish Poison)-producing species with fossil cysts, could be correlated with the fossil record (39). Despite the aid of the cellulosic thecal plates, the taxonomic positions of A. tamarense, A. catenella and A. fundyense have been problematic. Molecular clades in phylogenetic trees of the Alexandrium tamarense species complex, constructed from LSU rDNA sequences are consistent with geographic distribution, but do not support the three morphologically defined species that constitute the complex [80]. Detailed analysis of LSU rDNA and SSU rDNA sequences of 48 strains from all over the world provided the basis for RFLP-based markers for some of the strains [81, 79]. A recent investigation based on nucleotide sequences of the ITS1, ITS2, 5.8S, 18S and 28S rRNA gene, added a tropical species A. tamiyavanichii Balech to the species complex. A. tamiyavanichii was found to be most affiliated to the Thai-isolates of A. tamarense [92]. Alexandrium genus-specific primers were designed on the 5.8S rDNA region [24]. Seven genera in four orders of dinoflagellates have representatives that have entered a symbiotic mode of existence [5, 63]. Among these, the most commonly encountered taxon is a species-complex loosely

Michael J. Bennett and Joseph T.Y. Wong



defined as Symbiodinium. Earlier studies based on LSU rDNA sequences demonstrated that Symbiodinium-like zooxanthellae represent a collection of distinct species and strains [63]. One host individual can include more than one genotype of Symbiodinium [10]. Seven clades (A-G) of Symbiodinium-like taxa, based on LSU rRNA sequences, are now generally recognized. Isolates from different regions of the world can be assigned to different clades, based on sequence affinities (for example, [75]). Of the eleven species included in the seven clades, four species (S. miroadriaticum, S. pilosum, S. kawaguti and S. goreaui ) are distinguishable based on morphological characters. Some of the other species names may represent synonyms. More recently, phylogenetic trees of the plastidassociated psbA (D1 protein of photosystem II) were compared to those constructed with LSU rRNA gene. The topologies of the two trees were similar, suggesting that these two genes from different organelles (nucleus and plastid) had evolved in parallel in the genus Symbiodinium [86]. These analyses were based on symbiotic dinoflagellates harvested from the host, and planktonic stages of the symbiotic Symbiodinium have not been analyzed until recently [9, 71]. A major review of the diversity of this species complex was published recently [4] and can be referred to for a detailed analysis. More recent studies are summarized in Table 2. INTRA-SPECIFIC VARIATION

Unlike terrestrial plants, phytoplankton can be transported across wide oceanic regions in their life span. Despite this potential mixing of populations, natural phytoplankton harbour high degrees of genetic diversity [23]. Earlier studies on physiological variations of different strains of phytoplankton (including dinoflagellates) have been summarised [96]. However, many of these studies involved only a few strains from wide geographical areas and may have overestimated the overall genetic diversity among taxa. Because “Species Complexes” have been uncovered for several groups of dinoflagellates, the demarcation between intra- and inter-specific variations has become blurred. At the intra-specific level, neither the LSU rDNA nor the ITS regions were observed to have significant genetic diversity in general. Among five physiologically diverse (growth rate and toxin contents) strains of Karenia brevis (=Gymnodinium breve) isolated from Texas and Florida, no sequence differences were observed at the 18S and the ITS regions [48]. Identical ribosomal DNA sequences (ITS1, ITS2, 5.8S and

AMOVA Within populations 87% Between regions 8% Between populations regions 5% Identical sequence to isolates from other parts of the world, despite unique toxin profiles. 27 polymorphic markers, the identification of 40 unique haplotypes. About 40% of the isolates in each sample were identified as one haplotype.

RAPD and allozymes

LSUr DNA D1-D2 sequence RAPD (Six PCR primers)

ITS

Gymnodinium catenatum

Gymnodinium catenatum

Prorocentrum micans (12 water samples in two years)

Scrippsiella trochoidea

The 16 S. trochoidea isolates grouped into five single-strain clades and three multi-strain clades.

within

18S rDNA sequence did not distinguish the isolates, except the West Indies isolate.

18S rDNA sequence

Gambierdiscus toxicus, 7 strains from French Polynesia, Japan and the West Indies

Diversity

RFLP of LSU rDNA PCR products

Method

Gambierdiscus toxicus; 20 isolates

Species

Table 2. Studies on intra-specific variation of dinoflagellates

[53] Some of the ITS haplotypes were distinguishable visually and there are probably cryptic species.

Table 2 Contd.

[82]

[33]

[7]

[65]

[2]

References

>92% of the genetic variance was partitioned within water samples, providing evidence of high levels of genetic diversity.

ITS and 5.8S sequences may be better in differentiate populations.

Other comments

 Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

The mean number of differences among sequences is about 7. Populations grouped into a Malacca Straits group and a South China Sea group.

Phylotypes A, B and C,

Seven unique sequence variants were identified.

D2, LSU rDNA

5.8S, ITS1,ITS2

EGGE coupled with PCR of SSrDNA SSU rDNA

SSU rDNA

Sequencing the flanking regions of two polymorphic microsatellites

Crypthecodinium cohnii species complex, 60 isolates

Ostreopsis ovata, O. lenticularis

Symbiodinium from a range of hosts

Symbiodinium from 7 hosts

Symbiodinium from Caribbean and Mediterranean

Symbiodinium clade B1/ B184 from the Caribbean Sea, from octocorals.

A greater diversity of symbiotic dinoflagellates from corals in South Pacific (clades A, B, and C) than that observed in the rest of the Pacific Ocean (clade C).

One host individual can include more than one genotype of Symbiodinium

15 alleles per locus and gene diversity between 0.632 and 0.974.

13 polymorphic microsatellite loci from this species

Alexandrium tamarense

30 strains were analyzed; there was little difference among strains in partial LSU rDNA for most species, but strains of A. carterae and A. massartii Biencheler differed by as much as 4%.

LSU rDNA

Amphidinium, Comprising 6 species

Table 2 Contd.

Demonstrate specificity of symbiosis in some of the isolates

Benthic dinoflagellates causing cigertera poisoning

Crypthecodinium data was compared to similar studies in Tetrahymena

3 novel species identified

[72]

[75]

[13]

[10]

[61]

[62]

[56]

[55] Michael J. Bennett and Joseph T.Y. Wong

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Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

LSU rDNA) were observed from multiple strains of toxic and non-toxic strains of Pfiesteria piscicda [91]. Only 2-3 base pair differences were observed in LSU rDNA sequences of Alexandrium catenella strains isolated in two separate algal bloom incidences separated by ten years [98]. A very low level of sequence diversity (ITS1, ITS2, 5.8S) was observed for Malaysian strains of Ostreopsis ovata, but the rRNA gene sequences were able to differentiate the South China Sea group from the Malacca Strait group of strains [61]. On the other hand, ITS sequences of 16 strains of Scrippsiella trochoidea could be grouped into 5 single-strain clades and 3 multi-strain clades. Some of the haplotypes were also distinguishable based on minor morphological features [53]. The chloroplast encoded SSU rRNA gene has a higher degree of sequence diversity than the nuclear homologue [90]. The existence of pseudogenes in some conserved loci also led to the generation of distinctive markers on a one-step single-cell PCR, as in the case of the LSU rDNA of Alexandrium catenella in the South China Sea [97], or the SSU rDNA of the North American A. fundyense [78]. For the globally distributed Alexandrium species complex, RFLP(Restriction Fragment Length Polymorphism)-based “strain”-specific genetic markers are possible [81, 79]. For closely related taxons, Random Amplification of Polymorphic DNA (RAPD) and microsatellite DNA seems to be the method of choice for generating differentiation markers. The large genomes of dinoflagellates are known to contain (harbour) substantial amount of non-coding and repeated sequences [1], which are the basis of genetic diversity observed for RAPD. The isolation of specific repeated sequences [54] from the genomes will enable fine scale studies of intraspecific variations. Among the toxic strains of the dinoflagellate Gymnodinium catenatum, very limited variations were observed at the allozyme level when compared with a substantial amount of variation in RAPD analysis. RAPD genetic distances suggest the Australian populations are equally related to the Japanese and Spanish/Portugese populations. Analysis of molecular variance indicated genetic variation was partitioned in the order of: within populations (87%), between the regions (8%), and between populations within region(5%) [7]. In another study of 19 Spanish strains of Gymnodinium catenatum isolated from Andalucia and Galicia, almost identical sequences were obtained for ITS1, ITS2, 5.8S and LSU rDNA. Analysis of RAPD patterns clustered the strains into three different groups, which were different from the dendograms based on toxin analysis [60].

Michael J. Bennett and Joseph T.Y. Wong

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It is widely accepted that genetic diversity is important for adaptation to a changing environment. For phytoplankton blooms, the prevailing environment must favour the blooming species. The availability of intraspecific markers would allow the temporal monitoring of possible population changes in the course of bloom development and decline. Prorocentrum micans is a widely distributed bloom-forming species. RAPD was utilized to assay the genetic variation of 166 single-cell isolates obtained from 12 water samples over the course of two P. micans blooms at Scripps Pier (La Jolla, CA, USA). Twenty-seven polymorphic markers were observed with 6 RAPD primers, identifying 40 unique haplotypes. Over 92% of the genetic variance was partitioned within water samples and 40% of the isolates in one sample belonged to one haplotype. High levels of genetic diversity were observed within temporally separated populations, with differences among bloom and non-bloom periods [82]. With the isolation of more single-cell derived clones, development of single-cell PCR, and the generation of RAPD markers, the intra-specific variation may be more accurately accessed in both a temporal and spatial manner. PLASTID GENOMES OF DINOFLAGELLATES

The first eukaryotic plastid was most likely acquired as a cyanobacterium about 1558 MYA [68], which then diverged to give rise to the separate lineages of the glaucophytes, the red algae, and the green algae (which includes the land plants). This is referred to as the primary endosymbiosis [50]. The ancestor of the alveolates and the chromists (the haptophytes, cryptophytes and stramenopiles) has been labeled a chromalveolate. This ancestral chromalveolate participated in a secondary red algal endosymbiosis about 1300 MYA [102]. With the acquisition of this secondary chromophytic plastid and its accompanying chlorophyll c, the stage is then set for further evolutionary delineation. The act of plastid acquisition and maintenance is generally sponsored by the transfer of plastid genes to the nucleus [30] and the concomitant development of a signal peptide that can deliver the nuclear encoded plastid genes back to the plastid. This system is even more complex where a secondary endosymbiont is concerned with a tripartite signal peptide being used to first ferry the protein to the outer ER-derived membrane of the plastid, and then a further signal to help the protein pass across the inner two membranes [57]. Taken together, these

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developments, however unlikely, have nevertheless allowed the incorporation of foreign plastids into a host organism [35]. With strong support for a successful primary and secondary endosymbiosis, one can reason that much of the necessary molecular machinery to facilitate these processes may still be present in some form or the other in the descendants of these organisms. This would then lead to the conclusion that additional viable endosymbioses are not only likely, but also inevitable. It is also worth noting that plastid loss is not uncommon [20], and it seems to have occurred in the ciliates, some stramenopiles, and has been reduced to an apicoplast in the apicomplexans. It appears that the ancestral secondary plastid of red algal origin [58] has been lost completely in many dinoflagellates [67], or is in a state of advanced degradation in others. The process of plastid loss is preceded by the transfer of plastid encoded genes to the nucleus. As such, these nuclear encoded plastid genes can be used to determine the prior existence of a plastid and its phylogenetic relationship to other plastid containing taxa. Recent examination of dinoflagellate EST (Expressed Sequence Tag) libraries have identified many genes of plastid origin [3, 30] in the nucleus. Phylogenetic analysis of some of these genes has shown that many of them (atpI, atpF, psbO [30, 37]) are indeed of red algal origin. What was surprising, however, was the identification of a number of genes (hemB, tufA, cox2) that were of green algal origin. The results indicated another early tertiary endosymbiosis between a green alga and the common ancestor of the dinoflagellates and the apicomplexans. Apicomplexan and dinoflagellate plastid-targeted glyceraldehyde-3phosphate dehydrogenase (GAPDH) sequences were found to be closely related to the plastid-targeted GAPDH genes of stramenopiles and cryptophytes [20], two other groups that contain secondary plastids of red algal origin. Phylogenetic analyses of GAPDH suggest a single origin of the cytosolic-derived GAPDH in an ancestral chromalveolate [31]. This plastid targeted GAPDH gene resulted from a duplication of the eukaryotic cytosolic form, and replaced the original GAPDH of cyanobacterial origin. Two other putative cytosolic forms of GAPDH (GapC3) are present in the dinoflagellates [19, 87]. In all, there are five different plastid types found in the dinoflagellates (so far). The first, most common and probably the most ancient plastid type, contains peridinin and chlorophyll c2 and

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purportedly represents the ancestral state for this group [20, 58]. The peridinin containing dinoflagellates also share in common a unique organization of the plastid genes on unigenic minicircles [18, 42, 103, 105]. These have been found in Ceratium horridum, Alexandrium tamarense, Amphidinium carterae, Lingulodinium polyedrum, Heterocapsa sp. and Symbiodinium sp. spanning the Suessiales, Gonyaulacales, Peridiniales and Gymnodiniales. Minicircles

The chloroplast genomes of plants and algae, consisting up to 250 genes, are usually encoded on a single circular DNA molecule over 100kbp in size. Chloroplast genes of peridinin-containing dinoflagellates studied so far are either encoded in the nuclear genome, or on mostly unigenic “minicircles” 2-3kbp in size(83, 84). Only 15 of the 45 core chloroplast genes have been found on the minicircles so far (85). One recent in situ hybridization study demonstrates the nuclear extrachromosomal location of minicircles in the dinoflagellate Ceratium horridum (80). Further studies are required to investigate the presence of minicircles in other taxa of dinoflagellates. More interestingly, the minicircles’ non-coding regions (NCRs) are highly conserved within the same species, but are highly diversified inter-specifically, even among species of the same genus. The second category of plastid contains chlorophylls c1 + c2 and 19’hexanoyloxy-fucoxanthin instead of peridinin [89, 100, 101]. Phylogenetic analyses of psbO, psaA and psbA indicate that this plastid arose via tertiary endosymbiosis of a haptophyte alga, probably by an ancestral peridinin containing dinoflagellate. The oxygen-evolving enhancer 1 (PsbO) protein reveals replacement of the original nuclearencoded plastid gene by that of a haptophyte tertiary endosymbiont [37]. rRNA studies of the fucoxanthin containing dinoflagellates Karenia sp. and Karlodinium sp. have not yet distinctly placed them in the dinoflagellate lineage (74). Support lies for either grouping them as a sister group to the rest of the dinokaryotic dinoflagellates [101], or as a more derived group that falls within the GPP complex (Unpublished data). The latter proposal seems the more parsimonious, given the various phylogenetic analyses of plastid SSU rRNA gene sequences from three other 19'-hexanoyloxy -fucoxanthin containing species (Gymnodinium galatheanum, Gymnodinium aureolum, and Gymnodinium

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breve: all members of the Gymnodiniales), that also place the plastid genes of these taxa within the haptophyte clade (89). The other three types of plastid that have been identified in dinoflagellates exhibit varying degrees of reduction of the original endosymbiont to the more streamlined and dependant form of plastid that is commonly imagined. The plastid found in Peridinium foliaceum and P. balticum, members of the Peridiniales, contains fucoxanthin as the primary carotenoid pigment, but stems from a diatom, not a haptophyte [11, 36]. The diatom endosymbiont can still be seen to possess much of its original cellular machinery: nucleus, mitochondria, plastids and ribosomes. These diatom endosymbionts may represent an evolutionary halfway point between engulfment and complete reduction to just a plastid. The plastids found in Lepidodinium viride and Gymnodinium chlorophorum, both Gymnodiniales, contain chlorophylls a and b, and prasinoxanthin rather than fucoxanthin or peridinin [85, 93]. It is the only plastid type identified that has not been sourced from the red algal lineage. The final type of plastid is found in the members of the genus Dinophysis. This plastid is of cryptophyte origin with both psbA and Cp16S rRNA data strongly indicating that the donor plastid is sourced through kleptoplastidy from a species of Teleaulax [38]. Kleptoplastidy is a process whereby the host digests the algal prey, but leaves the algal plastid intact. The type of plastid that a Dinophysis owns apparently depends on the local algal community and is not transmitted vertically through the Dinophysis population. In fact, Dinophysis may be able to acquire plastids from more than one species of cryptophyte. Interestingly, a second class of cloned coding regions was isolated from each population. These psbA and SSU rDNA sequences were evolutionarily more divergent and specifically related to the florideophyte red algae. The mixotrophic habit of Dinophysis raises the possibility that this second class may have come from red algal preys in the food vacuoles of the single-cell isolates [29]. It seems that the dinoflagellates are evolutionary plastidial slavers. Every endosymbiotic event is accompanied by the expansion of the host’s genetic potential after transfer of donor genes to its nucleus [85]. This “rapid” influx of new genes [30], or at least different versions of the same gene, would make this strategy a very powerful method of accruing fitness determinants [50], a sort of “mix and match” policy using plastid genes from different sources to best suit the current situation. One could

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suggest that owning a set of plastid genes is equivalent to having the ability to relate to different cultures, or speak different languages, so that future encounters with potential endosymbionts may yield another successful round of genetic addition. A phagotrophic event for some dinoflagellates is very much akin to having their cake and eating it too! [77]. The dinoflagellates, as a group, can demonstrate this process from the engulfment and digestion of food prey, through to the kleptoplastidy of the Dinophysiales [38], the enslavement of the fucoxanthin diatom, the indentured prasinophyte plastid, the obsequious haptophyte plastid, and finally as evolution rejects the residue of the peridinin plastid, leaving nothing but a few minicircles to mark its passing. CONCLUSION

The genomes of the dinoflagellates have continued to present surprises. Studies of genes and molecular markers of the group have revealed some past evolutionary history and redefined various taxa. However, much more is yet to be learnt from these mesokaryotic cells with the largest known genome sizes, which perhaps have deterred a whole-genome sequencing effort. Another major problem concerning the investigation of dinoflagellate diversity is the difficulty of their culture and isolation. The recent discovery of the selective stimulatory effect of mimosine on dinoflagellates, but lethal effects on other major groups of phytoplankton(99), will provide a powerful tool for the isolation of pure cultures. Another technical problem concerning the estimation of intraspecific variation of unicellular organisms is the detection of molecular markers (for example, RAPD) at the single cell level. With the development of single-cell PCR protocols for dinoflagellates, more studies will be able to address the variation of diversity in both a spatial and temporal basis. Such tools in the biodiversity research of dinoflagellates will also help in understanding the dynamics of harmful algal blooms and coral bleaching. Acknowledgments

Essential sections of this work were supported by grants HKUST6096/ 02M and HKUST6244/04M from the Research Grant Council of Hong Kong to Joseph Wong. Joseph would like to dedicate this paper to Prof. G.E. Fogg and Dr. Patsy Wong for introducing him to the research studies on evolution and diversity of dinoflagellates.



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[48] Loret P, Tengs T, Villareal TA, Singler H, Richardson B, McGuire P, et al. No difference found in ribosomal DNA sequences from physiologically diverse clones of Karenia brevis (Dinophyceae) from the Gulf of Mexico. J Plankton Res 2002; 24(7): 735–739. [49] Massana R, Balague V, Guillou L, Pedros-Alio C. Picoeukaryotic diversity in an oligotrophic coastal site studied by molecular and culturing approaches. Fems Microbiol Ecol 2004; 50: 231–243. [50] McFadden GI. Primary and secondary endosymbiosis and the origin of plastids. J Phycol 2001; 37: 951–959. [51] Moldowan JM, Talyzina NM. Biogeochemical evidence for dinoflagellate ancestors in the early Cambrian. Science 1998; 281(5380): 1168–1170. [52] Montresor M, Procaccini G, Stoecker DK. Polarella glacialis, gen. nov., sp. nov. (Dinophyceae): Suessiaceae are still alive! J Phycol 1999; 35(1): 186–197. [53] Montresor M, Sgrosso S, Procaccini G, Kooistra W. Intraspecific diversity in Scrippsiella trochoidea (Dinophyceae): Evidence for cryptic species. Phycologia 2003; 42(1): 56–70. [54] Moreau H, Geraud ML, Bhaud Y, Soyer-Gobillard MO. Cloning, characterization and chromosomal localization of a repeated sequence in Crypthecodinium cohnii, a marine dinoflagellate. Internat Microbiol 1998; 1(1): 35–43. [55] Murray S, Flo Jorgensen M, Daugbjerg N. Amphidinium revisited. II. Resolving species boundaries in the Amphidinium operculatum species complex (Dinophyceae), including the descriptions of Amphidinium trulla sp. nov. and Amphidinium gibbosum comb. nov. J Phycol 2004; 40(2): 366–382. [56] Nagai S, Lian C, Hamaguchi M, Matsuyama Y, Itakura S, Hogetsu T. Development of microsatellite markers in the toxic dinoflagellate Alexandrium tamarense (Dinophyceae). Mol Ecol Notes 2004; 4(1): 83–85. [57] Nassoury N, Cappadocia M, Morse D. Plastid ultrastructure defines the protein import pathway in dinoflagellates. J Cell Sci 2003; 116(Pt 14): 2867–2874. [58] Nozaki H, Matsuzaki M, Takahara M, Misumi O, Kuroiwa H, Hasegawa M, et al. The phylogenetic position of red algae revealed by multiple nuclear genes from mitochondria-containing eukaryotes and an alternative hypothesis on the origin of plastids. J Mol Evol 2003; 56(4): 485–497. [59] Onofrio GD, Bianco L, Montresor M, Marino D. rDNA-ITS region sequences as a tool for phylogenetic analysis: The case of calcareous dinoflagellates. Naples (Italy): Arti Grafiche Solimene Srl; 1997. [60] Ordas M, Fraga S, Franco JM, Ordas A, Figueras A. Toxin and molecular analysis of Gymnodinium catenatum (Dinophyceae) strains from Galicia (NW Spain) and Andalucia (S Spain). J Plank Res 2004; 26(3): 341–349. [61] Pin LC, Teen LP, Ahmad A, Usup G. Genetic Diversity of Ostreopsis ovata (Dinophyceae) from Malaysia. Mar Biotechnol 2001; 3(3): 246–255. [62] Preparata RM, Beam CA, Himes M, Nanney DL, Meyer EG, Simon EM. Crypthecodinium and Tetrahymena: An exercise in comparative evolution. J Mol Evol 1992; 34(3): 209–218. [63] Rowan R, Powers DA. Ribosomal RNA sequences and the diversity of symbiotic dinoflagellates (zooxanthellae). Proc Natl Acad Sci USA 1992; 89(8): 3639–3643. [64] Ruiz-Gonzalez MX, Marin I. New Insights into the Evolutionary History of Type 1 Rhodopsins. J Mol Evol 2004; 58(3): 348–358.

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[65] Sako Y, Vagi M, Chinain M, Legrand A, Nakahara H, Kurokawa TS, et al. Phylogenetic relationship of ciguatera-causing dinoflagellate Gambierdiscus toxicus with 18s ribosomal DNA sequence comparison. Paris (France): UNESCO; 1996. [66] Salais MF, Bolch CJS, Botes L, Nash G, Wright SW, Hallegraeff GM. Takayama gen. Nov. (Gymnodiniales, Dinophyceae), a new genus of unarmored dinoflagellates with sigmoid apical grooves, including the description of two new species. J Phycol 2003; 39: 1233–1246. [67] Saldarriaga JF, Taylor F, Keeling PJ, Cavalier Smith T. Dinoflagellate nuclear SSU RNA phylogeny suggests multiple plastid losses and replacements. J Mol Evol 2001; 53(3): 204–213. [68] Saldarriaga JF, Leander BS, Taylor FJ, Keeling PJ. Lessardia elongata gen. et sp. nov. (Dinoflagellate, Peridiniales, Podolampaceae) and the taxonomic position of the genus Roscoffia. J Phycol 2003; 39(2): 368–378. [69] Saldarriaga JF, McEwan ML, Fast NM, Taylor FJR, Keeling PJ. Multiple protein phylogenies show that Oxyrrhis marina and Perkinsus marinus are early branches of the dinoflagellate lineage. Int J Syst Evol Microbiol 2003; 53(1): 355–365. [70] Saldarriaga JF, Taylor FJR, Keeling PJ, Cavalier-Smith T, Menden-Deuer S, Keeling PJ. Molecular data and the evolutionary history of dinoflagellates. Eur J Protistol. 2004; 40: 85–111. [71] Santos SR. Phylogenetic analysis of a free-living strain of Symbiodinium isolated from Jiaozhou Bay, P.R. China. J Phycol 2004; 40(2): 395–397. [72] Santos SR, Shearer TL, Hannes AR, Coffroth MA. Fine-scale diversity and specificity in the most prevalent lineage of symbiotic dinoflagellates (Symbiodinium , Dinophyceae) of the Caribbean. Mol Ecol 2004; 13(2): 459–469. [73] Saunders GW, Hill DRA, Sexton JP, Anderson RA. Small subunit ribosomal RNA sequences from selected dinoflagellates: testing classical evolutionary hypothesis with molecular systematic methods. In: Bhattacharya D, editor. Origins of algae and their plastids. New York: Springer; 1997; 237–260. [74] Saunders GW, Hill DRA, Sexton JP, Andersen RA. Small-Subunit ribosomal RNA sequences from selected dinoflagellates: testing classical evolutionary hypotheses with molecular systematic models. In: Bhattacharya D, editor. Plant Systematics and Evolution. New York: Springer-Verlag Wien; 1997; 237–259. [75] Savage AM, Goodson MS, Visram S, Trapido-Rosenthal H, Wiedenmann J, Douglas AE. Molecular diversity of symbiotic algae at the latitudinal margins of their distribution: Dinoflagellates of the genus Symbiodinium in corals and sea anemones. Mar Ecol Prog Ser 2002; 244: 17–26. [76] Schneider EM, Roberts KR. A preliminary phylogenetic analysis of selected dinoflagellates using Wagner parsimony. In: Annu. Meet. of the Phycological Soc. of America, Toronto, Ont. (Canada), 6–10 Aug 1989. [77] Schnepf E. From prey via endosymbiont to plastid: comparative studies in dinoflagellates. In: Lewin RA, The origins of plastids. New York: Chapman & Hall; 1992. 53–76. [78] Scholin CA, Anderson DM, Sogin ML. Two distinct small-subunit ribosomal RNA genes in the North American toxic dinoflagellate Alexandrium fundyense (Dinophyceae). J Phycol 1993; 29(2): 209–216. [79] Scholin CA, Herzog M, Sogin M, Anderson DM. Identification of group- and strainspecific genetic markers for globally distributed Alexandrium (Dinophyceae). 2.

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[96] Wood AM, Leatham T. The species concept in phytoplankton ecology. J Phycol 1992; 28(6): 723–729. [97] Yeung PKK; Wong FTW, Wong JTY. Sequence data for two large–subunit rRNA genes from an Asian strain of Alexandrium catenella. Appl Environment Microbiol 1996; 62(11): 4199–4201. [98] Yeung PKK, Wong JTY, LSUrRNA sequence data from Alexandrium catenella strains isolated from Hong Kong. J Appl Phycol 2002; 14(2): 147–150. [99] Yeung PKK, Wong FTW, Wong, JTY. Mimosine, the allelochemical from Lucaena, selectively stimulates cell proliferation in dinoflagellates. Appl Environ Microbiol 2002; 68: 5160–5163. [100] Yoon HS, Hackett JD, Bhattacharya D. A single origin of the peridinin- and fucoxanthin-containing plastids in dinoflagellates through tertiary endosymbiosis. Proc Natl Acad Sci USA 2002; 99(18): 11724–11729. [101] Yoon HS, Hackett JD, Bhattacharya D. The monophyletic origin of the peridinin-, and fucoxanthin-containing dinoflagellate plastid through tertiary replacement. J Phycol 2002; 38(S1): 40–40. [102] Yoon HS, Hackett JD, Ciniglia C, Pinto G, Bhattacharya D. A molecular timeline for the origin of photosynthetic eukaryotes. Mol Biol Evol 2004; 21(5): 809–818. [103] Zhang Z, Green BR, Cavalier-Smith T. Single gene circles in dinoflagellate chloroplast genomes. Nature 1999; 400(6740): 155–159. [104] Zhang Z, Cavalier-Smith T, Green BR. A family of selfish minicircular chromosomes with jumbled chloroplast gene fragments from a dinoflagellate. Mol Biol Evol 2001; 18(8): 1558–1565. [105] Zhang Z, Cavalier-Smith T, Green BR. Evolution of dinoflagellate unigenic minicircles and the partially concerted divergence of their putative replicon origins. Mol. Biol. Evol. 2002; 19(4): 489–500.

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Evolution of the Diatoms WIEBE H.C.F. KOOISTRA1*, VICTOR CHEPURNOV 2, LINDA K. MEDLIN 3, MARIO DE STEFANO 1, KOEN SABBE2 & DAVID G. MANN 4 1

Stazione Zoologica, Villa Comunale, 80121 Naples Soizi Naples, Italy Laboratory for Protistology and Aquatic Ecology, Department of Biology, Ghent University, Krijgslaan 281 - S8, B-9000 Ghent, Belgium 3 Alfred-Wegener-Institute for Polar and Marine Research, Am Handelshafen 12, D-27570 Bremerhaven, Germany 4 Royal Botanic Garden, Edinburgh EH3 5LR, Scotland, UK 2

ABSTRACT Diatoms are unicellular photoautotrophic eukaryotic organisms whose hallmark is the elaborately shaped and ornamented composite cell wall. The group is highly diverse and species rich. These organisms are the best studied of all the micro-algal groups. In the present study, a phylogeny has been inferred from nuclear SSU rDNA sequences based on a large number of taxa throughout the diatom diversity. This tree is then used to reconstruct evolution of frustule- and protoplasm ultrastructures, and various aspects of the life cycle, such as sexual reproduction and auxospore formation. Possible morphological evolutionary hypotheses are then compared with what is ascertained from diatom palaeontology. Finally, the recent discovery of the occurrence of considerable cryptic diversity is discussed, in what was until recently perceived as a single taxonomically delineated species. Key Words: Diatoms, phylogeny, araphid pennates, raphid pennates, bipolar centrics, radial centrics

Abbreviations: SEM: scanning electron microscopy; SSU rDNA: small sub-unit ribosomal DNA *Address for correspondence: Wiebe H.C.F. Kooistra, Stazione Zoologica ‘Anton Dohrn,’ Villa Comunale, I-80121 Naples, Italy. Tel: +39-081-5833271, Fax: +39-081-7641355, E-mail: [email protected]

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INTRODUCTION

Diatoms (Bacillariophyceae or Bacillariophyta) are among the most diverse groups of eukaryotic microorganisms and there are probably well over 100,000 species [95, 98]. Their often beautifully shaped and ornamented silica cell wall, called the frustule, is the most conspicuous feature [137]. The architecture of the frustule is incredibly diverse [137] and has been the focus of attention of diatom taxonomists. However, organelle arrangements, protoplast ultrastructure, sexual reproduction, and the pattern of development of the auxospore (a unique type of cell, usually a zygote, involved in the regeneration of cell size and shape), also show considerable variation among diatoms [17, 115, 116, 137]. Since diatoms were discovered in the 18th century [1], diatomists have attempted to classify the bewildering diversity of shape and form into natural groupings. Until the 1950s, light microscopy (LM) was basically the only method of study available, but a vast amount of knowledge was nevertheless acquired about diatom structure, despite the relatively low resolution possible (c. 0.2 mm) (for example, [61, 119, 145]). Transmission electron microscopy revealed details of pore structure (for example, [58]), but it was the introduction of scanning electron microscopy in the 1960s (for example, [53]) that led to major surveys of frustule morphology (for example, [118]) and significant changes in classification (summarized in [137]). In the last two decades, Medlin and her co-workers have pioneered the use of molecular genetic data for studying the phylogeny of the group (for example, [108, 109, 110, 111, 112, 113, 114, 115, 116, 117]). Diatoms are of immense ecological importance. At present, they are ubiquitous in the plankton and benthos of marine and freshwater environments (including those with extremes of temperature and pH), as well as in temporarily humid surroundings, such as damp moss or rock [45, 95, 135] they occur from the tropics to polar regions. Several species have managed to carve out peculiar niches: for example, the brine channels of the sea ice, copepod talons, whale skins, the body feathers of diving birds, and lichens [81, 135, 137]. A few species have even become endosymbionts of dinoflagellates [18] and foraminifera [12, 13]. Except for a few species of Nitzschia [83], diatoms are autotrophic. Quite a number of species, however, are capable of taking up small organic molecules, such as amino acids and monosaccharides [121].

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Their contribution to productivity and nutrient cycling is huge. Just a few hundred species (according to current taxonomy, but see section on cryptic diversity below) of marine planktonic diatoms are responsible for around a fifth of all the photosynthetic fixation of carbon on Earth, and they are also the most important biological participants in the silica cycle [44, 46, 57, 95]. Models that seek to predict the effects of increased atmospheric carbon dioxide levels must therefore take into account the ecophysiology of these organisms [8, 31, 154]. Just how long diatoms have been having a significant impact on the biosphere is unclear and insights into the geological age of the group, from molecular or fossil evidence, would be valuable to climatologists in their afforts to discover if there are any historical analogues for the current Earth system, with atmospheric CO2 now at levels unknown since the mid-Tertiary period (see for example, [32]). Diatoms have an extensive fossil record. Their silica cell walls can be identifiable to species level after millions of years (for example, [5, 40, 49, 50]). Moreover, organic molecular fossils specific for particular diatom lineages, such as Rhizosolenia, are detectable in ocean sediments, providing alternative dating points for the origin of diatom lineages [133, 153]. Diatom fossils are used for stratigraphic correlation [4, 39, 43, 158] and also used to infer the existence of particular lineages at particular times in the past. The development of molecular systematics allows us to test morphology-based classifications and they also permit molecular clock calculations [7, 73]. Moreover, if features of the silica frustule in extant diatoms can be shown to be adaptively correlated with environmental parameters, then inferences can be made about the ecology of extinct species, thus permitting reconstruction of palaeoenvironmental conditions. MORPHOLOGY OF THE SILICA FRUSTULE

Diatoms are unicellular or colonial eukaryotes. Their cells are generally ~ 20–200 mm in their maximum dimension, but extremes of c. 2 mm and 4,000 mm have been recorded. The most characteristic feature of diatoms is their unique silica cell wall, the frustule (see Figs. 1-35), which consists of two intricate and highly ornamented structures called thecae. The thecae are slightly unequal, one (the epitheca) being slightly larger than the other (the hypotheca), which it overlaps like the lid and base of a Petri dish (see [137]). Each theca is itself compound and consists of a

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Figs. 1-9.

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large boat- or dish-like valve (epivalve or hypovalve; Figs. 1, 6, 10, 13, 16, 18 and 25) and one to many narrow girdle bands (Figs. 3 and 10) [137]. The primary function of the girdle bands is to allow for expansion during the cell cycle: new bands are added to the distal end of the hypotheca so that the protoplast is at all times enclosed. The valves are often differentiated into a flattened upper surface, called the valve face, and down-turned sides, called the valve mantle (Figs. 1, 16, 18 and 25). Valve and girdle bands are composed predominantly of silica, but they also possess a matrix and envelope of organic material [161], and most species secrete organic material, either as dissolved matter or in the form of gels, mucilage pads or stalks, or chitin threads. The valves consist of a system of silica ribs, which grow out from a circular (Figs. 1, 6 and 13), elongate ([137], p. 341) or linear (Figs. 18, 21 and 25) pattern-centre during valve formation, which takes place within a special membrane-bound sac, the silica deposition vesicle [137]. Small lateral ribs then link the primary ribs to delimit pores (Figs. 16, 25 and 31), which are often occluded by fine networks, flaps or struts of silica (Fig. 35). For the detailed terminology used to describe such pores, see Anonymous 2 [2], Round et al. [137] or Hasle and Syvertsen [57]. In many species, a second layer of silica is laid down above, or occasionally below, the primary system of ribs and pores (Figs. 4 and 24). The secondary layer usually has a different structure and pattern from the initial system of ribs (Figs. 4, 5, 21 and 24) often approximating to Figs. 1-9. Examples of radial centric diatoms (Coscinodiscophyceae sensu Round & Crawford 1981 [136] but excluding Thalassiosirales) and secondarily radial centric diatoms (Thalassiosirales) and their valve features. Fig. 1. Actinocyclus sp., a typical radial centric diatom in internal view. Note the marginal ring of labiate processes. Scale, 5 mm. Fig. 2. Detail of labiate process in 40° tilted view. Scale, 5 mm. Fig. 3. Open valvocopula showing prominent ligula. Scale, 10 mm. Fig. 4. Cross-section of valve showing loculate chambers. Scale, 5 mm. Fig. 5. Detail of external cribra. Scale, 2 mm. Fig. 6. Secondarily radial centric diatom Cyclotella sp. External view. Scale, 5 mm. Fig. 7. Detail of a minute labiate process in the valve mantle of a Thalassiosira sp. Scale, 1 mm. Fig. 8. External opening of a strutted process (strutted processes) of Cyclotella sp. Scale, 0,5 mm. Fig. 9. Internal view of strutted process. Scale, 0,5 mm.

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a hexagonal lattice (Figs. 4 and 5). There may also be extra ribs (Fig. 24), collars (Fig. 11), buttresses and spines (Fig. 10), making the finished valves look like the bizarre creations of a crazed mediaeval architect (see several examples in Round et al. [137]). The girdle bands are generally simpler, but may bear internal flanges (septa or fimbriae) that partially subdivide the cell lumen (see for example, Grammatophora in Round et al. [137] and Campyloneis in De Stefano et al. [22]). As the frustule encases the protoplasm completely and the silica is solid and basically impermeable to water, communication and exchange between diatom and environment are restricted to the lines of pores (‘striae’) lying between the primary rib system of the valves (see for example, Figs 1, 2, 13, 16, 21 and 25); the girdle bands are often porous. In many species, there are clusters of larger or less occluded pores at the periphery of the valve face, forming specialized structures called ocelli (Figs. 14 and 15), pseudocelli, ocellulimbi (Fig. 23) or apical pore fields (Fig. 19), depending on their structure and the taxonomic group in which they occur. These structures are involved in the secretion of mucilage stalks or pads. There are also several types of isolated pores or tubes that are not part of the rib–stria system and do not appear to play much part in stalk or pad formation. These ‘processes’ [57, 137] include some, for example, the ‘tubular process’ that are nothing more than simple tubes. The ‘stigma’ of some raphid diatoms is also a simple tube, except that its inner aperture is sometimes covered by a porous froth of silica. The most common type of process is the labiate process (Figs. 1, 2, 7, 12, 21 and 23). Externally, this is quite variable, opening via a minute hole, a small tube (Fig. 11), or an elongated hollow spine but on the inside it is always flattened to produce a structure resembling tightly pursed lips (Figs. 2, 7, 21 and 23). Where present, there can be one or many labiate processes per valve. Their arrangement is taxon-specific, for example, in a circle near the junction of valve face and mantle (Fig. 1), scattered across the valve face, or clustered in the centre (Fig. 12), or, where there is only one or two, positioned in characteristic places on the valve or mantle (Figs. 7, 21 and 23). A further type of process, the occluded process, consists of an external tube that has been blocked completely at its intersection with the valve face. The functions of tube, labiate and occluded processes and stigmata are largely unknown, although labiate processes have been implicated in cell locomotion in Actinocyclus (Fig. 1) [107]. The function of another

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structure – the strutted process (fultoportula; Figs. 8 and 9) – is better known, having been linked to the secretion of chitin threads [59], which either link sister cells in chains or extend out into the water, increasing drag and promoting suspension [168]. The final structure deserving mention is the raphe (Figs. 25-34), which might be considered as a sixth type of process. It generally consists of a pair of longitudinal slits through the valve (Figs. 25 and 28), each beginning near the centre (Figs. 26 and 29) and extending towards the apex of the valve (Figs. 27 and 32). In almost all cases, possession of a raphe is associated with motility. The raphe slits overlie a differentiated zone of cytoplasm containing prominent bundles of microfilaments, which generate movement over a solid substratum by displacing transmembrane elements that are themselves attached to polysaccharide trail material secreted into the raphe slits [27]. The shapes and positions of the raphe slits vary considerably among raphid diatoms (compare for example, Figs. 25 and 34), but are highly constant within individual species. The raphe represents a structural weakness of the valve. In many species, especially larger ones, the raphe is reinforced with flying buttresses, called fibulae (Fig. 31; [137], p. 45, Fig. 37a-e). In some of these species, the raphe is almost enclosed in a canal (Figs. 33 and 34; [137], p. 45, Figs. 37a-e). TAXONOMY BASED ON CHARACTERISTICS OF THE SILICA FRUSTULE

The varied shapes and wealth of detail in the silica frustules provide many characters that can be used as a basis for delimiting taxa and inferring relationships between them, and these characters are always available for study, even in fossil material, using light or electron microscopy. Indeed, morphology provides much more information for systematics than in many other groups of protists, for example, chlorococcalean green algae or euglenids. Protoplast organization and auxospore form and development also provide useful data in diatoms (for example, [91, 92, 115, 116]), but there are too many gaps in our knowledge for these characters to be used extensively. Diatoms have traditionally been classified into two major groups, based on the organization of the pattern of ribs and pores on the valve face. In centric diatoms, the ribs generally radiate from a ring (annulus).

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Figs. 10-17.

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In the simplest (though not necessarily the most primitive) cases, the annulus is more or less circular and central, and the valve outline is circular (for example, see Fig. 6). These have been referred to as the ‘radial centrics’ (Fig. 1). In the ‘bipolar’ or ‘multipolar’ centrics, on the other hand, the valve outline is not circular and has only one or a few axes of symmetry (for example, Figs. 10, 13 and 16). Here, the annulus may or may not be positioned in the centre and it may not be circular. For instance, the annulus is elongate or elliptical in Attheya [20] and dumbbell-shaped in some Odontella species [127]. All of the different kinds of processes, except the raphe and stigmata, are found among the centric diatoms. Another major group consists of the pennate diatoms, which are organized like a feather (Latin ‘pinna’ or ‘penna’), with an elongate riblike pattern-centre, called the sternum (Figs 18, 21, 25 and 28), from which ribs extend out more or less perpendicularly. Pennate diatoms are usually elongate and their valves resemble the hulls of ships. Pennates can be subdivided into two types: raphid pennates, possessing one or two raphe slits within or closely associated with their sternum (Figs. 25 and 28); and araphid pennates, which lack such slits (Figs. 18 and 21). The types of processes present in pennates include labiate processes – present in most araphid pennates (Figs. 21 and 23) and a few genera of raphid pennates (Eunotia and its allies), and stigmata, present only in some raphids. Clusters of pores involved in mucilage secretion are found in many different genera of both centrics and pennates (Figs. 14, 15, 19 and 23). Figs. 10-17. Examples of bipolar centric diatoms and their valve features. Fig. 10. External view of Biddulphia sp. Scale, 10 m m. Fig. 11. External view of Lithodesmium sp. Note the tubular outer part of the central labiate process. Scale, 5 m m. Fig. 12. Internal view of the central labiate process in Lithodesmium sp. Scale, 1 m m. Fig. 13. Valve exterior of Lampriscus sp. Scale, 10 m m. Fig. 14. External view of ocellus (pore field) of Lampriscus sp. Scale, 10 m m. Fig. 15. Internal view of ocellus (pore field) of Lampriscus sp. Scale, 10 m m. Fig. 16. External view of Ardissonea crystallina, a bipolar centric diatoms originally believed to belong to the pennates. Scale, 10 m m. Fig. 17. Internal view of A. crystallina. Scale, 10 m m.

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Some genera are difficult to fit into these broad, morphology-based groupings. First, the border between the multipolar centrics and the araphid pennates is unclear. Certain diatoms, such as Dimeregramma, Talaroneis and Subsilicea, have parallel ribs on their valve faces, apical pore fields, and what appears to be a sternum – as in araphid pennate diatoms – but they totally lack labiate processes [77, 137]. Toxarium [75] has been considered a pennate diatom because of its elongate shape and benthic habitat, but the best-known species T. undulatum lacks a midrib, parallel ribs, or any trace of labiate processes. The pattern-centre here is neither a simple annulus nor a sternum. Instead, Toxarium seems to have a pattern-centre lying at or near the valve face–mantle junction and running around the whole perimeter of the valve face. This subtends rows of pores outwards, on the mantle, and encloses a more or less irregular scatter of pores on the valve face. A similar arrangement is found in Climacosphenia, Synedrosphenia and Ardissonea (Figs. 16 and 17), which are probably closely related to Toxarium (they share similar girdle and chloroplast morphology), except that on the valve face the pores are organized into rows perpendicular to the peripheral pattern-centre. This type of pattern-centre has been called a bifacial annulus [89], because it subtends ribs on both sides, instead of only outwardly, as in the true annulus. These three genera also lack labiate processes and clearly defined apical pore fields (Figs. 16 and 17; [75]). A final example of a diatom that cannot easily be fitted into the centric or pennate groups is Psammodiscus. This has circular valves and radial symmetry, but appears to have neither an annulus nor a sternum. It has been suggested that Figs. 18-24. Examples of araphid pennate diatoms and their valve features. Fig. 18. Catacombas gaillonii in external view. Scale, 10 mm. Fig. 19. Detail of Synedra capitata showing the apical pore field. Scale, 1 mm. Fig. 20. 60° tilted image of two frustules of Fragilaria cf. crotonensis with interlocking spines. Scale, 2 mm. Fig. 21. Internal view of Fragilaria cf. crotonensis showing apically oriented labiate process. Scale, 2 mm. Fig. 22. Internal view of Tabellaria flocculosa with valve apex lacking a labiate process. Scale, 1 mm. Fig. 23. 60° tilted image of C. gaillonii in internal view. Note the protruding labiate process. Scale, 1 mm. Fig. 24. Chamber structure of C. gaillonii with an internal plain layer of silica in which two marginal rows of silica are located. Scale, 2 mm.

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Figs. 25-35.

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Psammodiscus is related to a group of araphid pennate diatoms (Rhaphoneis and its allies), but that the sternum has been reduced to a point [89, 137]. Another grey zone also occurs between raphids and araphids. Pseudohimantidium [137] possesses a short slit externally that resembles a raphe but internally the slit opens into a row of fused labiate processes [41, 131, 137, 151]. Raphe slits of more normal appearance are present in Eunotia, Actinella and Semiorbis [137] and Eunophora [167], but they are much shorter than in most raphid diatoms and they are not fully integrated into the sternum. Furthermore, labiate processes are present in most of the Eunotia group, whereas they are apparently never found in raphid diatoms with a fully developed raphe system. Greater integration between pattern-centre and raphe is present in Peronia, which is considered to be closely related to Eunotia. Like Eunotia, Peronia is a freshwater genus typical of acid oligotrophic habitats, and it too has short raphes and labiate processes. Comparative morphology has suggested that the raphe originated from labiate processes [54] and that it only later became associated with and incorporated into the pattern-centre [89]; thus the Eunotia group represents an intermediate stage. Figs. 25-35. Examples of raphid pennate diatoms and their valve features. Fig. 25. 60° tilted image of Navicula sp. sensu lato in external view. Note the raphesternum, the pattern of striae. Scale, 5 mm. Fig. 26. Detail of the external, central raphe endings of Cocconeis clandestina. Scale, 1 mm. Fig. 27. Detail of external apical raphe ending of C. clandestina. Scale, 1 mm. Fig. 28. Navicula sp. sensu lato in internal view. Scale, 10 mm. Fig. 29. Cocconeis clandestina in internal view. Detail of the raphe endings converging on the central nodule. Scale, 1 mm. Fig. 30. External view of Bacillaria sp., a genus with a keeled raphe system. Scale, 2 mm. Fig. 31. Internal view of Bacillaria sp., with fibulae subtending the raphe. Scale, 2 mm. Fig. 32. Detail of Fig. 28 showing the internal, apical raphe ending with its helictoglossa. Scale, 2 mm. Fig. 33. Entomoneis sp. with elaborate keeled raphe system. The raphe is located in a subraphe canal connected to the cell lumen through a narrow tube. Scale, 2 mm. Fig. 34. Tryblionella sp. with elaborate keeled raphe system. The space between the connecting tubes is created by fusion and simplification of the keel walls. Scale, 1 mm. Fig. 35. Detail of a hymen, a common areola occlusion in raphid pennates. Scale, 0,1 mm.

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(a) Fig. 36. Contd.

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Fig. 36. Contd.

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(c)

Fig. 36. Neighbour-joining (NJ) phylogeny inferred from maximum likelihood pair-wise distances among nuclear small subunit nuclear ribosomal DNA sequences (SSU rDNA or 18S rDNA) of various diatom species. Sequences of Bolidomonas spp., were included as outgroup. Maximum likelihood calculations were constrained with substitution rate parameters: AÛC = 0.9, AÛG = 2.4, AÛT = 1.2, CÛG = 1.1, CÛT = 3.8 versus GÛT set to 1.0. Assumed proportion of invariable sites along the alignment = 0.4 and a gamma distribution of rates at variable sites with shape parameter (alpha) = 0.5. Assumed nucleotide frequencies: A = 0.25, C = 0.18, G = 0.25, T = 0.32. Bootstrap values (1,000 replicates) have been generated using the same settings as above and have been indicated in the tree. Bootstrap values associated to minor clades to the right have been omitted to avoid clutter. Fig. 36a highlights the centric diatoms, Fig. 36b the araphid pennates, and Fig. 36c the raphid pennates.

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PHYLOGENIES BASED ON CHARACTERISTICS OF THE SILICA FRUSTULE

Several diatom phylogenies have been inferred solely from morphological information (for example, [66, 67, 68, 69, 70, 71, 130, 152, 170]. At first (for example, [148, 152]), this was done by informal arguments, based on subjective (though often logical) analysis of the characteristics of living and fossil diatoms, with consideration also of the geological record. More recently, the cladistic approach has been preferred, in which a matrix of carefully chosen characters is used to reconstruct the most parsimonious hypothesis of relationships. Such a hypothesis is represented as a tree diagram, called a cladogram, which is composed of nested sets of clades, each of them defined by shared derived character states, also called synapomorphies. The results of such studies often provide new insights into the evolutionary history of taxa, allowing adjustments to be made to classifications, but the method also suffers from a few shortcomings. For example, in order to root the cladogram (so that the tree diagram becomes a hypothesis about ancestral states), one needs a group of taxa that is phylogenetically close to, but assumed to be outside the group of organisms being studied. Such a group is called outgroup. However, choosing an outgroup presupposes that we already know a considerable amount about genealogical relationships and there is a danger that the outgroup selected may, in fact, belong within the ingroup. In order to avoid this pitfall, several outgroup taxa should be included, so that any taxa that do belong to the ingroup find their correct place. Phylogenetic inferences of diatom morphology can also be affected by names given to frustule structures [77]. As an example, the sieve-like mucilage secreting structures of bipolar centric diatoms are called ocelli or pseudocelli (Figs. 14 and 15), whereas the very similar structures in araphid pennates are called apical pore fields (Figs. 19, 22 and 23; [137], p. 40). Again, the isolated pore (‘stigma’) present near the central raphe endings in the raphid diatom Luticola would almost certainly be called a labiate process if it had been found in a centric or araphid pennate diatom valve. Apparently, the names assigned to structures do not always depend on what is actually observed, but sometimes where a taxonomist believes that a species belongs in the taxonomic system. Finally, although morphology yields many characters in diatoms, relative to other groups of microscopic eukaryotes, the number is still

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small and interpretation is often arguable. Therefore, morphology-based cladograms often lack resolution. Several to many equally or almost equally parsimonious hypotheses about relationships can be drawn from each data-set, implying extensive parallelism and convergence (homoplasy); recent examples are analyses of Aulacoseira [28] and Petroneis [63]. One can study evolution of a particular characteristic of valve shape or structure via cladistic analyses based wholly on morphological data. However, due to the difficulties outlined above and because the character is itself part of the morphological data used for the construction of the tree, it is desirable to find other, richer and more independent information sources to reconstruct phylogeny, such as DNA sequence data. Here, the characters consist of DNA positions, called sites, and there are only four possible character states (A, C, G and T), which facilitate mathematical modelling of changes amongst them and allows little or no room for a priori taxonomic interpretation and taxonomic authority. The main uncertainty is in the alignment of those DNA sequences that do not code for proteins. Phylogenetically informative characters in sequence data generally outnumber those in morphological data sets by far (for a general discussion, see Scotland et al. [147]). A disadvantage is that only a small part of the genome is sampled and the evolution of a particular sequence may not reflect species evolution. This problem can be minimized through comparison of trees inferred from unrelated sequence regions or separate genomes, such as those located in plastids, mitochondria and the nucleus. A PHYLOGENY INFERRED FROM NUCLEAR SSU rDNA SEQUENCES

A neighbour-joining (NJ) phylogeny inferred from maximum likelihood pair-wise distances among nuclear small subunit nuclear ribosomal DNA sequences (SSU rDNA or 18S rDNA) is presented in Figs. 36a, 36b and 36c. Technical details of the analysis are given in the figure legend. The alignment for this data set is not based on a secondary structure model and these results differ slightly from those obtained from a secondary structure model, as presented in Medlin and Kaczmarska [116] and in Medlin et al. [117]. Also, the number of nucleotides used in the analysis will affect the monophyly of the various major clades. The picoplanktonic flagellates Bolidomonas mediterranea and B. pacifica were selected as

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outgroups because previous analyses have shown a close relationship with diatoms [21, 47]. The tree has been cut into three parts to fit onto multiple pages. Any clade obtaining insufficient bootstrap support (< 50%) has been collapsed into a polytomy onto the clade to the left of it with proper bootstrap support. Therefore, any combination of clades sprouting from such a polytomy could, in fact, represent a monophyletic group, as was found in Medlin and Kaczmarska [116] and in Medlin et al. [117]. Figure 36a shows an almost basal separation with Paralia sol separating from the remainder of the diatoms. The latter group forms a polytomy, bearing several clades of radial centrics and a moderately well supported clade with all multipolar centrics and pennates. The results of the phylogenetic inference of the 18S rDNA data suggest radial centrics are paraphyletic but note that this group is monophyletic in Medlin and Kaczmarska [116] and in Medlin et al. [117]. Furthermore, the multipolar centrics in their turn constitute several clades, that together with a clade containing all pennates, ramify from a polytomy. However, this group is monophyletic in Medlin and Kaczmarska [116] and in Medlin et al. [117]. So, the multipolar centrics could be either paraphyletic or a monophyletic sister group to the pennate clade. The pennate clade, however, has high bootstrap support (98%); a result in agreement with all other diatom phylogenies published to date. Figure 36b shows that the pennates in their turn separate into two major clades, a well supported one (91%) with the araphid pennate diatoms Asterionellopsis, Asterioplanus, Rhaphoneidaceae and Talaroneis (Plagiogrammaceae), and a weakly supported one (59%) with the remainder of the pennates. The latter clade consists of a polytomy with several araphid pennate clades and a weakly supported clade with all raphid pennates (78%). Araphid pennates are therefore paraphyletic. Fig. 36c shows that the raphid pennates are monophyletic (78%). This group consist of a polytomy with several small and one diverse clade. The tree topology is broadly in agreement with those in several other recent studies [30, 75, 76, 115, 116, 153, 155, 156], although we have chosen to collapse branches with < 50% bootstrap support. Raphid pennates are generally monophyletic, whereas araphid pennates are paraphyletic. There is disagreement on whether radial centrics and multipolar centrics constitute mono- or paraphyletic groups, depending on the in- and outgroups chosen, as well as the alignment strategies and

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phylogenetic inferences used. Overall, however, the tree presented here suggests that (i) the radial centric diatoms are more basal (ancestral) than the multipolar centrics and pennates; (ii) one clade within the radial centrics gave rise to all multipolar centrics, including some that have subsequently reverted to radial morphology [the Thalassiosirales, represented here by Thalassiosira, Skeletonema, Cyclotella (Fig. 6), Detonula, Porosira and Lauderia]; (iii) in its turn, one clade within the multipoplar centrics became the pennates; (iv) araphid pennates are basal in the pennates; and (v) the raphe evolved in one clade within this araphid group, to give the raphid pennates. Or, in short, radial centrics begot multipolar centrics, multipolar centrics begot araphid pennates and araphid pennates begot raphid pennates. All of the molecular phylogenies published to date show that centric diatoms are paraphyletic. They are therefore incompatible with the classification proposed by Round et al. [137], if monophyly is, as most people accept, a necessary condition for higher taxa to be acceptable, because Round et al. [137] divided diatoms into three classes: (i) the Coscinodiscophyceae, containing all centric diatoms, (ii) the Fragilariophyceae, containing araphid but not raphid pennates, and (iii) the Bacillariophyceae (the raphid pennates). Only the Bacillariophyceae is a natural, monophyletic group according to results of most molecular phylogenetic analyses, including the one in Fig. 36. To obtain a natural classification, i.e. a classification in agreement with phylogenetic results, Medlin and Kaczmarska [116] have proposed a basal separation in a subdivision Coscinodiscophytina (radial centrics) and a subdivision Bacillariophytina (pennates and multipolar centrics), with the latter being divided into two classes, (i) Mediophyceae, and (ii) Bacillariophyceae. The data are equivocal in several respects, however, because some genes and analyses show topologies incompatible with the idea of a basal dichotomy into radial centrics on the one hand, and multipolar centric + pennates on the other hand. For example, Bayesian analysis of 18S rDNA data shows the radial centrics to be paraphyletic ([116], Figs. 2 and 3 but not in their Fig. 1, which is a subtree of a dataset composed of over 7,500 sequences) and a recent re-analysis of 18S rDNA data also indicates the paraphyly of radial (and multipolar) centrics [156]. However, a Bayesian analysis of 438 taxa using the entire 18S rDNA gene shows Coscinodiscophytina and Bacillariophytina to be monophyletic, with two monophyletic classes: (i) the Mediophyceae or

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the multipolar centrics, and (ii) the Bacillariophyceae [117]. In Fig. 36a, which is also based on 18S rDNA data, monophyly of the Coscinodiscophytina is also rejected but that of the Bacillariophytina obtains mild support (65%). Mono- or paraphyly of the radial centric diatoms depends on ingroup and outgroup choice, the alignment methods (for example, use of secondary structure models), the number of nucleotides included in the alignment and types of phylogenetic algorithm to infer the trees. Note that the trees in Sörhannus [156] and Sinninghe-Damsté et al. [153] do not show any bootstrap values and therefore do not provide information on the support for groupings in their trees, meaning that monophyly is not necessarily rejected for groups that are depicted as being paraphyletic in their trees. They also do not indicate how many nucleotides were used in their analyses. Trees in Medlin et al. [115], Medlin and Kaczmarska [116] and SinningheDamsté [153] have been generated with an alignment obtained using secondary structure models as guiding tools in places where multiple alignment solutions were encountered, whereas those in Sörhannus [156] and the present one have not made use of these models because they include a whole series of unproven assumptions on their own behalf. Within the Bacillariophytina, Medlin and Kaczmarska [116] proposed grouping the multipolar centrics in a class Mediophyceae. This class is monophyletic in their tree taken from the main tree used in the ARB database, but not in their Bayesian or maximum likelihood trees, however, it could be so in the trees present here because the bipolar centric Attheya cf. septentrionalis groups with pennates; nevertheless, it does so with only meagre bootstrap support (57%). Finally, Medlin and Kaczmarska [116] merged the two classes of pennate diatoms proposed by Round et al. [137] – the monophyletic Bacillariophyceae and the paraphyletic Fragilariophyceae – into a single monophyletic class Bacillariophyceae. Such an arrangement receives strong support from Fig. 36 (98%). Other gene regions have been used to address phylogenetic relationships above the generic level. In trees inferred from an extensive rbcL dataset by Mann and co-workers, the topology was not resolved as well as with nuclear 18S rDNA (Mann et al. [102] and unpublished), but the results broadly support the phylogenetic superstructure inferred from 18S sequences. Identical relationships have been shown by the 16S

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the mitochondrial [116], tuf A [114], cox 1 [30], and rpoA gene [33], with the first three from the plastid genome and the latter from genome. Basal centrics

The phylograms published by Medlin et al. [115] and Medlin and Kaczmarska [116], and also our tree (Fig. 36), suggest that Paralia sol may form a separate lineage, basal to all other diatoms. Bootstrap support for this topology is variable – for example, 74% in Medlin et al. [115] but only 65% in our NJ tree (Fig. 3). Sinninghe Damsté et al. [153] did not include Paralia because it was not published at the time. Sörhannus’s [156, fig. 1] parsimony analysis also shows Paralia to be basal, this time together with Leptocylindrus. Paralia and Leptocylindrus are both chainforming centric diatoms with circular valves and a central annulus, but we can think of no morphological synapomorphies (shared derived characters) that might link them. Leptocylindrus has the feature, unusual among diatoms but common in other groups of algae (for example, [36]), that sexual reproduction is linked to the formation of a dormant resting stage [35]. This could perhaps be a primitive character (plesiomorphy) in diatoms. Unfortunately, nothing is known of sexual reproduction and life cycle dynamics in Paralia. In Medlin et al. [117] Paralia is included with other radial centrics. The Radial Centrics (Fig. 36a)

The radial centric diatoms (Figs. 1-5) sprout as a series of clades in the basal polytomy of the tree in Figure 36a. None of these clades correspond exactly to orders in the morphology-based classification of Round et al. [137], except for the two species of Corethron (Corethrales, placed in a separate subclass Corethrophycidae). The Melosira– Aulacoseira–Stephanopyxis–Hyalodiscus clade combines Round et al.’s Melosirales and Aulacoseirales. On the other hand, Round et al. [137] placed Rhizosolenia, Proboscia and Guinardia in the same family, whereas our analysis does not support a close relationship. However, many relationships among radial centric clades remain unresolved and many genera and several families listed by Round et al. have yet to be sequenced. These include some with unusual valve morphology, such as Chrysanthemodiscus (with an extremely simple valve structure and no processes), Arachnoidiscus (with web-like thickenings and a central ring of structures that may or may not

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be labiate processes) or Stictocyclus (with buttressed mantle and large numbers of labiate processes scattered near the centre). Radial centric diatoms have a single pattern centre (Fig. 1), from which the ribs radiate out to the valve perimeter. The valves are almost always circular. There are apparently some interesting exceptions, viz. the semicircular Hemidiscus and Palmeria, which have not been sequenced but are classified close to Coscinodiscus and Actinocyclus, respectively, in Round et al.’s system, on the basis of several similarities in valve ultrastructure. As we describe below (‘Multipolar centrics’), development of bilateral symmetry in diatoms is usually associated with anisometric expansion of the auxospore, brought about by unequal hardening of the auxospore wall by systems of special silicified bands. Such systems are unknown so far in radial centrics, and so the production of bilateral symmetry in Palmeria and Hemidiscus is unexplained (their life cycles have not been studied). The annulus of radial centrics can be small and central, or it may lie around the junction of valve face and mantle, thus enclosing the whole of the valve face, as in some Melosira and Aulacoseira species. Labiate processes, where present, are located in a ring around the valve perimeter or scattered over the valve face. Several genera possess spines at the edge of the valve face, which interlock firmly with similar structures on the valve face of the adjacent sister cell, linking the cells firmly in chains. The Multipolar Centrics (Fig. 36a)

The multi- (bi, tri- etc.) polar centrics (Figs. 6-17) form a series of clades, which together with a clade containing all Thalassiosirophycidae (Figs. 69) and a clade with all pennates, sprout from a polytomy (Fig. 36a). Many members of this apparently more advanced group of centrics and pennates share some form of specialized mucilage-secreting region in their cell walls. These regions take slightly different forms in different groups, but in all cases they appear to be formed by modification of the valve pores in well-defined areas, rather than by modification and grouping of processes, such as labiate or tube processes. The areas of modified pores are usually located at the poles of the valves and in the true pennates, they are therefore called apical pore fields (for example, and Figs. 19, 22 and 23); in bipolar and multipolar centrics, on the other hand, they are mostly referred to as ocelli (Figs. 14 and 15) or pseudocelli. The function of the mucilage secreted through the special

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areas of pores is clearly attachment, either to sibling cells to form chains, or to anchor the diatom to a substratum. In Chaetoceros and Bacteriastrum, however, linkage between cells to form chains is achieved through very fine, silicified, hair-like structures called setae, which are extensions from the valve poles that are extruded out into the surrounding medium after cell division, through holes in the girdle [124, 125]. Figure 36a indicates that Chaetoceros is related to Eucampia and the Cymatosiraceae, both of which have ocellus-like structures and form chains. It is likely, therefore, that the linkage mechanism in the Chaetoceros lineage, involving setae, has replaced an earlier one based on mucilage; indeed, some Chaetoceros species have ocellus-like areas of pores at the bases of their setae [137], but these obviously cannot still function like the exposed ocelli of Eucampia. The Thalassiosirophycidae (sensu Round et al. [137]) constitute an especially interesting group within the multipolar centrics because, as in the basal, radial centric diatom clades, they generally have radially symmetrical, circular valves with a central round annulus (Fig. 6). Indeed, for many years, several species of Thalassiosira were classified within the radial centric genus Coscinodiscus. Molecular phylogenies consistently place the Thalassiosirophycidae within the multipolar centric clade. Furthermore, many trees based on 18S rDNA data show this group to be nested well within the multipolar centrics (for example, [153, 156]), sometimes with moderate statistical support (for example, [116]). Hence, the similarity to radial centrics is apparently deceptive and Thalassiosirophycidae must have acquired radial symmetry only secondarily, with loss of poles and special areas of modified pores. There must also have been loss of the special siliceous elements that create anisometric expansion of the auxospores in multipolar centrics (see the section on auxospore formation). The Thalassiosirophycidae are easily recognized and circumscribed, because they are uniquely characterized by the presence of strutted processes (Figs. 8 and 9), a derived character state shared only by species in this group. In some species (for example, in the abundant marine planktonic diatom Skeletonema), the strutted processes are developed into long tubular extensions externally, which interdigitate and fuse with similar tubes of the adjacent cell to link the cells into chains (see Round et al. [137], pp. 140–141, [139]). These linking structures have evolved independently from those found in radial centrics. Elsewhere, the strutted processes themselves are not linked, but

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create cell–cell linkages by the chitin threads they secrete, or produce the threads as aids to suspension. Labiate processes are also present, but often only one (Fig. 7), positioned on the valve mantle or near the centre of the valve. Bellerochea and Helicotheca (Hemiaulales, Biddulphiophycidae) group in a clade with Ditylum and Lithodesmium. If our topology is correct, this would make the Lithodesmiophycidae (sensu Round et al. [137]) paraphyletic, but this grouping agrees exactly with von Stosch’s [163] concept of the Lithodesmiaceae, based on ultrastructural and life history data. All these diatoms have a centrally located labiate process (Fig. 12), often adorned with an elongate exterior tube (Fig. 11) that lies within the annulus. Often there is a sheet-like ridge at the periphery of the valve face, called the lamella (Fig. 11). The lamella is usually involved in linking cells together into chains. The latter structure generally connects with a similar structure protruding from the adjacent cells. Ditylum, however, is solitary. A central labiate or tubular process, located within the annulus, is also a feature common to the Chaetoceros–Eucampia clade and at least some of the Cymatosirales clade. Both generally have bipolar valves. Biddulphiopsis, Lampriscus, Odontella and Pleurosira, on the other hand, do not have processes lying within the annulus. These genera were placed in different orders of the subclass Biddulphiophycidae by Round et al. [137], a group characterized only by the formation at the corners of the bi- or multiangled valves of elevations bearing ocellus-like areas of pores. Given that such structures are also found in the Cymatosirophycidae and Chaetocerotophycidae (these, but not the Biddulphiophycidae, having synapomorphies that characterize them uniquely), perhaps it is not surprising that molecular data do not support monophyly of the Biddulphiophycidae (Fig. 3, [116, 156]). The bipolar centrics Odontella and Pleurosira (placed in the Triceratiales by Round et al. [137]) possess well-developed ocelli surrounded by a distinct rim and have two (groups of) labiate processes in their valve face, one on either side of the centre. Morphological transformation from one into the other is not difficult to imagine (the principal difference is that the ocelli of Pleurosira are not raised on horn-like elevations of the valve face, whereas those of Odontella are), but molecular data do not encourage this idea. Biddulphiopsis (Biddulphiales) and Lampriscus (Triceratiales) do not group together either, although they both have apical pore fields that lack the

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rim and clear differentiation from the remainder of the valve face seen in Pleurosira and Odontella (i.e. they are ‘pseudocelli’, not ‘ocelli’). These genera have no trace of central or subcentral processes like those in the Chaetoceros–Eucampia and Lithodesmiaceae clades, or the Pleurosira– Odontella clade, respectively. Instead, Biddulphiopsis possesses a row or ring of minute labiate processes near each pseudocellus, somewhat reminiscent of the arrangement in radial centrics. One of the major surprises of recent years [75, 76, 156], reflected also in Fig. 36a, is the recovery of the highly elongate, pennate-like diatom Toxarium as sister to the multipolar genus Lampriscus. Classically Toxarium has been considered to belong to the pennates because of its general shape and habitat [137]. However, its lack of labiate processes and a sternum could have suggested, perhaps, that a classification within the pennates was doubtful. The link with Lampriscus received moderate support (72%) in an analysis with all available multipolar centric sequences, and inclusion of the radial centrics Stephanopyxis nipponica and Rhizosolenia setigera, and of the pennates Asterionellopsis and Asterionella [75]. In other words, Toxarium seems not to be a pennate at all, but an extremely elongated bipolar centric related to Lampriscus. Although the valve shape of Toxarium could scarcely differ more from that of Lampriscus, there are a few similarities in valve structure. For example, both genera lack processes of any kind and the valve structure is simple, lacking any trace of the chambering present in other wellsilicified multipolar centrics [137]. As other multipolar centrics do generally possess processes, their absence in Toxarium and Lampriscus could be due to a secondary loss in this lineage. Moreover, in contrast to true pennates, Toxarium does not have a sternum – the bar-like pattern centre that subtends two series of ribs, one on either side, as in a feather. Instead, the valve face bears a haphazard arrangement of pores [75, 137]. However, there are two longitudinal rows of evenly spaced, ordered pores at the junction of the valve face and mantle; at the apices these rows bend inwards towards each other and meet, apparently forming a complete ring [75]. In fact, one could argue that the genus is really an elongate bipolar centric diatom, with an elongate annulus. Toxarium is probably not an isolated genus within the multipolar centrics. The highly elongate diatoms Climacosphenia¸ Synedrosphenia and Ardissonea, which occur in similar habitats to Toxarium (attached to rocks or seaweeds in the marine sublittoral), also lack a central sternum,

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labiate processes and apical pore fields (despite the fact that all are attached to substrata by mucilage pads). In these three genera (which were classified in three different orders by Round et al. [137]), the nature of the pattern-centre is more obvious than in Toxarium and consists of two longitudinal ribs, one on either side of the valve, near the margin of the valve face, each subtending systems of transverse ribs on either side that meet along the midline of the valve (creating a fault in the pattern that mimics a sternum). The longitudinal ribs are apparently continuous with each other at the apices of the valves (clearly demonstrated in Climacosphenia Mann [89]) and therefore could be interpreted as parts of an elongate annulus. The structure of the girdle in Toxarium and Ardissonea is also similar, and both these genera exhibit rapid locomotion, generated by secretion of mucilage at the poles [75, 128]. However, we must also stress that the annulus of Climacosphenia, Synedrosphenia and Ardissonea differs significantly from that of the centrics like Lampriscus, not only in shape, but also because it subtends ribs and striae both outwardly and inwardly, i.e. the annulus is bifacial and, almost uniquely in diatoms, there is strong centripetal development of the valves (the principal other group in which this occurs is in the Surirellales, a very late-evolving lineage of raphid diatoms). Molecular data suggest, then, that two clades of elongated to extremely elongated diatoms have evolved from multipolar centric ancestors. One pennate-like lineage developed a bifacial annulus, visible today in Toxarium, Climacosphenia and their allies. The other clade is what we will call the true pennates and these developed a different kind of pattern-centre – the rib-like sternum. The origin of this structure is unclear. It is possible that it arose through extreme elongation of the unifacial annulus present in multipolar centrics. An analogy for the pattern-centre in the ancestor to the pennates might be the elongate, dumbbell-like annulus of some Odontella species ([127], Fig. 40e), which is very close to a sternum in its form, except that it still encloses a few pores. One might suggest, therefore, that the true pennates evolved from elongated bipolar centrics with differentiated areas of pores at the poles and pericentral or apical labiate processes. The position of Attheya in our 18S rDNA tree as sister to the true pennates, though only weakly supported, is particularly interesting, because Attheya also has an elongate annulus (though this still encloses pores, as in Odontella). However, other models of sternum evolution can be formulated and

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these cannot be ruled out on the basis of current molecular data. For example, the sternum of the araphid pennate Fragilaria apparently forms by fusion of many aligned, annulus-like structures [104], so that the ancestor of the pennates might best be sought among centric diatoms with multiple circular annuli (if such exist), rather than those with elongate annuli. Overall, however, it must be remembered that sampling of multipolar centric taxa for molecular studies is sparse and patchy, particularly among the genera classified by Round et al. in the subclass Biddulphiophycidae. The araphid pennates (Fig. 36b)

The “true” pennates comprise a well-supported small clade containing Asterionellopsis and Asterioplanus (family Fragilariaceae sensu Round et al. [137]), Rhaphoneis (Rhaphoneidaceae) and Talaroneis (Plagiogrammaceae), and a weakly supported clade consisting of a polytomy of several clades of araphid pennates and a clade containing all the raphid pennates. The well-supported clade (which appears also in other analyses, for example [156]) is a strange amalgam of dissimilar taxa that exhibit considerable diversity of colony shape and frustule structure. Almost the only things they are known to have in common are their pennate organization and simple rib–stria construction, and the fact that all (including the likely relatives of Rhaphoneis: Delphineis, Diplomenora, Sceptroneis, Perissonoë, etc.) are strictly marine. On the whole, these diatoms have apical pore fields, with which any labiate processes present are closely associated, but there are exceptions to every generalization. Molecular data indicate that Asterionellopsis and Asteroplanus are sister taxa, so that the erection of a separate genus for Asteroplanus karianus (= Asterionellopsis kariana) may have been unnecessary. Among the many clades arising from the polytomy in Fig. 36b, there are several that can be reconciled quite easily with other characteristics. For example, the monophyletic genera Hyalosira (called Microtabella in some papers) and Grammatophora group together in one clade with very high support, considering that these marine genera both have internal partitions within the cell (formed by inward extensions of the girdle bands), apical pore fields associated with a labiate process, and zig-zag colonies. However, each of these characteristics is also found in some other araphid pennates; for example, internal partitions occur in the

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freshwater genus Tabellaria and the marine Rhabdonema, and no molecular support has yet been found for a link between these taxa and Grammatophora or Hyalosira, confirming one aspect of the classification of Round et al. [137]. Round et al. also suggested that Grammatophora or Hyalosira belonged with Striatella in Striatellaceae, but this is unlikely, judging by earlier work [76, 116, 156], and receives no support from the present tree, though the possibility of a monophyletic relationship is not excluded. Nevertheless, these three genera share well-developed and well-defined apical pore fields, with pores ordered into a more or less trigonal grid. The pore fields of Striatella are markedly depressed below the surrounding part of the valve and are surrounded by a rim of plain silica; those of the other two genera, on the other hand, are not sunken. None of these taxa possess spines along the valve face perimeter. The next araphid pennate clade in the araphid polytomy constitutes in its turn a polytomy with a series of clades. For convenience, we will refer to this polytomy as the ‘secondary araphid polytomy’; it contains five subordinate clades according to our analysis (Fig. 36b). The uppermost clade contains Fragilaria, Fragilariforma and Synedra (the nomenclature of Synedra and related genera is currently very controversial: the name Ulnaria has been proposed for the freshwater group previously regarded as nomenclaturally typical of Synedra). These taxa share well-developed apical pore fields in which the pores are ordered into a rectangular grid, or in the case of Fragilariforma, staggered. In some species, spines along the perimeter of the valve face link adjacent cells into ribbon-like colonies, but the freshwater ‘Synedra’ species, including Synedra (Ulnaria) ulna lack spines and are not linked into chains. The next clade of the secondary araphid polytomy contains further ‘Fragilaria’ species, although these, unlike those in clade 1, are apparently marine or brackish diatoms, and Synedropsis, which is also a marine genus. None of these taxa bear spines. The Fragilaria lineage within this clade possesses small apical pore fields with pores in a rectangular grid. In Synedropsis, on the other hand, the pore fields are composed of a series of narrow slits parallel to the sternum. All taxa in these two upper clades possess uniseriate striae composed of small pores. Within the secondary araphid polytomy, there is a marine clade with Tabularia and Catacombas, and a separate one comprising Hyalosynedra (all Fragilariaceae in Round et al.’s classification). The apices of these taxa look extremely similar in SEM, with a distinct and markedly sunken apical pore field composed of a rectangular grid of pores. As in other

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clades of the secondary araphid polytomy, there is a labiate process close to the pore field at each apex. The frustules of Tabularia, Hyalosynedra and Catacombas are robust, with a well-marked sternum. The complex areolae are round or transversely elongate and are occluded externally by cribra with distinct cross-bars. In Catacombas and Hyalosynedra, the valves are chambered (Figs. 23 and 24), with development of a layer of plain silica internally that covers the stria, apart from a line of round openings (foramina) along the valve mantle, near the margin. These three genera are brackish (Tabularia only) or marine. The lowermost clade of the secondary araphid polytomy comprises the marine Thalassionemataceae [137]. These taxa possess a single row of areolae along the perimeter of the valve face, which are complex chambered structures occluded externally by cribra, although they are simpler and possibly reduced in many species of Thalassionema. Both Thalassionema and Thalassiothrix form spines along the rim of the valve face but in contrast to those in all other araphid pennates, they develop from the cribra, rather than from the ribs separating the striae. The apical pore fields appear to be reduced to a single pore. The close relationship and uncertain status between paraphyletic Fragilaria – which occurs in chains – and some ‘Synedra’ species – which do not form chains but instead occur in bunches or as solitary cells, – was noted by Lange-Bertalot [82] and point to the need for further taxonomic changes beyond those advocated by Williams [169] and Williams and Round [171, 172, 173], Hasle et al. [56] and Round et al. [138]. Chain-forming linkage structures in these genera do not signify relatedness. On the other hand, it is clear that the classification of almost all Synedra- or Fragilaria-like diatoms into a single undifferentiated Fragilaria, as initially suggested by Krammer and Lange-Bertalot [78], is untenable. Next in the primary araphid polytomy are two small marine clades with species of Licmophora (Licmophoraceae) and a clade with Cyclophora and Pseudohimantidium. All of these are attached benthic marine diatoms. Pseudohimantidium and Protoraphis possess elaborate composite labiate processes opening into a groove externally [41, 42] and it was initially suggested that the raphe might have evolved by further fusion of such multiple labiate processes [79, 151]. Our molecular tree renders such a hypothesis not very likely. We can find no morphological synapomorphies linking Cyclophora and Pseudohimantidium, but these

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genera and Licmophora (which were placed in different families by Round et al. [137] and have markedly different valve shape) share a similar striation pattern and simple rib-stria structure, and their apical pore fields are generally somewhat reduced or modified, to: (a) a single row of slitlike pores perpendicular to the sternum in Licmophora [60], (b) a series of slits fanning out from a hyaline area near the termini of the sternum in Cyclophora, or (c) what appears to be a series of rows of small pores surrounding the composite labiate processes in Pseudohimantidium. None of these genera possess spines. The penultimate clade in the primary araphid polytomy branches into an upper clade with the freshwater taxa Asterionella, Diatoma and Tabellaria and a lower clade with the marine genus Nanofrustulum and the freshwater genus Staurosira (formerly classified within Fragilaria because of the characteristic which they share, i.e. both form long ribbon-like chains). Staurosira may be paraphyletic. All the genera mentioned share uniseriate or multiseriate striae composed of tiny pores. The Asterionella–Diatoma–Tabellaria clade includes taxa with short, simple spines placed in a row lying along the perimeter of the valve face and extending through the apical pore fields. There are also taxa that lack spines. All of these taxa tend to form zig-zag or stellate colonies, which are linked by mucilage pads secreted through the apical pore fields at one or both poles. The pores in these pore fields are organized in vertical rows, which are offset from those in neighbouring rows in a seemingly random fashion. They have simple rib-stria systems, tiny round pores in the striae, and the striae are rather irregularly spaced, compared to those of most other pennate diatoms (accompanied by thickening of the wider ribs in Diatoma and the presumably closely related Distrionella). Nanofrustulum and Staurosira possess highly reduced apical pore fields (in Nanofrustulum they consist only of a simple pore) and lack labiate processes entirely. The valve face perimeter is adorned with elaborate spines that link cells into ribbon-like colonies. Rhabdonema is the last branch from the primary araphid polytomy. Details on this diatom’s complex frustule ultrastructure can be found in Round et al. [137] and Pocock and Cox [129]. The solitary position of Rhabdonema corroborates the suggestion by Round et al. [137] of a lack of affinity with any other araphid diatoms. The complex girdle structure and the presence of many labiate processes along the sides of the sternum are apparently without parallel elsewhere, although some araphid taxa,

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such as Entopyla, have yet to be studied in detail using SEM. von Stosch [159] suggested that the genus is transitional (a “Bindeglied”) between araphids and centrics, (i.e. the sister lineage to all other pennates), because of peculiarities of its sexual reproduction (referred below). The tree topology does not support this idea, but the bootstrap support (59%) for the primary araphid polytomy is not high and so a more basal position of this genus remains possible. Hence, although molecular data do support some aspects of morphology-based classifications of araphid pennates, for example, links between Thalassionema and Trichotoxon, and the need for re-classification of Synedra sensu lato and Fragilaria sensu lato, they also show considerable deficiencies, especially in the Fragilariales sensu Round et al. [137]. This is currently a large heterogeneous order characterized only by the absence of any of the more sharply-defined diagnostic features (putative synapomorphies) of each of the remaining families and orders. The Fragilariales sensu Round et al. [137] is not monophyletic, because it contains Asterionellopsis as well as some but not all taxa in the primary araphid polytomy (it contains all taxa in the secondary araphid polytomy except the clade with Thalassionema and Trichotoxon, and all taxa in the clade with Asterionella, Diatoma, Nanofrustulum and Staurosira). The Raphid Pennates (Fig. 36c)

Raphid pennate diatoms (Figs. 25-35) form a single clade in Fig. 36c. There is only moderate bootstrap support (78%) for the clade, but other published trees (for example, [116, 156]) also show the raphid group as monophyletic. Apparently, therefore, the true raphe has been acquired only once. It is believed to have evolved through elongation of labiate processes at the distal ends of the valve in an ancestral araphid pennate lineage ([54, 89] [137] p. 41); which lineage this was and whether it was marine or freshwater, are unclear. In most cases, the raphe is fully integrated into the pattern-centre of the diatoms, forming a composite ‘raphe-sternum’ [19, 89]. The raphid group is the largest among the diatoms, in terms of genera and species, even though it is the youngest of the major lineages, with no confirmed fossil evidence for the group before the Tertiary period. We will not undertake an in-depth treatment of evolutionary patterns in the raphid pennates, because many relevant sequences remain unpublished and another study is under preparation dealing with this topic. We will instead select a few of the more important conclusions that can be drawn from published data and Fig. 36c.

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The nature of the Eunotiophycidae, which includes the genera Eunotia, Eunophora and Actinella (Fig. 36c) has long been discussed. Traditionally, they have been placed at the beginning of the raphid system, as ‘primitive’ raphid pennates (‘raphidioids’) because their raphe slits are very short and simple, being apparently less evolved than the longer slits present in most other raphid diatoms, which are generally more actively motile. Detailed examination of their morphology by electron microscopy and consideration of valve morphogenesis has shown that the Eunotia raphe is not enclosed within the sternum, as in other raphid pennate diatoms, but lateral to it, interrupting the rib–stria system subtended by the sternum [89, 104]. In Fig. 36c this genus is recovered in a clade on the basal polytomy of the raphid pennates. If this group is indeed basal within the raphid pennates, then the more developed type of raphe system, characteristic of all other raphid diatoms, may have developed from the raphidioid raphe through elongation and reflexing of the slits until they lay alongside and then became integrated into the sternum [89]. Some molecular trees (for example, in [156]) support the idea that Eunotia and its allies are basal. However, in Fig. 36, their position is unclear. The raphidioids are clearly a monophyletic group, but the data leave open the possibility that they are a derived group, their curious raphe system being produced via secondary reduction and modification (cf. further reduction and loss of the raphe in some raphidioids: [72]). In Medlin et al. [117], the raphid diatoms diverge initially into two clades: one is the Eunotiophycidae and the other contains all remaining raphid diatoms. As with the araphid pennate and centric diatoms, molecular data sometimes reveal relationships that were previously unsuspected but sometimes confirm or extend conclusions drawn from morphological data. For example, the close relationship between Navicula, Seminavis and Pseudogomphonema predicted from valve, raphe, girdle and chloroplast data (summarized by Round et al. [137]) is confirmed by 18S rDNA data (Fig. 36) and rbcL data [63], as is the ‘gomphocymbelloid’ cluster (Round et al. [137]) of Anomoeoneis, Cymbella, Encyonema, Placoneis and Gomphonema (Fig. 36c) [63, 116, 156]. Many of the latter group of taxa possess apical pore fields and stigmata, which are not found in their apparently closest relatives, represented in Fig. 36c by Cocconeis. A further confirmation of previous ideas is the link between Pinnularia, Sellaphora and Fallacia, which all have a similar raphe system and share

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several features of chloroplast organization, including invaginated pyrenoids [137]. Among unexpected results is the demonstration that the Surirellales, traditionally considered the most ‘advanced’ of the main lineages of raphid diatoms (because the raphe system is both subtended internally by silica bridges and highly modified in position and extent, so that it occupies the whole perimeter of the valve face), is allied to a group of Amphora species (Fig. 36c) [156]. Re-examination of the characteristics of Amphora and the Surirellales in the light of molecular data does not reveal any features that are obviously synapomorphies for this clade. Both usually have one chloroplast per cell, but that is where the resemblance ends. However, incorporation of other genera into molecular phylogenies, such as Auricula with its Amphora-like symmetry but Surirellales-like valve and raphe structure, may help to reveal the structural and developmental changes that have occurred during the evolution of this group. In Medlin et al. [117] the Surirellales are the last raphid lineage to diverge. Last but not least, molecular data show that previous attempts [137] to revise the classification of the ‘monoraphid’ diatoms were too timid. In monoraphid genera, the frustules have dissimilar valves, one bearing a raphe system and the other lacking one, so that the pattern-centre resembles the sternum of araphid pennates. Sometimes, these diatoms have been regarded as early divergences within the raphid group, but there is no evidence of this from molecular data and in fact the rapheless valve develops by infilling of the raphe-system during valve formation [9, 87]. The only two monoraphid genera of any size recognized until 1990 were Cocconeis and Achnanthes; Cocconeis had elliptical valves and the frustules were straight in girdle view, whereas Achnanthes generally had more elongate valves and the frustules were bent in girdle view. Achnanthes was split when it was realized that there were major differences in pore structure, internal raphe structure and chloroplast morphology between different groups with the genus (for example, [137]). However, both Achnanthes and the newly resurrected Achnanthidium were retained within the same family. Figure 36c shows, however, that the monoraphid condition has evolved at least twice independently, because Cocconeis (and some diatoms previously classified in Achnanthidium, for example, Planothidium lanceolatum: [116, 156]) group with the gomphocymbelloid genera, whereas true Achnanthes

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species appear in a very different region of the raphid tree. Clearly, these genera cannot continue to be classified in the same family or order. PHYLOGENETIC SIGNAL IN THE LIFE CYCLE AND AUXOSPORE ONTOGENY

Diatoms are diplontic organisms. Their life cycles usually consist of a prolonged period of mitotic division (months to years), followed by a short period of sexual reproduction comprising gametogenesis (hours) immediately followed by cell fusion and the formation and development of auxospores (hours to days). The haploid gamete stage does not undergo asexual mitotic divisions, except in rare cases of haploid parthenogenesis in culture [17]. Mitotic cell division in diatoms shows several peculiarities. The location of the rigid parental epi- and hypotheca dictates the orientation of the new thecae (such a process is termed autopoiesis [48]) and dividing cells cannot alter their plane of division without producing cells of abnormal shape and size. Daughter cells are formed back-to-back, with two possible outcomes: separation of the cells once the new thecae are sufficiently mature, or the formation of uniseriate filaments. Diatoms are unable to form branching multicellular thalli, like those encountered in green, red and brown algae [95], although a close approach to multicellularity is achieved by those diatoms (a few species, belonging to disparate clades of raphid diatoms) that produce mucilage tubes [137]. Another consequence of the rigid silica cell wall and the fact that new wall elements are produced within the cell (in silica deposition vesicles), is that mitotically reproducing diatoms almost inevitably decrease in size, although usually by only a fraction of a micrometre per division. The rim of the epitheca fits over the rim of the hypotheca and is therefore slightly larger. Consequently, the daughter cell inheriting the parental hypotheca will be smaller than its parent because the parental hypotheca now has become the epitheca (see [137]). The principal way for a diatom to avoid diminishing to nothing is through the expansion of a specialized zygote, the ‘auxospore’. During the life cycle, large cells are at first unable to undergo sexual reproduction and size restoration via an auxospore. As they continue to reduce in size, however, they reach a critical size threshold, after which they gain the potential to become sexual, if other conditions (light, nutrients, internal factors, the presence of sexually compatible partners in the case of

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heterothallic species) permit. During the formation of gametes or shortly thereafter, the old cell walls are shed. The zygote (auxospore) formed by fusion of the gametes then becomes surrounded by a wall unlike that formed at any other stage of the life cycle, which is either wholly organic or composite, consisting of organic and silica elements (scales or bands); in either case, the auxospore wall allows expansion, which is a wellcontrolled process, during which the basic outline of the new enlarged cells is regenerated, although there is often some further modification of shape during the formation of the initial cell (the first thecate cell produced by the auxospore) or its first few divisions (for example, [16, 99]). Once expansion is complete, new silica cell walls are produced within the auxospore wall. Just a few diverse species of centric and pennate diatoms are known to be able to perform a similar trick also during their vegetative phase [17, 160]. In this ‘vegetative enlargement’, cells shed their cell casing, inflate (often irregularly), and then construct an entirely new pair of thecae; a special wall like that of the auxospore is apparently not produced. Diatoms exhibiting vegetative enlargement can usually also enlarge via sexually produced auxospores. Rarely, sexual formation of auxospores via allogamy is replaced by automixis or apomixis [17], which seem to represent modifications of allogamous pathways of development. In diatoms, sexual reproduction was believed always to be possible between genetically identical clonal individuals [23]. Recent data, however, have shown that accepted generalizations about the life cycle require considerable modification [17]. Mechanisms to prevent or hinder inbreeding have been demonstrated in many pennate diatoms (for example, [132]), whereas knowledge of mating systems in centric diatoms is still very poor. In addition, new observations indicate that the link between size and sex in diatoms is much more complex than had been thought [14, 17]. The formation of dormant resting stages or cysts, a key adaptive strategy in many other algae, where it is often linked to sexual reproduction, is known in diatoms but it has been confidently reported in only relatively few taxa. They are easily detected if the stages differ from vegetative cells not only physiologically but also morphologically, in which case they are referred to as ‘resting spores’. Resting spores are principally found in radial and multipolar centrics, with only rare records among pennates [106]. In other cases, the dormant stages are only

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physiologically differentiated, in which case they can be referred to as ‘resting cells’. These are much more difficult to detect and are probably more common than it currently appears. The functions of resting stages are probably varied (for example, [105, 106]). No convincing phylogenetic signal is yet detectable in the occurrence and types of resting stages in diatoms. We emphasize that auxospores are not resting stages, despite their name; however, in Leptocylindrus, a resting spore is formed within the auxospore [35]. Gamete Formation

Meiosis and gamete formation vary considerably among diatoms [17, 23, 29, 137]. All centric diatoms studied so far produce motile flagellated micro-gametes (sperm) with one anterior flagellum, and nonmotile naked macro-gametes (eggs). In some centrics, the sperm develop merogenously, i.e. they separate from the parental cell, leaving the plastids behind in a remnant cell without nuclei. In Stephanopyxis (a radial centric), the elimination of the plastids into a residual cell devoid of nuclei occurs before the remaining cell proceeds to form the gametes. In other centric diatoms, particularly in multipolar centrics (including Thalassiosirophycidae), the sperm form hologenously: the whole of the spermatogonial cell divides into equal portions, plastids included. In our earlier review [76], we found a reasonable correlation between the method of sperm formation and phylogeny, most radial centrics producing sperm merogenously and most multipolar centrics producing sperm hologenously. The more complete review by Jensen et al. [62] shows a more complex pattern, with merogenous and hologenous spermatogenesis sometimes occurring in different species of the same genus (for example, Melosira, Odontella). In Medlin and Kaczmarska [116], it was concluded that the Coscinodiscophytina had predominantly merogenous, and the Bacillariophytina had predominantly hologenous spermatogenesis, despite the many exceptions. The presence of the ability to form both types of sperm in both clades even in the same species was interpreted as an ancestral polymorphism. Apparently, the life cycles of many additional species need to be examined before phylogenetic pattern can be distinguished with respect to spermatogenesis. The araphid pennate Rhabdonema produces macro-gametes and micro-gametes, like centric diatoms, but here the micro-gametes lack a

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flagellum [159]. As far as is known, all other pennates reproduce sexually by means of isogamy [17, 93]. Each cell produces one or two similar gametes and there is no differentiation into large immobile and small mobile gametes, although there may be differences in gamete behaviour (‘physiological anisogamy’). Thus, the paraphyletic centric diatoms are oogamous and have flagellated micro-gametes, whereas the monophyletic pennates reproduce via non-flagellated gametes. This independent corroboration of the molecular phylogeny is convincing because it is based on many reports of sexual reproduction in many different genera (for example, 17, 38). However, it should be noted that there is as yet no information on sexual reproduction in the well-supported Asterionellopsis–Asterioplanus– Rhaphoneis–Talaroneis clade. The fact that Rhabdonema produces microgametes, even though these are non-flagellated, can be interpreted in two ways. The species may represent a position between the centrics and other araphid pennates. In that case, the micro-gametes of the araphid pennate ancestors first lost the flagellum, then they gave rise (1) to the lineage with Rhabdonema, which retained the micro-gametes, and (2) to the lineage containing the remainder of the pennates, which subsequently evolved towards isogamy. On the other hand, the microgametes of Rhabdonema may result from a reversal to the ancestral state in an otherwise advanced lineage. In fact, the lobed plastid of Rhabdonema could hint at a position in between araphids and raphids. Sexual reproduction in the pennate-like centric diatom Toxarium and its relatives has never been studied. Yet, given its pennate lifestyle and general shape, knowledge of its mode of sexual reproduction becomes highly significant. If, for whatever reason, isogamy is advantageous in pennates, or more generally in benthic attached diatoms, then Toxarium may have developed such a mode of reproduction independently. Like the pattern of spermatogenesis, the method of production of the macro-gametes also varies within centric diatoms, and there is little obvious concordance with the molecular phylogenetic tree, although there is no comprehensive review for this character. Round et al. [137] describe the fate of nuclei at the two meiotic divisions. In many centric diatoms, only one of the four haploid nuclei produced by meiosis ends up in a functional macro-gamete. However, taxa differ in whether or not there is a cell division at meiosis I. In the multipolar centrics Chaetoceros and Skeletonema, and in the radial centrics Melosira and Stephanopyxis

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both meiotic divisions occur in an undivided cell, the superfluous nuclei being resorbed into the cytoplasm. In Cerataulus and Biddulphia, which are presumably multipolar centrics, there is an unequal cytokinesis after meiosis I, producing a ‘polar body’ that subsequently degenerates. In a third variant, there is an equal cell division after meiosis I and two egg cells are produced. This is known to occur only in multipolar centrics, such as Lithodesmium and Odontella. The interesting feature is that two eggs are produced per mother cell only in multipolar centrics, and it is this group of centrics that is sister to the pennates, which we can be fairly sure (from the distribution of different types of gamete formation among lineages) primitively produce two isogametes per mother cell. In Rhabdonema, which is intermediate between pennates and centrics in having differentiated macro- and micro-gametes but not flagella, two species produce two macro-gametes per mother cell, whereas the third produces a single macro-gamete and a large polar body [23]. Pennate diatoms produce large and sedentary or rather slow-moving gametes and fertilization can only occur if the gamete-producing cells are somehow brought close together. Thus, gametogenesis is closely linked to the pairing up of compatible partners [17, 93, 95, 137]. The actual fusion involves swelling of gametes to bring them into contact, or amoeboid movement of the gametes between the partner cells, or a complete shedding of the walls before fusion. The modes of pairing and gamete behaviour vary greatly among pennates, although particular genera or families often show consistent patterns of behaviour. For example, Actinella and Eunotia, clearly related according to molecular data (Fig. 36c), also share a similar type of sexual reproduction involving formation of just one gamete per gametangium and plasmogamy via an apical papilla [101]. More species need to be included in the molecular phylogeny before the patterns of distribution of sexual characteristics can be examined in detail and further information is still required on the ‘exceptional’ sexual behaviour of Rhabdonema [17]. Auxospore development

After gamete fusion and a short period of internal rearrangement of nuclei and chloroplasts, the auxospore begins to expand, to restore its original cell size (see for example, [17, 87, 88, 97, 146, 162]. Round et al. [137] summarize the principal features of auxospore wall ontogenesis, noting that zygotes first form an outer polysaccharide layer. In many

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centric genera, in Rhabdonema and other pennates [65, 140, 141] numerous silica scales are added to this layer as it is stretched by the expanding auxospore. Usually, the scales are flat structures with an annulus, like simplified centric valves [136]. However, Odontella possesses scales bearing dichotomously branching spines [162], which occur nowhere in the frustules of vegetative cells. In most radial centrics and possibly in Thalassiosirophycidae, this is all there is to the structure of the auxospore wall, and the auxospore itself is a simple expanding ball (see references in Table 2 of Medlin and Kaczmarska [116]) and the first (‘initial’) valves formed by the auxospore are hemispherical. The phylogenetic tree agrees with this observation. The formation of each valve is accompanied by a mitosis, but no cytokinesis occurs and one daughter nucleus degenerates (it appears to be a universal rule in diatoms, that no valve is produced without a preceding round of DNA replication and usually mitosis). After the first normal, cytokinetic mitotic division, the two daughter cells are heterovalvar, with oddly convex epivalves but more normal hypovalves. In most multipolar centrics (except Thalassiosirophycidae), a second layer is laid down by the auxospore, underneath the primary one. This layer, the properizonium, consists of a complicated system of silica bands and hoops ([137, 162], see references in table 2 of Medlin and Kaczmarska [116]), which force the auxospore to develop anisometrically, producing the characteristic shape of the future vegetative cell. Modification of shape can also occur during the formation of the initial valves, through contraction of the auxospore away from its silicified envelope (summarized by Round et al. [137]). This observation also agrees with the molecular tree. In pennates, the auxospore is at first usually globular or ellipsoidal (though there are exceptions, for example, [90, 96]) and is again enveloped in a primary organic wall. This wall ruptures when the auxospore starts to swell, the two more or less equal halves remaining as caps on the ends of the elongating cell (for example, [97]). As the cell expands, a primary silica ring is often formed surrounding the equator of the auxospore. Subsequently, more transverse bands are laid down around the auxospore, one after the other on both sides of the primary band, forcing expansion to occur solely along one axis, like the extension of a telescope (for example, [87, 88, 97, 162], see references in Table 2 of Medlin and Kaczmarska [116]). When elongation is complete, a

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second series of bands is often added beneath the first. These bands, unlike the first series, are orientated longitudinally. Together, the transverse and longitudinal series constitute the ‘perizonium’. Then, as in centric diatoms, the nucleus divides mitotically twice without cytokinesis, the two superfluous nuclei degenerating. After each mitosis a new theca is produced, to create the frustule of the newly enlarged cell. More rarely, silicified elements are apparently absent, as in Licmophora and some Eunotia [15, 101], the auxospore wall being largely or wholly organic. The properizonium and the perizonium are probably homologous, being different states of the same character (Kaczmarska et al. [64, 65] review the known distribution of these structures among diatoms). Our molecular tree indicates that this form of auxospore encasing came into being along the internode leading to the clade containing the pennates and the polytomy of multipolar centrics. The properizonium appeared at the origin of the multipolar centrics, whereas the slightly more differentiated form of covering present in pennates – the perizonium – (which is present in Rhabdonema and has been demonstrated recently in the araphid genus Gephyria by Sato et al. [140]) was acquired only once, apparently along the internode leading to the pennates. Again, however, we must point out that there is as yet no information on life cycles in the Asterionellopsis–Asterioplanus–Rhaphoneis–Talaroneis clade. Alone among ‘multipolar’ centrics, the Thalassiosirophycidae lack a properizonium in their auxospores and their auxospores expand isodiametrically. This must represent secondary loss or an ancestral polymorphism, because properizonia are present in other multipolar lineages, including Lithodesmium, Odontella and Chaetoceros, representing three of the lineages of multipolar centrics in Fig. 36a. Interestingly, a few Cyclotella (Thalassiosirophycidae) species are known which have oval or polygonal outlines (for example, [25, 122]) and it will be interesting to see whether they also have differentiated, properizonium-like auxospore coverings. PHYLOGENETIC SIGNAL IN CYTOPLASMIC ULTRASTRUCTURAL FEATURES

Three or possibly four arrangements of the Golgi apparatus can be distinguished in diatoms, relative to the endoplasmic reticulum (ER) and the mitochondria [115, 116]. The radial centric diatoms Coscinodiscus,

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Stephanopyxis and Ellerbeckia (the last is probably a close relative of Paralia, whose possibly basal position has been discussed above) have been found to possess Golgi bodies associated with a cisterna of ER and a mitochondrion. This association is called the G–ER–M unit in Schmid [142] and was correlated with the tree in Medlin et al. [115] and Medlin and Kaczmarska [116]. In the cortical cytoplasm, the mitochondrion is oriented towards the plasmalemma, whereas the Golgi apparatus is located on the other side of the ER. Near the nucleus, the situation is the reverse, with the mitochondrion on the side of the ER facing the nucleus whereas the Golgi apparatus faces the plasmalemma [142]. The G–ER–M association is also found in the Oomycetes and may, therefore, be an ancestral state inherited from the common ancestor of Oomycetes and Heterokontophyta (including diatoms). This feature is also found in red algae [142], however, and therefore Medlin and Kaczmarska [116] suggested that the feature was acquired from the red algal co-ancestor when it became an endosymbiont of the heterotrophic ancestor of at least all Heterokontophyta. Yet it remains difficult to understand how the endosymbiont’s endo-membrane system could have been transferred to the host and another consequence of a red algal origin of the G–ER–M unit in diatoms and Oomycetes is that the latter must have lost plastids secondarily. But see references in Medlin and Kaczmarska [116] regarding other organelles that have been retained in endosymbioses. In most multipolar centrics and in all pennates that have been examined with respect to Golgi location, the numerous Golgi bodies present are not dispersed through the cell in G–ER–M units but surround the nuclear envelope, thus forming a shell. In this perinuclear arrangement, the forming side of the Golgi bodies is oriented towards the nucleus [115]. There is no obvious association with mitochondria. The perinuclear arrangement is referred to as type 2 by Medlin et al. [115]. There are two modifications of type 2 Golgi arrangement. In one type (Golgi type 2.1), which has been found in Ulnaria (Synedra) ulna, the nuclear envelope, together with the nuclear matrix (with RNA in transit, but without DNA) forms two tentacles reaching the cell poles [143]. Golgi-bodies are aligned along one side of the tentacle, thus constituting the Plattenband (Platte = Golgi-body) sensu Geitler [143]. The other side of the tentacle possesses nuclear pores. This organization solves a distribution problem by providing a highway for products from and towards the nucleus in strongly elongated pennates [115]. Another

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modification of type 2 (type 2.2) is found in the raphid diatoms Pinnularia [24] and Caloneis [26], and in the araphid pennate Striatella [34], where there are many nuclear tentacles pointing out in various directions from the nucleus. Here, the tentacles are much thinner and thin out distally into ER cisternae. Pores are distributed all over the nuclear envelope, as usual, whereas the tentacles do not appear to possess pores. Golgi-bodies group in pairs alongside the tentacles, that is, they group in pairs on both sides of the tentacles, thus forming a series of what were originally called ‘Doppelplättchen.’ The G–ER–M arrangement is found primarily in radial centric diatoms, but not all radial centrics investigated possess this arrangement. Aulacoseira and its sister Melosira possess a perinuclear Golgi arrangement as in type 2, suggesting that this arrangement arose twice independently from the G–ER–M arrangement. Medlin and Kaczmarska [116] have also interpreted this as an ancestral polymorphism because both types of the Golgi are present in outgroups. Moreover, the G–ER– M arrangement is also observed in the bipolar centric diatom Odontella sinensis [126], which is definitely an advanced bipolar centric diatom phylogenetically embedded in a lineage where perinuclear Golgi-body arrangements are also present (see [142]); even the related O. aurita has a perinuclear arrangement. The occurrence of G–ER–M units in O. sinensis may be a reversal from type 2 to type 1, or an ancestral polymorphism. In Biddulphiopsis titiana, only the outer membrane of the nuclear envelope forms tentacles along which Golgi-bodies are paired up, whereas the remainder of the nuclear envelope is porose just as in group 1 [115]. In Medlin et al. [115] and Medlin and Kaczmarska [116], this arrangement is referred to as type 3. The arrangement in Biddulphiopsis might be an evolutionary step between types 1 and 2. In the phylogeny of Medlin et al. [115] this species forms, together with Lampriscus, the sister lineage of all other bipolar centric diatoms as well as the pennates. The evolutionary scenario for the cytoplasmic arrangement would then be as follows. In the common ancestor of all pennates and all bi- and multipolar centrics including Biddulphiopsis, the outer membrane of the nuclear envelope formed tentacles, with the Golgi-bodies becoming paired on either side of them. This state was then retained in Biddulphiopsis (and possibly in Lampriscus), whereas in all other bi- and multipolar centrics and their pennate ‘offspring’ the Golgi-bodies grouped

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around the nuclear envelope. The modifications of the group-2 arrangement then arose and are synapomorphies for particular taxa; here, both membranes of the nuclear envelope formed the tentacles and not only the outer membrane of the envelope. In fact, in terms of other aspects of its structure too, Biddulphiopsis exhibits characteristics that could be regarded as ancestral in bi- or multipolar centric diatoms. For example, its labiate processes are located in the valve mantle, like those of radial centric diatoms. In conclusion, the organization of ER, Golgi-bodies and mitochondria shows a pattern of states more or less in concordance with the topology of the molecular phylogeny. The G-ER-M unit is found in the Coscinodiscophytina and the perinuclear shell in the Bacillariophytina. Yet the patterns are not perfectly congruent and it is unlikely that Aulacoseira and Odontella are misplaced in the molecular trees, because frustule morphology also shows that Aulacoseira is a radial centric and Odontella is a bipolar centric diatom. The molecular tree also gives no support to the idea that the lineage containing Aulacoseira and Melosira might be sister to the bi- and multipolar centrics. Although in Medlin et al. [117] they are the last radial centric lineage to diverge. Possibly, the arrangement of Golgi bodies and other organelles is strongly influenced by relative and absolute nuclear and cytoplasmic volumes. It is noticeable that in pennate diatoms, the unusual Golgi arrangements (types 2.1 and 2.2) occur only in large-celled species. Therefore, perhaps in relatively small cells, such as those of many pennates and most Melosira and Aulacoseira, there may be enough space around the nucleus and the cell volume may be sufficiently small overall so that products from perinuclear Golgi bodies can be produced and disseminated rapidly through the cell to the location where they are required. Evolution of G– ER–M units could be a compensating adaptation to large cell size and highly vacuolate cytoplasm (this latter feature helping to produce low density cells: many of the radial centrics studied thus far are planktonic). This idea can be probed further by examination of radial centric diatoms with different cell shapes and sizes. Alternatively, the cells with a G-ERM unit could be involved in certain processes that require a high amount of energy, thus the close juxtaposition of the mitochondria with the Golgi. Plastid shape and arrangement vary greatly among diatoms. Generally, radial centrics have many small chloroplasts, as do many multipolar centrics and some araphid pennates. Raphid pennates, on the

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other hand, usually have a few large chloroplasts, whose shape and position in the cell is often genus or family-specific [94]. Recent work on pyrenoid ultrastructure by Schmid [144] on species drawn (albeit sparsely) from across the diversity of diatoms indicates some trends in concordance with the molecular phylogeny. Plastids of radial, bi- and multipolar centric diatoms possess single, membrane bound pyrenoids that are traversed by one or more sets of lamellae not connected to thylakoids (see references in Medlin and Kaczmarska [116]). Notably, these species generally possess multiple small chloroplasts in their cells. The most conspicuous difference between radial centrics and bi- and multipolar centrics assessed by Medlin and Kaczmarska [116] from the review by Schmid [144], is that the chloroplasts of the latter tend to possess multiple pyrenoids. The pennates possess pyrenoids that are either embedded or protruding from the chloroplast and there is a wide variety in pyrenoid morphology [94, 144]. PALAEONTOLOGY AND PHYLOGENY

It is generally accepted that gene trees, if carefully constructed and analysed, can allow us to reconstruct aspects of the past from information available from living organisms. However, many lineages are no longer represented among extant diatoms, and fossils offer the only direct method of checking phylogenetic hypotheses. Diatoms possess an excellent fossil record, from which much can be deduced concerning their evolutionary history [5, 40, 166]. Nevertheless, interpretation of fossils must always be done in the context of the more complete knowledge which can be acquired by relating to living forms. Similarities between fossils and extant forms may result from a shared evolutionary history, but could also reflect convergence (for example, [37]). If aspects of morphology show great phenotypic plasticity or homoplasy among living diatoms, we must show great caution in assuming they will be of any greater taxonomic value when comparing extant and fossil taxa. Many fossil diatom taxa are available in large quantities from facies all over the world, especially during the Tertiary and to a lesser extent, the Upper Cretaceous Period (for example, [49, 165, 165, 166]). The first appearances of morphological synapomorphies or genera, as deduced from molecular phylogenies, can thus be dated using the fossil record (for example, [73, 112, 113, 115]). However, fossils become rare or difficult to interpret in pre-Cretaceous facies. Reports of diatoms in strata older

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than the Jurassic (for example, [80, 149, 150]) are not generally accepted, either because the material is too fragmentary to allow confident interpretation, or because contamination cannot be ruled out. The diatoms reported in these very early deposits look identical to modern ones, which would contrast greatly with the unusual morphologies found in the deposits studied by Gersonde and Harwood [40]. Round and Crawford [36] suggested that the early fossil record of diatoms has been lost because of diagenesis, and that the origin of the diatoms might be as far back as the Precambrian era. However, this does not agree with molecular data, which indicate an origin c. 250 Ma, possibly around the time of the Permian–Triassic mass extinction [73, 113]. The first plausible reports of diatoms are currently from the Jurassic and are of small helmet- or thimble-like valves [52, 134]. The most ancient deposits in which well-preserved diatoms have been recovered in large quantities are from c. 120 Ma Lower Cretaceous sediments around Antarctica [40, 50]. The survival of oceanic sediment of this age is very unusual; most of the oceanic crust that existed in the Lower Cretaceous Period has been destroyed by subduction or compression through tectonics or overlying volcanic deposits. All of the Lower Cretaceous Antarctic fossils resemble modern diatoms in their overall organization (i.e. they have valves and girdle) and an annulus is visible in several cases (for example, [40], pl. 14, Fig. 3), indicating a centric type of organization of the valve pattern. Some of the fossils possess interlocking ornamentations emanating from the valve perimeter, whereas a few have centrally located simple tubes. Generally, the valves are circular, but Bilingua has bipolar valves (but a radial pattern of striae) and resembles modern multipolar diatoms. These fossils could perhaps be interpreted, therefore, as members of the same lineages as some of the modern radial and multipolar centric diatom genera. If so, it would imply that the primary radiation that produced the various lineages of radial centric diatoms, and also the first multipolar diatoms, occurred in or before the Lower Cretaceous Period. What is questionable, however, is that none of the ancient fossil ‘radial’ centrics possess peripheral labiate processes, whereas most extant radial centric ones do. Some of the Lower Cretaceous diatoms, like Rhynchopyxis and Praethalassiosiropsis [40], have a central tube-like structure which, as in some modern multipolar centrics, appears to lie within the annulus. However, although the tube-like process of

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Rhynchopyxis superficially resembles the labiate process found in a similar position in extant Ditylum, it opens internally by a round pore, with no trace of tightly appressed lips (for example, [51]). The central process of Praethalassiosiropsis does not resemble anything found in modern diatoms, since it opens internally by a scatter of small round pores, as in a pepper pot. Indeed, none of the Lower Cretaceous fossils have anything that is easily interpretable as a labiate process. Furthermore, there is no sign of any specialized pore fields for mucilage secretion. Of course, one could invoke independent and sudden acquisition of labiate processes and pore fields in all existing diatom lineages during the mid or late Cretaceous Period but such an assumption is not particularly parsimonious. In Medlin and Kaczmarska [116], a scenario in which a central process, such as that found in Praethalassiosiropsis, could evolve into a modern labiate process was proposed. We must therefore be cautious in interpreting the Lower Cretaceous flora. Overall shape and the broad lay-out of the striation pattern and processes may be poor guides to relationships. Molecular phylogenies have already revealed many cases of parallel evolution: for example, Fig. 36c illustrates clearly, even without formal parsimony analysis, that a wedge-shaped cell shape, with heteropolar valves, has arisen independently in at least three lineages of raphid diatoms: Gomphonema (related to the isopolar diatoms Cymbella, Anomoeoneis and Cocconeis), Pseudogomphonema (related to the isopolar diatoms Navicula, Seminavis and Pleurosigma) and Actinella (related to the isopolar diatoms Eunotia and Eunophora). Inspection of Fig. 36 will reveal many similar examples, involving independent evolution of circular valves, bilaterally asymmetrical valves, apical pore fields, etc. Such character states may have appeared and then been lost over and over again. Consequently, there must be detailed correspondence in many respects between fossil and modern genera, and there must be plausible demonstration that these resemblances are synapomorphies, before we can conclude that a close phylogenetic relationship exists between them. Deceptive parallelism among fossils, or between fossils and living taxa, has been encountered in other groups of organisms, such as ammonites [120], foraminifera [10] and calcified macroalgae [74]. If the Lower Cretaceous diatoms are not closely related to any modern diatoms, it becomes even more difficult to determine when key evolutionary events took place. Heterokontophyte phylogenies do not

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reveal long branches (= comparable to extended time, assuming a stochastically constant rate of sequence change) between the separation of the bolidophycean lineage – so far as is known, these are the nearest living relatives of the diatoms – and the basal ramification of the extant diatom diversity [47]. Nevertheless, an internode under thybasal ramification of the extant diversity implies that there is some time for an older diversification and the Antarctic Lower Cretaceous flora may be composed predominantly, or wholly, of representatives of these earlier, now extinct lineages. Medlin and Kaczmarska [116] have proposed links between certain groups of these taxa and modern taxa. In this case, and if Medlin and Kaczmarska [116] are correct in asserting that there is a basal dichotomy among extant diatoms between radial centrics and multipolar centrics + pennates (rather than the paraphyly of Fig. 36a), it could be suggested that all extant diatoms have evolved from just one lineage of radial centric, diatoms with a simple central (labiate) process. In the modern radial centrics the process was replicated many times and moved to the periphery in many cases (but not in all: clusters of central processes are present in for example, Stellarima), whereas in the other clade, cells acquired the capacity to produce mucilage pads via specialized areas of pores, but the labiate processes generally remained central. Alternatively, there might have been two lineages instead of one, one with a central process and one without, evolving into the Bacillariophytina and the Coscinodiscophytina, respectively. As a third alternative, all extant diversity may have evolved from lineages so rare that they have not yet been found in Lower Cretaceous oceanic sediments, or from lineages that were confined to habitats adverse to fossilization, for example, high-energy inshore environments or even freshwater or terrestrial habitats, as suggested by Harwood et al. [52]. The marvellously preserved diverse fossils observed by Gersonde and Harwood [40] may have been the predominant diatoms at the time, at least in the Cretaceous oceans surrounding this part of Antarctica, but for whatever reason, their offspring apparently did not survive to the present. Unfortunately, there are some large gaps in the diatom fossil record. These gaps could simply indicate a hiatus in fossilization in lineages that were nevertheless present throughout, but they could also represent extinction events and the subsequent evolution of superficially similar but unrelated taxa. One particularly frustrating gap occurs just after the occurrence of the exceptionally well-preserved flora from the Lower

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Cretaceous system described above, during which there is a period which diatoms are poorly preserved and often pyritized. Well-preserved fossil diatoms then reappear at ca. 90 Ma, in the mid Cretaceous Period. By the Upper Cretaceous, many of the diatoms prevalent at that period are so similar to extant genera (for example, [164, 165]) that there seems no question that they belong to the same clades. There is apparently no major extinction of diatom lineages at the K/T boundary (65 Ma), but perhaps it is no coincidence that the explosive radiation of pennates, maybe including the evolution of raphid diatoms, seems to have occurred at around this time. Pennate diatoms have never been observed with certainty in Lower Cretaceous facies. They appear only after 90 Ma in the fossil record and raphid diatoms are not known before the Palaeocene [165], which ended 55 My ago. Molecular evidence for Toxarium suggests that pennate like morphology and behaviour have evolved twice independently and the fossil record may be riddled with many more such ‘experimental’ designs. However, the earliest confirmed reports of pennates show diatoms with a simple sternum (for example, [164]), not a bifacial annulus. There is a recent report [11] of a freshwater diatom flora, including pennate diatoms, from 70 My-old deposits in Mexico. The material (preserved in chert) does not allow confident identification of the lineages represented, but if the organisms are confirmed to be both diatoms and autochthonous, the diversification of the pennates may have occurred earlier than is currently believed. The pennate specimens in this deposit resemble modern species, which in the molecular tree diverge very late. Therefore, if these diatoms are not contaminants, then they represent an earlier extinct lineage of pennate diatoms and the similarities in morphology represent parallel evolution. Overall, however, the order in which the major groups arose (as inferred from molecular phylogenies i.e. centrics first, then araphid pennates, followed by raphids), agrees with the stratigraphic evidence of the fossil record. NEW DIRECTIONS IN DIATOM PHYLOGENY

It is expected that sequence and structural information provided by the draft genome of the diatom Thalassiosira pseudonana [3] will play an important role in the development of new markers for phylogenetic analyses. For instance, the use of low, ideally single copy gene families can be explored for constructing multigene phylogenies, which may enhance

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the resolution of the deeper branches in the phylogeny of diatoms. Analysis of the evolution of functional gene families and structural aspects of diatom genomes may lead to a better understanding of the processes that gave rise to the major clades within the diatoms. Recently, a major new thrust of phylogenetic research into diatoms has begun to emerge that focuses not on the ‘big picture’ of macroevolutionary trends, but on the fringe between what should be regarded as a single species and what should not. Of course, determining the limits of particular species of diatoms is only a part of the much wider debate about what constitutes ‘a species’ and what species concepts could be applied. Traditionally, diatom species have been defined somewhat arbitrarily on the basis of morphology, with no reference to other criteria, such as genetic similarity or reproductive isolation [95]. In the 1980s, studies of a few ‘model species’ showed a lack of congruence between morphologically defined taxa and reproductive isolation, indicating that there were many more species than had previously been recognized (for example, [92, 95]). The introduction of molecular genetic methods has confirmed this conclusion and greatly extended our capacity to detect and study the subtleties of microevolution in diatoms. To date, most molecular genetic studies have been of marine phytoplankton and, although relatively few have been published thus far, almost all concur in suggesting that ‘morphological’ plankton species are heterogeneous, consisting of monophyletic or paraphyletic assemblages of sibling species (174, 115, 139). So far, results suggest that sibling species within a ‘morphological species’ are ‘pseudo-cryptic’, i.e. that they do show stable morphological differences, but that these differences are so subtle they were previously overlooked or interpreted as ecophenotypic variations. If confirmed across a wider spectrum of phytoplankton diversity, these exciting new findings would have wide-ranging implications for our concepts of phytoplankton taxonomy, ecology and evolution. An example of how new insights are changing our views of what defines a species is shown in Skeletonema costatum (sensu lato). This taxon was believed to be a morphologically plastic, ecologically versatile and ubiquitous species occurring in almost every marine and brackish water habitat imaginable, except at very high latitudes in the Arctic and Antarctic [57]. Detailed DNA sequence comparisons, using the nuclear 18S and 28S rDNA regions, have revealed, however, that this taxon

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%$consists of several genetically distinct species [109, 139]. Subsequent careful examination of the strains has revealed subtle morphological differences among the molecular clades and Sarno et al. [139] have, therefore, described several new taxa (S. dohrnii, S. grethae, S. japonicum, S. marinoi). The newly described species formerly included in S. costatum do not even form a single clade, because some among them are more closely related to other previously recognized species in Skeletonema (S. tropicum, S. menzelii, S. subsalsum). Unlike S. costatum (sensu lato), the new Skeletonema taxa appear to have limited distributions. Skeletonema grethae appears to be found only in the North-western Atlantic, whereas S. japonicum seems to abound in the temperate zones except Europe where some species seem to be pantropical, whereas others appear to be confined to the temperate zones (Kooistra et al., unpublished). The closely related species S. marinoi and S. dohrnii are both abundant in the Mediterranean Sea in winter but they have not been found together: Skeletonema marinoi forms major blooms in the northern Adriatic whereas S. dohrnii is found in the Gulf of Naples (Tyrrhenian Sea), on the opposite side of the Italian peninsula [139]. There are several other examples where molecular phylogenetic investigations of marine planktonic diatoms have shown that species need to be re-defined. In the most recent review of the genus Thalassionema, Hasle [55] recognized five morphologically distinct species: T. bacillare, T. frauenfeldii, T. nitzschioides, T. pseudonitzschioides and T. synedriforme. However, subsequent molecular and morphological research into the diversity of this genus has already revealed at least 13 genetically distinct species. In this case, unlike Skeletonema, many of the new species are truly cryptic, being morphologically apparently indistinguishable, even in SEM (Sarno and Kooistra, unpublished results). In Rhizosolenia setigera, 18S rDNA sequences show considerable divergence among strains [53], but examination of type material of this species shows that none of the isolates sequenced correspond to the type and so none can properly be called R. setigera. The genus Pseudo-nitzschia is responsible for harmful algal blooms worldwide because of its capacity to produce the potent neurotoxin domoic acid. Many genetically and morphologically distinct species have recently been described and many more are being discovered (for example, [84, 85, 86]). But that is not the whole picture; many of these so-called species appear to consist of assemblages of cryptic or semi-cryptic species [85, 86, 123]). Similar

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diversity has also been found in Fragilaria islandica, Grammatophora oceanica, Phaeodactylum tricornutum, Cyclotella spp., Eucampia antarctica and Melosira varians (Medlin, Jung, Bezsteri, Sato, unpublished). For freshwater diatoms, and for benthic diatoms of all kinds, there is much less information, although these represent by far the majority of diatom species [95]. However, existing information shows the same pattern as in marine phytoplankton: traditional morphologically defined taxa consist of complexes of cryptic and semicryptic species. Within the Sellaphora pupula species complex, molecular data have revealed unsuspected diversity, even within clades already studied using reproductive, ultrastructural and morphometric data [100, 6] and it is likely that what was originally considered a single species will end up being split into many tens or hundreds of species [6, 103]. Members of this clade have been evolving separately for so long in some cases that rapidly evolving parts of the genome, for example, ITS rDNA, which are often used for species-level studies in other groups of organisms, are of limited value here. One remarkable feature of the S. pupula complex is that up to ten or more members of the complex often coexist in the epipelon of what appear to be more or less homogeneous silty sediments, raising interesting questions about niche separation in diatoms. Those found together in the same locality are not always those that are most closely related phylogenetically; a wide geographical range is likely in two cases (S. capitata and the ‘pseudocapitate’ deme; [6, 95, 103). Several other morphologically defined benthic ‘species’ have also been shown, by studies of reproductive isolation and/or molecular data, to be complexes of semicryptic or cryptic sibling species. These include the marine Achnanthes brevipes and the freshwater Neidium ampliatum and Eunotia bilunaris ([95] and unpublished rbcL data; [99] Vanormelingen, unpublished data). Hence, pioneering studies in diatom species diversity strongly support the hypothesis that the actual number of diatom species of ca. 10,000 usually cited (for example, [157]) is actually more in the order of 100,000 [95, 98, 100]. Many published claims about the ecology or biogeography of particular species must be treated with great circumspection, because they refer not to a single species, but to a species complex and perhaps not even to a monophyletic one. The claim that some species are ‘globally distributed’ needs to be tested. Perhaps there are no truly cosmopolitan diatoms and the idea that some species have

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high ecological plasticity may be an artifact of poor taxonomy. Instead, suites of sibling species may be common, each adapted to a tightly defined ecological niche and with a restricted horizontal, vertical or temporal distribution. Acknowledgements

Wiebe Kooistra thanks Carmen Minucci for assistance with culture maintenance, Gandi Forlani for help with EM, Elio Biffali and the molecular service of the SZN for sequencing. Koen Sabbe and Victor Chepurnov thank Sylvie Cousin for sequencing and Ghent University for financial support (BOF-GOA 12050398). David Mann thanks Gillian Simpson, Michelle Hollingsworth, Alex Ponge and Michael Möller for help with molecular phylogenetics. References [1] Anonymous. Two letters from a gentleman in the country, relating to Mr. Leeuwenhoek’s letter in Transaction, no. 283. Communicated by Mr. C. Phil, Trans Roy Soc London 1703; 23: 1494–1501. [2] Anonymous 2. Proposal for a standardization of diatom terminology and diagnoses. Nova Hedw. Beih. 1975; 53: 323–354. [3] Armbrust EV, Berges JA, Bowler C, et al. The genome of the diatom Thalassiosira pseudonana: Ecology, evolution, and metabolism. Science 2004; 306: 79–86. [4] Baldauf JG. Middle Eocene through early Miocene diatom floral turnover. In: Prothero DR, Berggren WA, eds. Eocene-Oligocene climatic and biotic evolution. Princeton USA: Princeton University Press, 1992; 310–326. [5] Barron JA, Mahood AD. Exceptionally well-preserved early Oligocene diatoms from glacial sediments of Prydz Bay, East Antarctica. Micropaleontol 1993; 39: 29–45. [6] Behnke A, Friedl T, Chepurnov VA, Mann DG. Reproductive compatibility and rDNA sequence analyses in the Sellaphora pupula species complex (Bacillariophyta). J Phycol 2004; 40: 193–208. [7] Bhattacharya D, Medlin LK. Dating algal origin using molecular clock methods. Protist 2004; 155: 9–10. [8] Boyd PW, Watson AJ, Law CS, et al. A mesoscale phytoplankton bloom in the polar Southern Ocean stimulated by iron fertilization. Nature 2000; 407: 695–702. [9] Boyle JA, Pickett-Heaps JD, Czarnecki DB. Valve morphogeneis in the pennate diatom Achnanthes coarctata. J Phycol 1984; 20: 563–573. [10] Cifelli R. Radiation of Cenozoic planktonic Foraminifera. Syst Zool 1969; 18: 154– 168. [11] Chacón-Baca E, Beraldi-Campesi H, Cevallos-Ferriz SRS, et al. 70 Ma nonmarine diatoms from northern Mexico. Geology 2002; 30: 279–281.

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[159] Stosch HA von. Kann die oogame Araphidee Rhabdonema adriaticum als Bindeglied zwischen den beiden grossen Diatomeengruppen angesehen werden? Berdt Bot Ges 1958; 71: 241–249. [160] Stosch HA von. Manipulierung der Zellgrösse von Diatomeen im Experiment. Phycologia 1965; 5: 21–44. [161] Stosch HA von. Structural and histochemical observations on the organic layers of the diatom cell wall. In: Ross R, ed. Proceedings of the 6th Symposium on Recent and Fossil Diatoms. Koenigstein, Germany: Koeltz, 1981; 231–52. [162] Stosch HA von. On auxospore envelopes in diatoms. Bacillaria 1982; 5:127–156. [163] Stosch HA von. Some marine diatoms from the Australian region, especially from Port Phillip Bay and tropical north-eastern Australia. II. Survey of the genus Palmeria and of the family Lithodesmiaceae including the new genus Lithodesmioides. Brunonia 1987; 9: 29–87. [164] Strel’nikova NI. Diatomei pozdego mela (Zapadnaya Sibir’). ‘Nauka’: Moscow, USSR, 1974. [165] Strel’nikova NI. Paleogenovye diatomovye vodorosli. St Petersburg, Russian Republic: Saint Petersburg University, 1992. [166] Strel’nikova NI. Evolution of the family Coscinodiscaceae. In: John, J. ed. Proceedings of the 15th International Diatom Symposium, Perth 1998. Koenigstein, Germany: Koeltz, 2002; 381–394. [167] Vyverman W, Sabbe K, Mann DG, et al. Eunophora gen. nov. (Bacillariophyta) from Tasmanian highland lakes: description and comparison with Eunotia and amphoroid diatoms. Europ J Phycol 1998; 33: 95–111. [168] Walsby AE, Xypolyta A. The form resistance of chitan fibres attached to the cells of Thalassiosira fluviatilis Hustedt. Br Phycol J 1977; 12: 215–223. [169] Williams DM. Comparative morphology of some species of Synedra Ehrenberg with a new definition of the genus. Diat Res 1986; 1: 131–152. [170] Williams DM. Cladistic analysis of some fresh-water araphid diatoms (Bacillariophyta) with special reference to Diatoma and Meridion. Plant Syst Evol 1990; 171: 89–97. [171] Williams DM, Round FE. Revision of the genus Synedra Ehrenb. Diat Res 1986; 1: 313–339. [172] Williams DM, Round FE. Revision of the genus Fragilaria. Diat Res 1987; 2: 267–288. [173] Williams DM, Round FE. Notes. Fragilariforma, nom. nov., a new generic name for Neofragilaria Williams & Round. Diat Res 1988; 3: 265–267. [174] Zechman FW, Zimmer EA, Theriot EC. Use of ribosomal DNA internal transcribed spacers for phylogenetic studies in diatoms. J Phycol 1994; 30: 507–512.

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Ascomycota: Introduction to Biodiversity, Evolutionary Genomics and Systematics WOLFGANG SCHWEIGKOFLER Department of Plant Protection, Research Center for Agriculture and Forestry. Laimburg, Italy

ABSTRACT The Ascomycota form a monophyletic division of the true fungi with more than 70,000 described and probably several 100,000s non-described unicellular, dimorphic and filamentous species, which are important saprophytes, symbionts or parasites of plants and animals worldwide. Whereas traditional systematics based on morphological characters fail to integrate asexual groups, considerable progress towards a unified phylogenetic system was achieved during the last two decades using molecular sequence analysis, especially of the nuclear ribosomal DNA unit. In order to resolve deep (for example ordeal) relationships among the Ascomycota and to improve bootstrap support, single gene phylogenies are more and more replaced by multi-gene phylogenies. This strategy uses protein-coding and non-coding single-copy and multi-copy gene regions, for example, the 18S and 26S rRNA genes, the ITS1 and 2 regions, the mtSSU rDNA, RNA polymerase I subunit B (RPB1), RNA polymerase II subunit B (RPB2), elongation factor (EF)-1a, >-tubulin gene, and others. Fungal genomes are relatively compact, ranging from approx. 8 Mb to 42 Mb in length. The yeast Saccharomyces cerevisiae was the first eukaryote, for which the whole genome sequence was published. The recent completion of genomes sequences from members of all three main ascomycetous lineages (Hemiascomycetes, Archiascomycetes, and Euascomycetes) opens up the way for comparative genomic analysis. Key Words: Ascomycota, fungi, biodiversity, phylogenetics, genomics

Address for correspondence: Department of Plant Protection, Research Center for Agriculture and Forestry Laimburg, I-39040 Auer (BZ), Italy. Tel: ++ 39 0471 969630, Fax: ++39 0471 969599, E-mail: [email protected].

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Abbreviations: bp: base pair; EAS: Extracellular Amyloid Substances; GCPSR: Genealogical Concordance Phylogenetic Species Recognition; IGS: Intergenic Spacer Region; ITS: Internal Transcribed Spacer; kb: kilobase; Ma: million year; Mb: Megabase; mtDNA: mitochondrial DNA; ORF: Open Reading Frame; RIP: Repeat Induced Point Mutation; RPB1: RNA polymerase I subunit B; RPB2: RNA polymerase II subunit B; RST: Random Sequence Tag; SSU rDNA: small subunit ribosomal DNA; YLS: Yeast-like Endosymbionts INTRODUCTION

The Ascomycota (or sac fungi) are a group of the true fungi defined by the presence of an ascus, the typical “sac-like” cell, in which ascospores (meiospores) are produced by “free cell formation” after karyogamy and meiosis [74]. Based on molecular analyses of the small subunit ribosomal RNA gene (SSU rDNA or 18S rDNA) the true fungi constitute a monophyletic group [160], which is more closely related to the animals than to the plants, where they have been placed traditionally in ”premolecular” times. Phylogenetic analysis of several protein-coding genes (elongation factor, alpha- and beta tubulin, actin, dihydrofolate reductase, thymidylate synthase) support this view [8, 31, 45, 195, 225]. Fungi, together with the Animalia and the Choanozoa, are now considered to form the monophyletic group of the “ophistokonts”, named after the locomotion in unicellular stages [195]. Phylogenetic analysis of a particular gene fusion involving dihydrofolate reductase (DHFR) and thymidylate kinase (TS) that is found only among some eukaryotes, suggests that the “ophistokonts” separate very near the basic root in the eukaryotic tree [195]. Fungal-like organisms belonging to the Oomycota, Hyphochytridya, Labyrinthuloida and Thraustochytrida with simple hyphal morphology, have been shown to form a distinct clade within the Stramenopila and are not closely related to the true fungi. Due to historical reasons, this group, which contains many important plant pathogens (especially in the genera Pythium, Phytophthora, Peronospora and Plasmopora) is still widely referred to as “fungi” by many researchers in the applied fields. Besides the Ascomycota, the true fungi also include the Chytridiomycota (the only fungal group with flagellate cells), the Zygomycota and the Basidiomycota. These four fungal lineages share an exclusively absorptive or lysotroph nutrition [236], chitinous cell walls

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[12, 13], the =-aminoadipic acid lysine biosynthetic pathway [221, 222], glycogen as storage carbohydrate [36] and plate-like mitochondrial cristae [36]. The Microsporidia, traditionally regarded as primitive members of the eukaryotic crown groups, cluster within the true fungi based on molecular analyses of the gene for the largest subunit of the RNA Polymerase II (RBP1) [78], and are now considered to be a highly specialized derived group, which lost many typical fungal characters due to an extreme adaptation to an obligate endoparasitic life style. The Chytridiomycota are often regarded as the most primitive true fungi, because they are zoosporic (monoflagellate or more rarely polyflagellate) and have different modes of sexual reproduction (isogamy, anisogamy, oogamy, gametangiogamy and somatogamy) [1]. The approx. 800 described species in five orders are mainly aquatic saprobes or parasites, members of the Neocallimastigales are obligate anaerobes in the rumen of herbivores. Molecular phylogenetic analyses [133, 162, 200, 209] suggest that the Chytridiomycota are not monophyletic; the relationship between the main chytridiomycetous lineages and the evolution towards the other fungi and the presumably polyphyletic losses of the flagella need further studies. The Zygomycota, a group consisting of circa 1,000 species, contains many ecologically important mycorrhiza fungi as well as saprobes and obligate biotrophic parasites of insects. Traditionally they have been characterized morphologically on the basis of mainly nonseptate hyphae and the production of zygospores (resting spores), formed by fusion of similar gametangia. Molecular analyses showed that they do not form a monophyletic group, but that the arbuscular mycorrhiza forming Glomales and the Endogonales-Mortierellales may lie outside the main Zygomycota group, either more basal to the AscomycotaBasidiomycota [32] or closely related to the Chytridiomycota [45]. The Basidiomycota are a monophyletic group characterized by the presence of a basidium, from which after karyogamy and meiosis, haploid basidiospores are produced externally [74]. Hawksworth et al. [74] accepted 41 orders with approx. 23,000 species. Based on distinct cell wall pattern [153-155] and sequence analyses of the 18S rRNA and 26Sr RNA genes [17, 162, 182, 206, 207] three classes can be distinguished: the Ustilaginomycetes (“smut fungi”), the Urediniomycetes (“rust fungi”) and the Hymenomycetes. Whereas the Ustilaginomycetes and Urediniomycetes comprise mainly plantpathogenic species, the Hymenomycetes comprise saprophytic, mycorrhizal and wood-decay fungi. Basidiomycota are typically mycelial and many Hymenomycetes

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develop complex fruiting bodies (mushrooms), but yeasts and yeaststages are known from all three classes. Fossil Record of the First Fungi

Based on molecular clock analyses of multiple protein sequences, the main fungal lineages diverged during the proterozoic age ca. 1,000 Ma ago in marine habitats [75]. The first fossilized fungal structures associated with early terrestrial plants have been detected from the Ordovician (circa 460 Ma) and strongly resemble modern arbuscular mycorrhizal fungi of the Glomales [165]. Ascomycota-like fossils associated with microarthropods have been described from the Silurian (438-408 Ma) [189]. The first lichens have been documented for the Devonian [215], but are speculated to have been present already in the Precambrian [169]. Fungal Biodiversity

Together with the Insecta, the fungi are probably the most diverse and species-rich group of organisms. About 70,000 species of true fungi have been described so far, of which ca. 33,000 belong to the Ascomycota and 15,000 to the mitosporic fungi (or Deuteromycota) with mainly ascomycetous affiliation [74]. Estimation of the total number of species differs widely, partly due to different circumscription of species and the method used for calculating bio-diversity. The existence of up to 1.5 million fungal species is predicted by some authors [73]. Most ascomycetous taxa are known from terrestrial habitats, with only relatively few groups from aquatic or marine habitats. Saprophytic soil fungi and plant-associated fungi (being parasitic, endophytic, lichenized) are ubiquitous and species rich, whereas parasites of vertebrates are known in relatively few groups (for example, Onygenales, Saccharomycetales, and Eurotiales). Associations of Ascomycetes with insects are numerous and diverse: invasive pathogens, which are able to penetrate the host, multiply within the hemocoel and kill the insect are known mainly from the Clavicipitaceae with more than 700 species mainly from the tropics (for example, Cordyceps, Beauveria, and Metarhizium), but occur also in other ascomycetous lineages. The Laboulbeniales (ca. 2,000 species) are-with a few coprophile exceptionsobligate ectoparasites of insects and some other arthropod groups (Arachnida and Diplopoda). The whole fungal life cycle is completed on

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the insect cuticle without killing the host. Insect guts are inhabited by numerous hemiascomycetous and a few unicellular euascomycetous species [159, 202]. Insects also play a major role as vectors of spores of plant-pathogenic fungi, the best known examples being Ophiostoma ulmi and O. novo-ulmi (Ophiostomatales), the causal agents of Dutch Elm Disease, which are associated with the bark beetle Ips spp. (Coleoptera:Scolytidae). Lichenized fungi comprise ca. 13–18,000 species of ascomycetous fungi belonging to 13 orders, of which only four orders are exclusively lichenized [74]. Smaller numbers of lichenized fungi are also found within the Basidiomycota (for example, Dictyonema, Multiclavula, and Omphalina). Lichens originated in several independent lineages, as has been shown by Gargas et al. [57] using sequence analyses of the 18S rRNA gene. Most lichenized Ascomycetes produce asci in ± open apothecia and have been grouped within the Discomycetes (Table 1). The extensive detection of the “missing fungal species” is a challenging goal for further research on biodiversity. Numerous new species can be expected from soils, as endophytes in plants (both temperate and tropical) and associated with insects. For example, the group of Blackwell at the University of Louisiana recently isolated a surprising variety of new hemiascomycetous yeasts species from basidiocarp-inhabiting beetles. An unexpected diversity of major new ascomycetous lineages was recently detected from alpine soil samples from Colorado [178]. Anamorph, Teleomorph, Holomorph

Many ascomycetous fungi are pleomorph and produce states which are characterized by sexual (meiospores=ascospores) or asexual (mitospores=conidia) spores. Following the concept proposed by Hennebert and Weresub [76], the term teleomorph is used for the sexual (or “perfect”) state of a fungus, and anamorph for the asexual (or “imperfect”) state. The whole fungus, with all different states produced during its life cycle, is called holomorph and includes a teleomorph and often one, or rarely more anamorphs [74]. In some cases, a teleomorph is associated with more than one anamorphs, which then are called synanamorphs [56]. According to the International Code of Botanical Nomenclature [65], the name applied to the perfect state should also cover that of the imperfect state. Unfortunately, parallel names for both

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states are still widely in use and cause considerable taxonomic confusion (or ambiguity). The form group of the Deuteromycetes (also called Deuteromycota, Fungi imperfecti or mitosporic fungi) is characterized by the lack of any meiotic state, and therefore, has never been integrated into the classic fungal system (Table 1). For practical reasons, an artificial classification was introduced which does not reflect phylogenetic relationships [74]. According to this system, the Hyphomycetes comprise species which do not produce conidia (mycelia sterilia, “Agonomycetales”) or produce conidia on separate conidiophores (“Hyphomycetales”), on conidiophores which are aggregated as synnemata (“Stilbellales”) or on conidiophores developing from sporodochial conidiomata (“Tuberculariales”). The Coelomycetes are a form group of mitosporic fungi, in which conidia are formed within a cavity lined by fungal or fungal/host tissue [74]. Based on the morphology of the conidiomatal structure three form taxa are distinguished: the “Sphaeopsidales” producing pycnidia, the “Melanconiales” producing acervuli, and the “Pycnothyriales” producing pycnothyria. Using molecular analyses, mainly of the rRNA genes, the relationship of many purely mitotic or sterile taxa to ascomycetous or -to a lesser extent-basidiomycetous groups, has been unveiled unambiguously during the last 20 years. Interestingly, clusters of meiotic and strictly mitotic taxa were detected in several orders indicating that multiple independent losses of the teleomorphic state have occurred. Examples are the anamorphic genera Fusarium (Teleomorph T: Gibberella, Nectria; order Hypocreales), Trichoderma (T: Nectria, Hypocrea; order Hypocreales), Penicillium (T: Talaromyces, Eupenicillium; order Eurotiales), Aspergillus (T: Emericella, Eurotium; order Eurotiales), Botrytis (T: Botryotinia; order Helotiales) and Leptographium (T: Ophiostoma; order Ophiostomatales). The adaptive advantage of a non-sexual versus a sexual life cycle of pathogens is still a matter of debate. The very rare production of sexual stages in groups with predominantly asexual reproduction was shown for example, for the entomopathogen Cordyceps brongniartii (anamorph: Beauveria brongniartii) [186] and the forest pathogen Ophiostoma wageneri (anamorph: Leptographium wageneri var. ponderosum) [63]. Within these species sexual recombination is obviously still possible, but seems to play only a very restricted role in the life cycle. It cannot be excluded that teleomorphic stages of several taxa, of which at present only mitosporic

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stages are known, will be discovered in several groups, but nevertheless many apparently clonal fungal groups show a considerable ecological fitness. Still, the fact that no mitosporic ascomycetous fungus has passed strict population genetic tests for clonality [59] and the detection of mating-type loci in the asexual yeast Candida albicans [82] indicate a history of “hidden” recombination. Nonmeiotic genetic recombination by a parasexual cycle has been found in several filamentous mitosporic groups in vitro (for example, Aspergillus niger), but has not been demonstrated in nature on any fungus [59]. According to Hawksworth et al. [74], parasexuality is defined by: (1) the production of diploid nuclei in a heterokaryotic haploid mycelium (2) the multiplication of the diploid nuclei along with haploid nuclei in a heterokaryotic mycelium (3) the sorting out of a diploid homokaryon (4) segregation and recombination by crossing-over at mitosis (5) haploidization of the diploid nuclei The resulting genetic recombination is similar to that achieved by meiosis [74]. Species Concepts Used in Fungal Taxonomy

The species concept is central to biology in general, and to the estimation of biodiversity and phylogenetics in special. Morphological and Biological Species Concepts (MSC, BSC) have been traditionally used for circumscription of species and have been reviewed in detail [70, 214]. Phylogenetic species concepts (PSC) became more important since the general availability of molecular sequence techniques; their relevance for fungal systematics has been discussed by Harrington and Rizzo [70] and Taylor et al. [214]. Most ascomycetous species have been described by morphological characters [74] and-to a lesser extent-by other phenotypic characters, like pigmentation, secondary metabolites, growth at different temperatures or physiological tests [101]. The biological species concept has been applied to fungi mainly by using mating tests to identify mating compatible individuals. Mating tests, however, are not feasible with strictly asexual species or with species which produce sexual stages only in natural habitats and not in axenic cultures (approx. 20% of all fungal

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Limit of species

A

Limit of species

B

C

D

W

X

Y

Z

Incongruity within species

Fig. 1. Definition of a species according to the Genealogical Concordance Phylogenetic Species Recognition (GCPSR) (after Taylor et al. 2000).

species) [168]. The production of ascospores by itself is not sufficient to predict mating for homothallic species. The phylogenetic species concept is based on the criterion of monophyly and the requirement that a derived character be shared by the members of the species [214]. However, the delimitation of the gene can be difficult if the gene on which the phylogenetic analysis is based is polymorphic and has two alleles among individuals in a species. To avoid the subjectivity of species delimitation, a technique based on the comparison of more than one gene tree was proposed and named the Genealogical Concordance Phylogenetic Species Recognition (GCPSR) [214]. A species is defined by GCPSR if tree topologies of different genes have concordant branches. The method has been applied recently to several “difficult” species aggregations, resulting in the discovery of “cryptic” species, genetically isolated taxa which were not distinguished earlier based on morphology or mating tests. For example O’Donnell et al. [142] studied the Gibberella fujikuroi complex (Anamorph: Fusarium, Hypocreales) using nuclear sequences of three genes (>-tubulin gene exons and introns, mitochondrial small subunit (mtSSU) rDNA, and 5' portion of the nuclear 28S rDNA) and detected 45 species, among them 23 new to science. All biological species in the complex, defined previously by mating tests, were found by O’Donnell et al. [142] using GCPSR. The

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other species did not mate, and 23 of them had not been recognized by morphological characters. Further analyses of F. subglutinans, a member of the G. fujikuroi complex, showed that cryptic species were further subdivided into a number of smaller groups that appeared to be reproductively isolated in nature. This suggests that not only the existing F. subglutinans populations are in the process of divergence, but also that each of the resulting lineages are undergoing separation into distinct taxa [196]. Other examples of cryptic speciation were found within the meiosporic species complexes Botryotinia fuckeliana (Anamorph: Botrytis cinerea) [60], Histoplasma capsulatum (Anamorph: A. capsulatus) [88] and the mitosporic species complexes Candida albicans [203], Coccidioides immitis [93, 94] and Aspergillus flavus [59]. Examples among basidiomycetous fungi are Lentinula lateritia [77], and Filobasidiella neoformans (Anamorph: Cryptococcus neoformans) [51]. In all these cases, phylogenetic methods revealed species not detected by morphological characters or (in most cases) by mating tests. However, further analyses of these “cryptic” species often resulted in the identification of phenotypic differences, including pathogenicity. In this respect, the formae speciales of parasitic fungal species (for example, Fusarium oxysporum) which are defined by host preference but are usually not distinguishable by morphology, resemble the cryptic species. In general, GCPSR can be applied to all organismical groups and has been used successfully for prokaryotes. Among eukarya, however, few taxa outside the fungi were analyzed so far. Possible explanations are the haploidy of most fungi and that it might be easier to survey multiple genes in fungi because they have fewer paralogous genes and fewer complex multigene families [214]. Several studies indicate that in plants morphological differences may precede easily detectable genetic differentiation, in contrast to fungi, where the contrasting situation seems to prevail. SYSTEMATICS BASED ON MORPHOLOGICAL CHARACTERS

Traditionally, taxonomy and systematics of the fungi were mainly based on morphological (Euascomycetes) and physiological (yeasts) traits. Orders within the Euascomycetes were defined by the morphology, arrangements and methods of discharge of the asci and the development and form of the ascomata (the ascus-containing structure). There is considerable variation between the systems of higher categories proposed for Ascomycota [74] with the numbers and circumscription of recognized

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orders depending often on the interpretation of a single or a few characters, of which the evolutionary history and significance are not well understood. Still, these systems based on morphology are not only of historic interest, but through comparison with molecular data, might help to define the monophyly or paraphyly of certain combination of characters. As an example, the system according to Müller and Loeffler [128] is shown on Table 1. Morphology of the fruiting bodies is used to define the class of the Plectomycetes with ± globose, non ostiolate cleistothecia, the Discomycetes with mainly open, ± saucer or cup shaped apothecia, the Pyrenomycetes with flask-shaped perithecia, and the Loculoascomycetes with unwalled locules (pseudothecia). The class Loculoascomycetes is not accepted by a number of authors [for example, 74]. A single ascus type is typically found within the Plectomycetes (Prototunicate) and the Loculoascomycetes (bitunicate), whereas up to four different types of asci are found within the Discomycetes and Pyrenomycetes (Table 1). Ascus types are characterized by the thickness and modifications of the wall layers and the differentiation of the apical structures. The main types according to Müller and Loeffler [128] are: (1) Prototunicate: basically unitunicate, but with the wall lysing at or before maturity, lacking a differentiated apical structure; (2) Unitunicate: asci with a single functional cell layer. Apical structures are either inoperculate, when it opens with an irregular apical split to discharge the spores, or operculate, when it opens with an apical lid; and (3) Bitunicate: formed by two functional layers, that may or may not rupture or extend at discharge. The separation into very few basic ascus-types has been challenged by newer studies, which propose a much wider range of variation [18, 46, 81]. MOLECULAR APPROACHES TO FUNGAL PHYLOGENETICS

An excellent review on early stages of molecular fungal systematics was provided by Bruns et al. [31]. Methods like DNA-DNA-hybridization, restriction enzyme analysis and electrophoretic karyotyping only provided limited progress for the study of fungal relationships, but real breakthrough was achieved by the development of DNA sequencing techniques, especially since the introduction of PCR-technology [131] and the development of automated sequencing facilities. First sequences used were those of the ribosomal DNA cluster, facilitated by easy

Inoperculate Inoperculate

Phacidiales Lecanorales Ostropales

Heliotiales

Cyttariales

Unitunicateoperculate Inoperculate/ operculate Unitunicateinoperculate Inoperculate

Prototunicat Prototunicat

Ascosphaeriales, Elaphomycetales Peziales

Prototunicat

Ophiostomatales,

“Discomycetes”

Inordinate

Prototunicat

Microascales,

Hymenium Hymenium

Hymenium

Hymenium

Hymenium

Hymenium

Inordinate Inordinate

Inordinate

Inordinate

Eurotiales Onygenales Prototunicat

Arrangement

“Plectomycetes”

Asci Form Form

Order

Order

Single

Single Single

Single

Single

Single

Arrangement

Saprobes, Pathogens of plants and animals Saprobes, Pathogens of plants and animals Saprobes, Pathogens of plants and animals Pathogens of insects Obligate mycorrhiza

Occurrence

Table 1 Contd.

Saprobes, some plant pathogens Apothecium Fleshy Parasitic on stromata Nothofagus Apothecium Single, rarely Saprobes, plant in stromata pathogens Apothecium-like Single or in Saprobes or plant stromata pathogens Apothecium Single Lichenized Apothecium/ Single Lichenized or Perithecium saprobes

Apothecium

Cleistothecium Cleistothecium

Cleistothecium

Cleistothecium

Cleistothecium

Fruiting Bodies

Table 1. Systematics of the Euascomycetes based on morphological characters (mainly structure and arrangement of asci and fruiting bodies), after Müller and Loeffler,1982)

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Unitunicateinoperculate Inoperculate

Inoperculate

Sphaeriales Diaphortales

Clavicipitales Bitunicate

Prototunicate

Meliolales

Hymenium or single asci

Hymenium, asci of ten in bulk Hymenium

Hymenium

Lichenized Lichenized Lichenized or saprobes

Single or in stromata

Saprobes, plant pathogens, lichenized

Pathogens of plants, animals and fungi

Obligate ectopathogens of plants Single or Obligate accumulated endopathogens of plants Single Obligate ectopathogens of plants Single or Saprobes, pathogens in stromata of plants and animals Single or Single or in stromata in stromata

Single

Single Single Single

Perithecium-like, Single or Apothecium-like in stromata

Perithecium

Perithecium

Perithecium

Hymenium, Perithecium asci in different layers Hymenium Cleistothecium/ Perithecium

Prototunicate

Coryneliales

Cleistothecium

Hymenium Perithecium Hymenium Perithecium Hymenium, Apothecium partially with mazedium

Unitunicate, Few asci in operculate-like hymenium

Inoperculate Inoperculate Prototunicate/ unitunicate

Erysiphales

Graphidales Gyalactales Caliciales

“Loculoascomycetes” Dothideales

“Pyrenomycetes”

Table 1 Contd.

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amplification due to the high-copy number (approx. 100 copies per genome), the early availability of universal fungal primer sets [229] and the different substitution rate of the genes and spacers constituting the rRNA array, which enables to choose a target sequence well suited for analyzing a given taxonomic rank. In general, parts of the nuclear 18Sand 26S rRNA genes and the 5.8S rRNA gene are suited for studies of the relationships of higher taxonomic units, like phyla, classes, orders and families [21, 32, 44, 58, 126, 183]. Separation of genera and species is obtained using the Internal Transcribed Spacer (ITS) regions 1 and 2 [160], the D1/D2-domain of the 26S rRNA gene [102] and the Intergenic Spacer region (IGS) [6, 184]. Analysis of the 5S rRNA [64, 226] genes are no longer very popular, because they are too short and not very informative; mitochondrial small subunit (mtSSU) ribosomal RNA genes are used by several groups [118, 143, 144]. Differentiation of closely related species might require the use of at least two gene regions as has been shown by Scorzetti et al. [186] for biological distinct basidiomycetous yeasts, which could not be separated with D1/D2analysis alone. The status of the ribosomal DNA as the “golden standard of phylogenetic analysis” [164] is further underscored by the fact that most fungal genera, or even families, are represented in the GenBank only with rDNA sequences, in most cases by an approx. 600 bp long fragment spanning from the 3’ end of the 18S rRNA gene through the ITS1, the 5.8S rRNA gene, the ITS2 to the 5’ end of the 26S rRNA gene [5]. However, the base substitution rate of a given gene can show considerable variation and consequentially lead to taxonomic confusion. The most striking example are the Microsporidia, which were thought to be one of the deepest-branching eukaryotic lineages from rDNA analysis [224], but more recently were placed within the fungi based on combined analysis of multiple protein genes [78]. Such an artificial phylogenetic analysis caused by an extremely increased mutation rate of the gene under consideration is known as long branch attraction and results in a basal placement of the group [164]. Due to the repetitive nature of the rRNA genes and a weak selective constraint, relatively high intra-specific mutation rates are present among the multiple copies compared to single copy protein-coding genes [223]. However, paralogous copies of rDNA sequences do exist occasionally, as was shown by O’Donnell et al. [140] in the ITS2 region of Fusarium species. In addition, species delimitation in some fungal groups using rDNA is limited due to low sequence heterogeneity [149].

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In order to avoid systematic uncertainties, single-gene phylogenies of major eukaryotic lineages and important fungal groups are more and more replaced by a combined analysis of multi-copy and single copy genes of protein-coding and non-coding DNA sequences [118]. For example, the proponents of “deep hypha”, a consortium aimed to unveil major fungal relationships, propose to use information from a core set of seven genes, including 18S- and 26S rRNA genes, ITS1 and 2 regions, RNA polymerase I subunit B (RPB1), RNA polymerase II subunit B (RPB2), elongation factor-1 a, and mitochondrial ATPase subunit 6 [2]. The gene for the >-tubulin, coding for a subunit of the microtubules which are present in all eukaryotic cells, has been shown to be well suited for the estimation of deep-level phylogenies and the analysis of complex species groups [45]. Whereas the amino acid sequence of tubulin proteins is highly conserved, intron sequences of that gene are quite variable [116]. The recent development of universal primers of the >tubulin gene could further facilitate their use [45]. The RPB1-and RPB2genes are single copy genes with relatively slow evolutionary rates [110]. In parallel studies on the phylogeny of both asco- and basidiomycetous fungi [110, 111, 123] RPB1-and RPB2-genes resolve many internal branches that are unresolved in rDNA trees. These differences were explained by a deficit in sites evolving at slow- to moderate rates in rDNA [111]. Also used for fungal phylogenies are the nuclear glyceraldehyde 3phosphate dehydrogenase [24], the mitochondrial cytochrome oxidase (COX) genes, NADH dehydrogenase subunits [152], and the whole mitochondrial genome analysis [33]. Major breakthroughs for the understanding of evolutionary trends within the Ascomycota can be expected by comparative analysis of whole genomes, a new scientific field which is growing quickly. COMPARATIVE GENOMICS OF ASCOMYCETOUS FUNGI Genome analysis of Saccharomyces cerevisiae

The hemiascomycetous yeast S. cerevisiae (Saccharomycetales) was the first eukaryotic organism, of which the entire nuclear genome sequence was presented in 1996 [62]. Since then, analyses of the yeast genome has been invaluable as a reference in the sequencing of the human and other

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eukaryotic genomes, for comparative genomics, evolutionary studies, and for studies on gene functions and gene expression. S. cerevisiae occurs in both haploid and diploid strains, with the ellipsoid diploid cells being slightly bigger than the spherical haploid ones. On medium with limiting nitrogen resources, some diploid strains develop pseudohyphae with significantly elongated cells which do not detach after budding of the daughter-cell. The filamentous-form growth is regulated by a 26S proteasome [163]. Both heterothallic and homothallic strains occur in nature. Heterothallic strains are stable both as haploids and diploids, whereas the homothallic strains are stable only as diploids, because the transient haploids undergo mating type switching. The haploid nuclear genome of S. cerevisiae consists of approx. 13,500 kb, which are organized in 16 chromosomes with 200 to 2,200 kb each. A total of 6,183 open-reading-frames (ORF) of over 100 amino acids long were detected, approx. 5,600 of them are believed to be actual protein-coding genes [119]. In contrast, the human genome contains approx. 26,600 genes [175]. Similar to archae- and eubacteria, other Hemiascomycetes and S. pombe, the genome of S. cerevisiae is also highly compact. Circa, 72% of the total genome, represents actual genes, whereas relatively few repetitive sequences were found, in stark contrast to the genomes of plants and animals studied so far [175]. The average size of S. cerevisiae genes is 1.45 kb (ca. 483 codons). Only about 3.8% of the yeast ORF contains introns, compared to ca. 87.3% in human genes [175]. Ribosomal protein genes (rpg) contain an above-average amount of introns: the 137 yeast rpg genes contain 101 of the 234 introns present in the whole yeast nuclear genome. The ribosomal RNA genes are coded by approx. 120 copies of a single tandem array on chromosome XII. The nuclear genome contains 262 tRNA genes, 80 of which with introns. S. cerevisiae genes can be categorized into the following functional classes listed on Table 2. Genome analysis of Ashbya gossypii

A. gossypii (Syn. Eremothecium gossypii) is a filamentous plant pathogen which causes stigmatomycoses on cotton balls (Gossypium hirsutum) [7]. Based on analyses of nuclear rRNA genes, A. gossypii is a member of the Hemiascomycetes (Saccharomycetales), and clusters with the Eremothecium-group, which comprises unicellular and dimorphic plant pathogens transmitted by insects [160]. A. gossypii is one of the very few strictly filamentous members of the Hemiascomycetes which, together

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Table 2. S. cerevisiae genes related to functional categories (After Mewes et al. [4, 127, 128]) Functional Category Metabolism Energy Cell cycle and DNA processing Transcription Protein synthesis Protein fate (folding, modification, destination) Protein with binding function or cofactor requirement (structural or catalytic) Protein activity regulation Cellular transport, transport facilitation and transport routes Cellular communication/signal transduction mechanism Cell rescue, defense and virulence Interaction with the cellular environment Interaction with the environment (systemic) Transposable elements, viral and plasmid proteins Development (systemic) Biogenesis of cellular components Cell type differentiation Unclassified proteins

Approx. number of genes involved 1,488 363 995 1,060 473 1,130 1,019 237 1,028 233 552 457 8 123 70 844 448 2,054

with it’s small genome size and efficient gene targeting makes it an attractive model organism. Its biotechnological importance is based on the production of riboflavin (vitamin B2). The haploid genome is subdivided into seven chromosomes which contain 9,200 kb and encode for 4,718 proteins, 199 tRNA genes, and at least 49 small nuclear RNA (snRNA) genes [42]. So far, it is the smallest genome of any free-living eukaryote yet characterized. Only 40 copies of the ribosomal rRNA genes are present, compared to ca. 100 copies of the rRNA repeat unit in most other eukaryotes [228]. An extreme compactness of the genome is further obtained through the lack of transposons, subtelomeric gene repeats, and the rare occurrence of gene duplications. The average size of protein-coding genes in A. gossypii is only approx. 1.9 kb, compared to 2.1 kb in S. cerevisiae, 2.5 kb in S. pombe and 3.7 kb in N. crassa. Only 221 introns were identified in the

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whole genome, many of them at identical positions in S. cerevisiae homologues [42]. The gene number of 4,718 of this filamentous hemiascomycetous fungus is similar to that of the unicellular S. pombe, slightly smaller than that of S. cerevisiae and significantly smaller than that of filamentous Euascomycetes (Table 4), though a gene number between ca. 4,500 and 5,000 could be the minimum for a free-living eukaryote. For 95% of the protein-coding genes of A. gossypii, homologues in the S. cerevisiae genome were found [42], even if the overall GC-content differed significantly (A. gossypii: 58%, S. cerevisiae: 38%). Most of the homologue genes were found at syntenic locations with S. cerevisiae genes. Only 262 genes of A. gossypii did not show homology with S. cerevisiae genes, some of them instead showing homology with S. pombe genes. The amino acid identity of homologue genes of A. gossypii and S. cerevisiae varies considerably, ranging from less than 20% identity to nearly 100% [42]. No differences into highly conserved and less conserved genome regions could be detected based on analyses of the syntenic homologues. For most A. gossypii genes with homologues in S. cerevisiae, the synteny alters between one A. gossypii region and two S. cerevisiae regions (lying on two different chromosomes). When both S. cerevisiae regions are combined, the resulting gene order matches that of A. gossypii (“double synteny”). This pattern has been found for the overwhelming number of syntenic homologues of A. gossypii and S. cerevisiae. Non syntenic homologues were speculated to be former syntenic homologues, adjacent sequences of which were rearranged, resulting in insufficient evidence of synteny [42]. Analyses of the pattern of double synteny offers a way to postulate the gene order of the most common ancestor of the two species in question. Differences to that original pattern in the “modern” genome is a consequence of genome rearrangements and-in the case of S. cerevisiae-the loss of many genes after the genome duplication. By comparison of a limited number of genes of the Hemiascomycetes S. cerevisiae, Kluyveromyces lactis, K. marxianus, C. albicans and the Archiascomycete S. pombe, the extent of synteny was found to decrease with increasing evolutionary distance [90]. About 59% of adjacent gene pairs in K. lactis or K. marxianus are also adjacent in S. cerevisiae. Additional 16% of the Kluyveromyces neighboring genes are believed to correspond to the gene order in the ancestral species before the genome of S. cerevisiae underwent duplication. In contrast, only 13% of the gene

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linkages in the less closely related hemiascomycetous C. albicans are conserved, no linkages were found in the evolutionary distant S. pombe genome [90]. Analyses of gene order arrangements, chromosome numbers, and ribosomal RNA sequences suggest that genome duplication occurred before the divergence of the four species in Saccharomyces sensu strictu (S. cerevisiae, S. bayanus, S. pastorianus/carlsbergensis, S. paradoxus) with 16 chromosomes, but after this the lineage had diverged from S. kluyveri and the K. lactis/marxianus species group [90]. The pattern of double synteny between genes of S. cerevisiae and A. gossypii suggests that the two species originated from a common ancestor with seven or eight chromosomes. Later, a speciation event with an accompanying change in chromosome number generated the precursors of A. gossypii and S. cerevisiae, which diverged more than 100 million years ago [42]. Dietrich et al. [42] proposed a method for estimating relative evolutionary time scale which is based on the frequency of genome rearrangement events. Through analysis of the synteny breaks between the A. gossypii and the S. cerevisiae genome, they estimated 120 viable genome rearrangements in the A. gossypii lineage, and 180 genome rearrangements in S. cerevisiae, of which approx. 60 occurred before the genome underwent duplication. By assuming a similar rate of genome rearrangement in both species, the time span since the divergence of both species is approx. twice as long as the time span since the genome duplication in S. cerevisiae (which according to this estimation occurred ca. 50 million years ago) [42]. Comparative genomics among Hemiascomycetes

The Hemiascomycetes (“true yeasts”) comprise a relatively homogeneous and well characterized group of organisms sharing a small genome size and a low frequency of introns. Due to this and the availability of the whole S. cerevisiae genome, a comprehensive study on comparative genomics of that group so far unique among fungi (and eukaryotes) was conducted by a network of seven French laboratories. The project “Genolevures” [3, 50] was aimed to provide new data on molecular evolution, for example, to examine the conservation of chromosome maps, to identify “yeast-specific” genes and to review the distribution of gene families into functional classes.

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A random genomic DNA library was prepared by generating 4 to 5 kb long fragments from the following 13 yeast species: Candida tropicalis, Debaryomyces hansenii var. hansenii, Kluyveromyces thermotolerans, K. lactis, K. marxianus var. marxianus, Pichia angusta, P. sorbitophila, Saccharomyces bayanus var. uvarum, S. exiguus, S. servazzii, S. kluyveri, Yarrowia lipolytica and Zygosaccharomyces rouxii. The species were chosen representing the various branches of the Hemiascomycetes based on 18Sand 26S rDNA analyses. The ensuing phylogenetic relationships were in agreement with results of the decrease of synteny conservation [112], the amino acid sequence divergence [119] and the distribution of “Ascomycete-specific” genes [119]. Single-pass sequencing (up to 1.0 kb on average) of both extremities of each insert was performed, resulting in two random sequence tags (RSTs). Up to five ORFs were detected in a single RST. The genome coverage of the selected yeasts varied between 0.2 (using 2,500 RSTs) and 0.4 (using 5,000 RSTs). About 20,000 new genes were identified [188] and compared to the previously described approximately 6000 genes of S. cerevisiae. Two distinct groups of genes were found: genes commonly conserved through different phyla during evolution (“conserved genes”), and genes which did not show homologies to groups outside the Ascomycota (“Ascomycete-Specific”). A very low but significant level of species-specific genes [54] was also detected. In contrast to several prokaryotic groups, no strong evidence of gene acquisition through horizontal gene transfer was found [54]. Organization of Introns of Hemiascomycetous Yeasts

The compactness of hemiascomycetous genomes is based on the small number of genes, few repetitive sequences and the relatively low numbers of introns present in functional genes. Still, at least five distinct classes of introns were detected in the genome of S. cerevisiae (Table 3) based on location and mode of splicing [4]. Splicing of conventional nuclear pre-mRNA introns (or spliceosomal introns) occurs via a lariat intermediate through a spliceosome, a small nuclear ribonucleoprotein particle (snRNP) composed of five small nuclear RNAs (snRNAs or U [uridine-rich]-RNAs) associated with ca. 100 proteins. Splicing of tRNA introns involves the concerted action of three protein enzymes (without RNA co-enzyme): a site-specific endonuclease, a tRNA ligase, and a phosphotransferase. The HAC1 intron is a non-conventional pre-mRNA intron which uses protein

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Table 3. Major classes of introns found in S. cerevisiae (adapted from Mewes et al. 1997) Host genome

Intron class

Introns

Genes

Intron/Gene

S.cerevisiae genes

Nucleus

Pre-mRNA tRNA HAC1

270 61 1

263 61 1

1 or 2 1 1

ca. 5978a 274 ca. 5978a

Mitochondria

Group I Group II

9 4

3 2

1-4 1-3

2 8

a

ORF except questionable ORF

enzymes (i. e. the tRNA ligase among others). Mitochondrial Group I introns are mobile and show self-splicing, whereas Group II introns in general require a protein complex for splicing [4]. Introns in hemiascomycetous yeasts share a number of common features: 1. Hemiascomycetous genomes are intron-poor, only 1 to 5% of all nuclear genes have introns [27]. 2. Hemiascomycetous genes have only one (>96%) or two introns but never more [27]. In contrast, human genes contain on an average 7.8 introns per gene [175]. 3. Introns are generally short and can be divided into two lengthclasses: introns of ribosomal proteins can be close to 1,000 bp long, whereas non-ribosomal proteins in general have smaller introns (with 52 bp being the lower limit for an intron-size). Mean intron sizes varied, based on phylogenetic distance (for example introns in the Saccharomyces/Kluyveromces group are ca. 300 bp long, introns in the distant relative Pichia angusta: ca. 90 bp) [4]. Introns of human genes have an average size of 5,419 bp, but introns with more than 200,000 bp length also exist in humans. 4. Hemiascomycetous introns have conserved splice site motifs and specific contexts around the splice sites. 5. Analysis of the conservation of intron positions in homologous yeast species suggests that the ancestral nuclear yeast genes probably contained several introns, most of which have been lost since the ancestors of the Hemiascomycetes separated from the last common ancestor with S. pombe [27]. It has been speculated that the nuclear intron/exon mosaic of S. pombe resembles the

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original pattern, whereas the hemiascomycetous gene architecture might be a derived character [27, 89]. Variations from these common features were found within the introns of Yarrowia lipolytica, showing a larger size than was expected due to their phylogenetic distance to S. cerevisiae, a distinct 5’ splice site motif and a distinct nucleotide context around the splice sites [27]. Genome analysis of Schizosaccharomyces pombe

The fission yeast S. pombe (Schizosaccharomycetales) is used widely as a model system for classical and molecular genetics due to its simple cultivation and manipulation. Despite its name, S. pombe is not closely related to S. cerevisiae and other hemiascomycetous yeasts, but has been linked to a heterogeneous group of unicellular and filamentous fungi called Archiascomycetes by Nishida and Sugiyama [136]. The basal position of the Archiascomycetes compared to the Hemiascomycetes (“true” yeasts) and Euascomycetes has been challenged by newer analyses [162, 183] and is discussed further in this chapter. The sequencing of the whole nuclear genome was completed by a European consortium in 2002 [233]. With 4,824 genes on three chromosomes, the S. pombe genome is very compact and one of the smallest of eukaryotes found so far. Introns are present in 4,730 genes (43%). Intron length averages only 81 nucleotides, which is considerably shorter than those found in Metazoa and plants [233]. Nuclear rRNA genes are arranged as tandem arrays of 100-120 repeats. The average length of S. pombe genes is approx. 1,400 bp and the gene density for the complete genome is one gene every 2,528 bp, which is slightly less dense than in S. cerevisiae. Only 11 intact transposable elements were found in S. pombe, compared to 59 in S. cerevisiae. Gene numbers of the morphological simple fungi S. cerevisiae, S. pombe and A. gossypii are smaller than those in the largest prokaryotic genomes analyzed so far [5], i.e. Mesorhizobium loti has approx. 6,700 genes and Streptomyces coelicolor approx. 7,800 genes. In comparison, gene numbers of a parasitic prokaryote can be as low as approx. 500 (Mycoplasma genitalum), the smallest genomes of free-living prokaryotes contain approx. 1,500 genes (Aquifex aeolicus). The distinction between eukaryotic and prokaryotic cell organization, therefore, seems not to be determined by number of genes alone, but by the type of genes and expression pattern [233].

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Approximately two-thirds of the S. pombe genes have homologues in both the budding yeast S. cerevisiae and the nematode Caenorhabditis elegans, and are considered to represent a set of highly conserved “core” eukaryotic genes, among those coding for the cytoskeleton, compartmentation, cell-cycle control, proteolysis, protein phosphorylation and RNA splicing. “Yeast-specific” genes with homologues in S. cerevisiae but not C. elegans, make up 16% of the S. pombe genes. The remaining 14% of the genes identified in the S. pombe genome do not show homology with either S. cerevisiae or C. elegans genes and are considered to be “pombe-specific” [233]. Due to the little information available on genomes at the moment from closely related fungi belonging to the Schizosaccharomycetales or other Archiascomycetes, it is not clear whether these genes are really speciesor rather group-specific. Expression studies using DNA microarray spots hybridized with vegetative and meiotic samples [121, 122] showed that a disproportionate number of core genes is highly expressed in vegetative growing cells, at the same time many “pombe-specific” genes are expressed at low levels. During sexual development, especially during meiotic prophase, “pombe-specific” genes were expressed at high levels. Organism-specific genes involved in meiotic events could play a role for speciation by preventing chromosomal recombination between closely related species [121]. “Yeast-specific” genes, on the other hand, are overrepresented at the time of ascospore-development, which is a crucial step for all sexual Ascomycetes, but without homology in metazoic development. Genome analysis of Euascomycetes

Since the breakthrough in fungal genomic research, which was the sequencing of the complete S. cerevisiae genome in 1996 [62], relatively little progress was obtained in genomic research of filamentous, multicellular Euascomycetes. The first complete genome of a Euascomycete, that of Neurospora crassa (Sordariales) was presented in 2003 [55]. The sequencing of several other filamentous fungi (Table 4) is on-going, and high quality draft sequences are available for Aspergillus nidulans, A. terreus and Magnaporthe grisea. In contrast, over 50 bacterial genomes have been completed by this date [5]. Based on preliminary genomic analyses of members of only a few euascomycetous evolutionary lineages, the average genome size of a Euascomycete is between approx. 30 and 42 Mb, three to five- fold the

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Table 4. Genomes and Gene Complexity of selected ascomycetous fungi (modified after Kupfer et al. [101]) Order

Genome physical data Est. Chromosome Est. Gene Size Number number (MB) (complexity)

Hemiascomycetes Ashbya gossypii (syn: Eremothecium gossypii) Candida albicans Debaryomyces hansenii Saccharomyces cerevisiae Yarrowia lipolytica Archiascomycetes Pneumocystis carinii Schizosaccharomyces pombe Euascomycetes Plectomycetidae A. fumigatus A. nidulans Aspergillus oryzae Penicillium chrysogenusm Coccidioides posadasii Trichophyton rubrum Pyrenomycetidae Fusarium graminearum Trichoderma reesei Nectria haematococca (= Fusarium solani) Neurospora crassa Magnaorthe grisea

Saccharomycetales

8.7

7

4,700

Saccharomycetales Saccharomycetales Saccharomycetales Saccharomycetales

16 12.2 13.5 20.5

8 7 16 6

6,000 6,500 5,600 6,700

8 14

16(17) 3

n.d. 4,800

Eurotiales Eurotiales Eurotiales Eurotiales Onygenales Onygenales

29.2 30.1 36.7 34.1 29 30-40

8 8 8 4 4 4

10,000 9,900 14,000 8,200 n.d. n.d.

Hypocreales Hypocreales Hypocreales

36 33 40

4 7 10-16

11,600 7,800 8,800

Sordariales Sordariomycetes incertae sedis

43 39

7 7

10,000 11,000

Pneumocystidales Schizosaccharomycetales

n.d.: not determined

size of hemiascomycetous genomes. Assuming gene numbers of approx. 8–9,000 per euascomycetous genome [101], the typical gene densities vary between ca. 220 and 250 genes/Mb. This suggests a ca. 50% lower gene density in multicellular Euascomycetes compared to the

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predominantly unicellular Hemiascomycetes. The sizeable increase of gene numbers of Euascomycetes compared to the filamentous hemiascomycetous A. gossypii, could reflect a general increase in the complexity of morphological structures (for example, phialoconidiogenesis, and ascocarp development) and metabolic diversity. The relatively more pronounced increase of euascomycetous genome size, compared to the corresponding number of genes, is a typical feature of multicellular organisms, indicating increase of non-coding DNA sequences between and within protein (function) coding segments [101]. N. crassa (the bread mold) has been a well studied model organism for genetics and biochemistry for over 60 years since the seminal one gene-one enzyme model was established using this organism by Beadle and Tatum in 1941 [16]. N. crassa is a haploid organism with complex biological features, such as circadian rhythms, regulation of meiotic recombination, vegetative incompatibility reactions and response to light [83]. The N. crassa genome is segmented into seven chromosomes and contains approx. 39.9 Mb of DNA with a GC content of 54%. A total of 10,082 genes were predicted, corresponding to an average of 1 gene/ 3,700 bp, and an average of 1.7 short introns (134 bp long on an average) per gene. The nuclear rRNA gene complex consists of ca. 150 copies arranged in a tandem array [97, 98]. A mechanism for controlling genome expansion was detected in N. crassa [91, 187] which uses an induced mutagenesis of DNA repeats (“RIP”: repeat induced point mutation). RIP finds duplicated sequences that are longer than 400 bp in haploid nuclei of special premeiotic cells and introduces mutations from GC to AT, resulting in a high probability of nonsense codons and methylation of the mutated sequences [187]. The process eventually inactivates transposons and reduces increase of the genome size [91]. Multigene families, whose members have similar sequences, are also much rarer in N. crassa as would be expected from the genome size [83]. Only about 33% of the N. crassa genes have homologues in S. cerevisiae [30] and approx. 41% have no significant matches to other proteins. Of 1,421 genes with highest matches to either plant or animal proteins, 584 have no strong homology to genes of either S. pombe or S. cerevisiae, which led to the speculation that they may be involved with hyphal growth and multicellular development [83]. N. crassa lacks important genes involved in the control of asexual spores, which were found in the draft sequence of the A. nidulans genome. More comparative genomic

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studies of strictly asexual and strictly sexual fungi are needed to get a better insight into the evolutionary history of these diverse life styles. Ascomycete-specific Genes

Indications that S. cerevisiae may contain a high proportion of genes, which do not show homologies to genes from other phyla, date back to 1992 [145], when the sequence of the S. cerevisiae Chromosome III was proposed. Nearly half of the protein-coding genes of that particular chromosome could not be homologized with genes present at the sequence databases at that time [43]. Still, most of these ca. 3,000 socalled “orphan” genes are actively transcribed and ca 13% are essential for life [4]. Malpertuy et al. [119] compared the available sequence information from S. cerevisiae with the newly sequenced ca. 20,000 genes from 13 closely related hemiascomycetous species to detect the set of S. cerevisiae genes that do not share homologies in phylogenetic groups other than the Ascomycota. The term “maverick” for these genes was proposed in order to distinguish them from the term “orphan”, which has a functional connotation [119]. After comparison with 24 genomes available in the year 2000 (Caenoraphditis elegans, S. pombe, six archaeal and 16 eubacterial species) 2,743 S. cerevisiae genes out of 6,217 predicted ones (44%) remained without homologies. Additional comparison with human, mouse and rat sequences provided homologues to only 110 “maverick genes”. Malpertuy et al. [119] concluded that addition of novel organisms for comparisons should not significantly reduce the number of “maverick” genes in S. cerevisiae, as long as the novel organisms are not Ascomycetes. By comparisons of the S. cerevisiae genome with S. pombe 742 “maverick” genes were homologues. By comparing S. cerevisiae with the 20,000 sequences from the 13 hemiascomycetous yeasts, homologues of 3,680 S. cerevisiae genes of the “common” set (97.9% of the total) and 1,712 genes of the “maverick” set (69.8%) were identified. In conclusion, the genome of S. cerevisiae appears to contain 5,651 active genes of which ca. 32% have homologies only in other Ascomycota or, for most of them so far, only in other Hemiascomycetes. The proportion of “Ascomycete-specific” genes of S. cerevisiae with homologues in other Hemiascomycetes decreases with the increase in the phylogenetic distance of that species to S. cerevisiae increases [119]. The amino acid identities with S. cerevisiae were in the range from ca. 79% for S. bayanus var. uvarum to ca. 47% to the less

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closely related species D. hansenii var. hansenii, C. tropicalis and Y. lypolitica. Through comparison with the available functional classification of the S. cerevisiae genes, Gaillardin et al. [54] found that “Ascomycetespecific” genes are over-represented in the following biochemical pathways: cell wall biosynthesis, pheromone response, regulation of lipid, fatty acid and sterol biosynthesis, regulation of amino acid metabolism, regulation of nitrogen and sulphur utilization and regulation of carbohydrate utilization. PHYLOGENETIC RELATIONSHIPS AMONG THE ASCOMYCOTA AND THEIR ANAMORPHS

Numerous hypotheses on the phylogeny and evolution of higher fungi have been proposed (for references see [94, 117, 176, 216]. Phylogenetic trees inferred from multiple gene sequences (for example, 18S rDNA, >-tubulin) indicate that the Ascomycota and the Basidiomycota are sister taxa among the higher fungi, for which the term “dikaryophyta” was proposed, because both groups have pairs of unfused nuclei after mating and before nuclear fusion [for example, 32, 136, 162, 183, 200]. Savile [177] suggested that Taphrina was the closest survivor of a common ancestor of the Euascomycetes and the Basidiomycota. He suspected that two major lineages evolved from “Prototaphrina” a common ancestor. One major lineage led to the present day Taphrina and the higher Ascomycota – the Euascomycetes of today, whereas another major route led to the Basidiomycota (the Uredinales line and the parasitic Auriculariaceae line) through a “Protobasidiomycete”. The use of the Deuteromycota (or “Fungi imperfecti”) as a formal taxon is now decreased [74, 168, 200, 213]. Molecular sequence analyses do not always coincide with traditional definitions of fungal disorders or classes which were based mainly on the interpretation of ascoma characters. Plectomycetes, Pyrenomycetes, Loculoascomycetes, Discomycetes and other traditional class level categories are no longer used formally in the fungal classification system [74, 200]. The Ascomycota can be subdivided into three groups based on sequence analyses of the 18S rDNA and RPB2-genes [112, 162, 183], the qualitative and quantitative monosaccharide pattern of purified cell walls (Fig. 2) [183] and the ultrastructure of septal pores and urease activity [183, 219]. The three groups have been assigned to the taxonomic ranks of subphyla, containing a single class each by Eriksson et al. [49]: the

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Taphrinomycota (class Archiascomycetes), the Saccharomycotina (class Hemiascomycetes) and the Pezizomycotina (class Euascomycetes). The concept of a basal placement of the Archiascomycetes was proposed by Nishida and Sugiyama [134], based on sequence analysis of the 18S rRNA gene and confirmed by a number of authors. However, bootstrap support for the resulting tree topologies is generally low. Using a polyphasic biochemical and molecular approach, the group of Prillinger [162, 183] proposed an alternative view where the Euascomycetes and Archiascomycetes (for which they suggested the term “Protomycetes”), appear as a sister group (Fig. 2), whereas the Hemiascomycetes occupy a basal position. A similar tree topologies were obtained by Cai et al. [35] and Schweigkofler [180, 182] using a slightly different taxa sampling applying neighbour-joining and maximum methods. However, a recent maximum parsimony analyses of the 18S rRNA gene indicated that the Hemiascomycetes and Archiascomycetes (or Protomycetes) might be sister groups, although with low bootstrap support [115]. These contradictory results underline the limitation of ribosomal DNA sequences for the establishing of robust high-order phylogenies. S. cerevisiae and other Hemiascomycetes show a number of genetic, morphological and biochemical characters which could be interpreted as being primitive among the higher fungi: (1) the nuclear genome size of the Hemiascomycetes and of S. pombe appears to be about one-third the size of the Euascomycetes (Table 4). Three compact chromosomes are present in S. pombe, which resemble the chromosomes of higher eukaryotes. In S. cerevisiae 16 rather primitive chromosomes were found. (2) S. cerevisiae shows a rather unique cell cycle with the doubling of the spindle pole body and the formation of a short mitotic spindle already in the S-phase. The G2 phase is missing. On the other hand, the cell cycle of S. pombe resembles the higher eukaryotes with a characteristic G1, S, G2 and mitose phase [34]. (3) S. cerevisiae has a very compact nuclear genome with very few introns (223 introns, [4, 127, 128]). The higher frequency of introns in the genes of S. pombe (approx. 43%) resembles the situation in the higher eukaryotes. (4) S. cerevisiae is one of the few eukaryotes that can live without functional mitochondria. On the other hand, S. pombe requires functional mitochondria for survival like the higher eukaryotes.

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(5) The centromeres of S. cerevisiae are smaller and lack the repeated sequences, which are typical for higher eukaryotes. S. pombe has centromeres which resemble those in “higher” Ascomycetes and have a similar function [38]. (6) The length of the mating type loci S. cerevisiae has the shortest known idiomorphs for the mating type loci (a: 640bp, =: 750bp). The respective lengths for S. pombe are P: 1.1 kb, M: 1.1 kb; for the Euascomycetes Podospora pauciseta mat+: 3.8 kb, mat-: 4.7kb and Neurospora crassa A: 5.3 kb, a: 3.2 kb; for the Basidiomycetes Ustilago maydis a1: 4.5kb, a2: 8kb; and for the hymenomycetous yeast Cryptococcus neoformans mata: 35-45kb [129]). (7) The lack of fruit bodies among the Hemiascomycetes could be interpreted rather as a primitive than as a reduced character. Among the Protomyces/Schizosaccharomyces group, the Neolectales produce club-shaped fruit bodies, which are up to 7 cm tall and differ from other ascomycetous fruit bodies mainly by the absence of sterile hyphae (paraphyses) between the asci, the lack of ascogenous hooks (croziers) prior to ascus development, and the unusual combination of inoperculate asci having amyloid ascus walls [166]. (8) The coenzyme Q of the Hemiascomycetes contains a variable number of isoprene units ranging from Q-5 to Q-9 (Q-10 was found in Lipomyces lipofer). On the other hand, most strains of the Protomyces/Schizosaccharomyces clade analyzed so far contain coenzyme Q-10 (with the exception of Schizosaccharomyces octosporus, which has Q-9), resembling the Euascomycetes, which contain in most cases coenzyme Q-10 and Q-10 (H2). Coenzyme Q-9 was found only rarely in some euascomycetous strains (for example, Capronia parasitica, Symbiotaphrina spp.). No strain with less isoprene units was found within the Euascomycetes so far [183]. Basidiomycetous yeasts possess coenzyme Q systems with Q-7, Q-8, Q-9, Q-10 and Q-10 (H2) [102 and references therein]. (9) The Hemiascomycetes include morphologically primitive fungi (within the genus Eremothecium) with coenocytic “siphonal” [158] ontogenetic stages which resemble the Zygo-and Chytridiomycota. The yeast state, however, is thought to be a derived character, whereas the ancestral Ascomycetes were filamentous [112]. The

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hyphal growth was lost independently in the yeast forms, which are found in all three ascomycetous lineages. A yeast/hypha dimorphism is also common among all three lineages and is usually associated with a switch from saprophytic to parasitic nutrition. (10) The haplo-diplontic life cycle of S. cerevisiae also shows some similarities with the Chytridiomycota, where this type of a life cycle and diploid stages in contrast to the Euascomycetes are common [1]. (11) In a phylogenetic analysis using the >-tubulin, the PneumocystisTaphrina clade diverges in a paraphyletic manner from the Pezizomycotina, whereas the Saccharomycetales occupy a basal position among the Ascomycota [45]. S. pombe is paraphyletic to the Saccharomycetales, although separated by a long branch length. Hemiascomycetes

The Hemiascomycetes are a monophyletic group comprising predominantly unicellular (“true yeasts”) and a few dimorphic and filamentous taxa. Both strictly asexual and sexual species occur. Different from the Euascomycetes, sexual species of the Hemiascomycetes produce the asci never in ascocarps. Most of the approx. 700 species are saprophytic, but plant pathogenic (for example, Eremothecium, Ashbya) and human pathogenic (for example, Candida albicans) groups also occur. The seminal importance of S. cerevisiae as a model organism for genetics and molecular biology is as well known as it’s importance for biotechnology and human nutrition (“bakers yeast”, “beer yeast”, “wine yeast”). Within the Hemiascomycetes, Kurtzman and Fell [102] presently accept a single order, Saccharomycetales (Endomycetales), only. Based on the qualitative and quantitative monosaccharide pattern of purified yeast cell walls and complete 18S rDNA sequences (Fig. 2, [205]) Prillinger et al. [162, 183] proposed four different orders: Saccharomycetales, Dipodascales, Lipomycetales, and Stephanoascales. Whereas the Saccharomycetales and Lipomycetales can be delimited by the cell wall monosaccharide pattern and the presence or absence of extracellular amyloid compounds (Saccharomycetales: glucose, mannose, EAS: -; Lipomycetales: glucose, mannose, galactose, EAS: +) it is not possible to separate the Dipodascales and Stephanoascales by the cell

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Ustilago hordei Cystofilobasidium capitatum Mrakia frigida Auricularia polytricha Boletus satanas Filobasidiella neoformans Tremella moriformis

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ARCHIASCO MYCETES

Pyrenomycetidae EUASCOMYCETES

Plecto mycetidae

ASCOMYCOTA

Loculoasco mycetidae HEMIASCOMYCTES

Mixia osmundae Rhodosporidium toruloides Sporobolomyces roseus Microbotryum violaceum Schizosaccharomyces pombe Schizosaccharomycetales 88 Pneumocystis carinii Pneumocystidales Saitoella complicata 100 Taphrina populina Taphrina wiesneri 100 Taphrinales Taphrina pruni 6 lactucae-debilis 8 ProtomycesProtomyces macrosporus Protomyces inouyei 100 Neolectales Neolecta vitellina Neolecta irregularis Sclerotinia sclerotiorum Leotiales Blumeria graminis Erysiphales Xylariales 100 Xylaria carpophila Neurospora crassa Sordariales 100 Sordaria fimicola Gibberella pulicaris Hypocreales 98 Nectria aureofulva Microascus cirrosus Microascales Ceratocystis fimbriata Verticillium dahliae Phyllochorales Meliola juddiana 100 100 Meliolales Sporothrix schenckii 90 Ophiostomatales Ophiostoma ulmi 100 Leucostoma persooni Diaphortales Cryphonectria parasitica Phaeococcomyces exophiales Capronia mansonii Chaetothyriales Exophiala dermatitidis 98 Histoplasma capsulatum Onygenales Blastomyces dermatitidis 100 Ascosphaera apis Eremascus albus Ascosphaeriales Talaromyces bacillisporus Eurotiales Aspergillus flavus Mycosphaerella mycopappi 100 100 Pyrenophora tritici-repentis Cochliobolus sativus Pleospora herbarum 85 Alternaria alternata Pleosporales Alternaria brassicae 100 99 Cucurbitaria elongata Leptosphaeria maculans Septoria nodorum 89 Botryosphaeria ribis Aureobasidium pullulans Dothideales Dothidea insculpta Rhytisma salicinum Rhytismatales Symbiontaphrina kochii 97 Morchella esculenta Pezizales Helvella lacunosa Tuber magnatum 100 Lipomyces lipofer Lipomycetales Dipodascopsis uninucleata 100 Yarrowia lipolytica 100 Dipodascus albidus Dipodascales Galactomyces geotrichum 96 Clavispora lusitaniae Metschnikowia bicuspidata Pichia anomala Hanseniaspora uvarum 99 Saccharomycodes ludwigii Eremothecium sinecaudum Kluyveromyces lactis Candida glabrata 84 Zygosaccharomyces rouxii Torulaspora delbrueckii Saccharomyces cerevisiae Kluyveromyces polysporus Saccharomycetales 100 Debaryomyces hansenii 99 Candida albicans Candida tropicalis maltosa 100 Candida Saccharomycopsis capsularis Saccharomycopsis fibuligera 100 Dekkera naardensis Dekkera custersiana Dekkera bruxellensis 100 Issatchenkia orientalis Pichia membranifaciens 100 Candida blankii Stephanoascus ciferri Stephanoascales Candida valdiviana Candida bertae Candida edax

0.1

Fig. 2. Phylogenetic relationships of the Ascomycota. The unrooted Neighbor-Joining tree of 96 fungal species is based on the 18S rRNA gene; the Basidiomycota clade was used as the outgroup. The robustness of the tree was assessed by bootstrap analysis using 100 repeats. Only values greater than 65% are shown. Scale bar indicates accumulated changes per 100 nucleotides (adapted from Schweigkofler et al. 2002).

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wall monosaccharide pattern. Within both orders, glucose, mannose and galactose dominate, however, species with the glucose mannose pattern appear intermingled [205]. Sequences of the complete 18S rRNA gene are important to decide, whether a species belong to the Dipodascales or Stephanoascales. Extracellular amyloid compounds (starch formation) are absent in the Dipodascales and Stephanoascales. The order Dipodascales was already introduced by Batra [15]. However, based on molecular characteristics, many genera suggested by Batra cannot be included into this order. Additional multigene phylogenies are necessary to corroborate the orders Dipodascales, Lipomycetales and Stephanoascales [205]. Several genera within the Saccharomycetales are polyphyletic (for example, Candida and Pichia) [103, 205]. It was not possible to separate the genus Issatchenkia genotypically from the genus Pichia represented by its type species P. membranifaciens (Fig. 2 [103]). Genotypically, the genera Kluyveromyces, Saccharomyces, Torulaspora, and Zygosaccharomyces appear intermingled (Fig. 2, [35, 103]). K. delphensis is the closest teleomorphic species to Candida glabrata (Fig. 2, [35, 103]). C. albicans commonly occurs in the digestive tract. Candidiasis is by far the most important mycosis and vaginal infections are extremely frequent. C. albicans and S. cerevisiae were also considered as allergenic yeasts [81]. Although C. albicans is considered to be an asexual fungus, the recent identification of a mating-type locus [82] indicates the presence of (rare) sexual events. Based on ribosomal DNA sequencing, Messner et al. [125] and Prillinger et al. [160] included the two filamentous and plant parasitic species Eremothecium ashbyi and E. gossypii (= Ashbya gossypii) as well as the two dimorphic plant parasites E. coryli and E. sinecaudum in the family Saccharomycetaceae (Fig. 2). These data clearly indicate that yeasts cannot be separated taxonomically from filamentous fungi. Within the Stephanoascales Stephanoascus ciferrii and its anamorph C. ciferrii is the only pathogenic species so far. The species is often associated with animals and occasionally isolated from clinical specimens [53] as an agent of human onychomycosis. Some strains are strongly hyphal and produce conidia from characteristically inflated, denticulate heads. This anamorph has been described as Sporothrix catenata. No human pathogenic species is known so far from the order Lipomycetales. The order comprises typical soil yeasts (Babjevia, Lipomyces) and mycelial species (Dipodascopsis).

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Archiascomycetes

The Archiascomycetes are a very heterogeneous group of fungi comprising unicellular, dimorphic and strictly filamentous taxa which are saprophytes, plant pathogens or pathogens of vertebrates. Fruiting bodies are absent in most groups, but do occur in the Neolectales. Presently, four orders are accepted within the Archiascomycetes. These are the Neolectales, the Pneumocystidales, the Schizosaccharomycetales, and the Taphrinales [48]. Whereas the order Taphrinales was already introduced in 1928 from Gäumann and Dodge [99], the Schizosaccharomycetales, Neolectales, and Pneumocystidales were suggested recently based on molecular characters [47, 105, 154]. Based on the qualitative and quantitative monosaccharide pattern of purified cell walls, Prillinger et al. [154-157] considered the Archiascomycetes (Protomyces-Type, Schizosaccharomycetales) ancestral to the Euascomycetes and Basidiomycota, especially the Urediniomycetes sensu Swann and Taylor [204]. Morphological and ultrastructural data of Mixia osmundae [137] and 5S ribosomal RNA sequences from Protomyces inundatus [226] and Taphrina deformans [64] are additional characteristics which give support to the concept that the Archiascomycetes are ancestors of the Basidiomycota, as was originally suggested by Savile [176]. The anamorphic pigmented yeast Saitoella complicata was isolated only once from soil in Bhutan. Based on the qualitative and quantitative monosaccharide pattern of purified cell walls (glucose: 70, mannose: 23, galactose: 7) and ribosomal DNA sequencing (Fig. 2. [201]) the species belongs to the Archiascomycetes and can be excluded from the Urediniomycetes where it was included originally. A two-layered cell wall, a negative diazonium blue B test and positive urease activity are additional characteristics of yeasts and yeast stages which belong to the Archiascomycetes [183]. Enteroblastic budding of S. complicata [201], however, is a morphological trait shared with some groups of the Basidiomycota. The Neolectales are so far the only group of the Archiascomycetes where morphologically distinct fruiting bodies, apothecia similar to clavarioid basidiocarps, are produced. Landvik et al. [105] and Landvik [104] excluded the apothecial ascomycetous Neolecta vitellina and N. irregularis from the Euascomycetes (Fig. 2). Asci of N. vitellina are occasionally filled with numerous conidia and the ascospores become

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conidiogenous by producing a single apical collarette from which the phialoconidia emerge [166]. These features are similar to budding of ascospores within the ascus of Taphrina, and therefore, may not conflict with the proposed molecular phylogeny. The Pneumocystidales is the only order of the Archiascomycetes which harbors pathogens of vertebrates. Pneumocystis spp. are unicellular, obligate biotrophic organisms with a tropism for growth on respiratory surfaces of mammals [132]. Pneumocystis spp. have a number of features that are atypical for fungi (for example, cholesterol instead of ergosterol; [132]). Only a single copy of the nuclear ribosomal RNA gene cluster is present. Pneumocystis infection is widespread in AIDS patients and is often one of the first indications for the disease [41]. The molecular phylogeny and systematics of P. carinii have been controversial for a long time (Fig. 2, [132, 212, 213]) and its final placement within the fungi just occurred in the late 1980s. Four distinct species were described so far (P. jirovecii from humans, P. murina from mice, P. wakefieldiae and P. carinii from rats), but Pneumocystis isolates were detected from almost all mammalian species evaluated for their presence. A breakthrough in the understanding of the evolution and host specialization of these uncommon pathogens is expected by the outcome of the genome sequencing project of P. carinii, which is currently close to completion. The nuclear genome of P. carinii is haploid and contains approx. 8 Mb in 16 (or 17) chromosomes [39, 199]. Genes show a high AT-content (6065%) and usually contain introns, which are typically less than 50 bp long, but can be as numerous as 9 per gene [199]. The Schizosaccharomycetales comprise distinct unicellular saprophytic fermentative species which reproduce by fission. Many species of fission yeasts were described, but most have been found to be cospecific with one of three species: Schizosaccharomyces pombe, S. octosporus and S. japonicus [33]. Whereas S. pombe (originally known from tropical millet beer) is one of the best understood eukaryotic organisms on a genetic and biochemical level, relatively little is known about the two other species. Significant size length variation was found among the mitochondrial genomes of the Schizosaccharomyces species [33]; whereas the mtDNA of S. pombe is very compact with only about 19.4 kb, the mtDNA of S. octosporus is 44.2 kb and that of S. japonicus var. japonicus is over 80 kb. The size variation is largely due to noncoding regions [33].

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The phylogenetic position of the Schizosaccharomycetales within the Archiascomycetes is consistent using analysis of 18S rRNA- and RPB2genes [110]. However, certain phylogenies based on nuclear coded aminoacid sequences [9, 45, 107] and concatenated mitochondrial protein sequences [33, 109, 147] have placed S. pombe at the base of the Hemiascomycetes. Bootstrap support for these contradicting tree topologies are often low and depend on the genes used for analysis [33]. Comparative analysis of complete genomes of S. pombe and members of major fungal lineages (for example, S. cerevisiae, P. carinii, and N. crassa) is expected to unveil the phylogenetic position of the Schizosaccharomycetales. The Taphrinales (Fig. 2) comprise the Protomycetaceae and the Taphrinaceae [161]. All species are dimorphic fungi with the mycelial phase parasitic on ferns and especially woody dicotyledons, and the yeast phase saprophytic [95]. The mycelia and meiosporangia of the Protomycetaceae are polykaryotic, within the Taphrinaceae, mycelia are commonly dikaryotic and the young meiosporangia contain one nucleus only. Prillinger et al. [161] regarded the ascus of Taphrina as a “siphonal” germination state of a chlamydospore. This siphonal germtube acts as a meiosporangium, where an evolution from an indeterminate number of meiotic nuclei in the case of Protomyces to a single meiotic nucleus represented by the Taphrina species becomes obvious. Euascomycetes

The Euascomycetes, with comparatively well-developed fruiting bodies or ascomata, comprise the plectomycetes, pyrenomycetes, loculoascomycetes, laboulbeniomycetes, and discomycetes based on traditional morphological classifications (Table 1). In the monophyletic euascomycete lineage (Fig. 2, [198]) two major lineages, the Plectomycetidae with closed ascomata (cleistothecia) and the Pyrenomycetidae with flask-shaped ascomata (perithecia), appeared monophyletic, each receiving high bootstrap supports (Fig. 2). The tree topology in Fig. 2 supports the monophyly of the Plectomycetidae and Pyrenomycetidae as already detected by Berbee and Taylor [22] and Nishida and Sugiyama [136]. However, the concept of Gargas and Taylor [58] that the apothecial Pezizales (Ascobolus, Peziza, Gyromitra, Inermis, Morchella, Plectania) are ancestors of the cleistothecial and perithecial forms, is not supported by the tree topology shown in Fig. 2. Nannfeldt’s

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[134] phylogenetic hypothesis of a primitive apothecial ascomata with subsequent evolution of cleistothecial and perithecial forms is supported to some degree by phylogenetic analysis of both rDNA- and RPB2-genes [111, 162]. Within the Hypocreales, there are in addition some cleistothecial taxa, such as Heleococcum, Mycoarachis, and Roumegueriella, which have to be excluded from the Plectomycetidae [61]. Analyses of 18S rDNA support that both the loculoascomycetes (fissitunicate ascomycetes) and the discomycetes (apothecial ascomycetes) are not monophyletic (Fig. 2) [20, 57, 67, 68, 197, 198]. Within the fissitunicate ascomycetes, the loculoascomycete order Pleosporales appears as a monophyletic group, including the families Pleosporaceae and Lophiostomataceae, similarly the loculoascomycete order Dothideales may also constitute a monophyletic group, however, with weaker statistical support [20, 198]. On the other hand, the fissitunicate Chaetothyriales appear as a sister group of the Plectomycetidae or the lichen-forming Lecanorales and Peltigerales in the rDNA analysis [20, 183, 197, 198, 231]. However, a phylogeny based on the RPB2-gene suggests a sister relationship of the Chaetothyriales to a Pleosporales plus Dothideales clade, which would be in agreement with shared morphological characters (arcostromata, fissitunicate asci and apical pseudoparaphyses) [111]. Plectomycetidae

Based on morphological characters, six orders (Ascosphaerales, Elaphomycetales, Eurotiales, Microascales, Onygenales, and Ophiostomatales) were recognized within the class of the Plectomycetes [19, 130] (Table 1). Molecular data, however, suggest that only the Ascosphaerales, Elaphomycetales, Eurotiales, and Onygenales can be accepted within the subclass Plectomycetidae (Fig. 2). Presently it is not clear whether the Ascosphaerales, which lack ascocarps, and the hypogeous Elaphomycetales can be accepted as distinct orders or as families within the Eurotiales (Fig. 2, [1, 48, 106]. Eriksson and Winka [48] suggest five families comprising approx. 220 species within the Eurotiales (Ascosphaeraceae, Elaphomycetaceae, Eremascaceae, Monascaceae and Trichocomaceae) based on molecular characterization. The plectomycetous family Trichocomaceae within the Eurotiales includes cleistothecial teleomorphic genera which are associated with economically and medically important anamorphs, such as Penicillium,

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Geosmithia, Merimblia, Aspergillus, Paecilomyces and related genera [150, 151]. The teleomorphic genera associated with a Penicillium anamorph are Talaromyces, Hamigera, Eupenicillium, Trichocoma, Penicilliopsis, and Chromocleista [200 and literature cited therein]. Berbee et al. [23] and Sugiyama [200] gave a good overview about molecular phylogenetic studies in the Trichocomaceae. According to these studies, the genus Penicillium is not monophyletic; one group diverged first within the Trichocomaceae cluster and contains different Talaromyces species with the Penicillium-producing Talaromyces flavus and the Geosmithiaproducing T. bacillisporus. The second group consists of the Penicilliumproducing Eupenicillium javanicum, the Aspergillus-producing Eurotium rubrum and Neosartorya fischeri, as well as the Basipetospora-producing Monascus purpureus. The genera Penicillium and Aspergillus comprise a number of important human pathogens, some of which are aggressive invaders of immunocompromised patients. A. flavus produces aflatoxin, the most carcinogenic mycotoxin known. A. oryziae, used traditionally for soy fermentation in Asia, is a sister taxon of A. flavus; a cospecificity cannot be excluded [59]. The genus Paecilomyces contains several rare opportunistic human pathogenic fungi [41]. The entomopathogen P. tenuipes can be excluded from the genus based on 18S rRNA sequences data and instead it seems to be the anamorph of an entomopathogenic fungus of the genus Cordyceps (Hypocreales) [52]. The Onygenales comprise keratinophilic or cellulolitic soil fungi and numerous, often dimorphic pathogens of humans and other animals. Plant pathogenic taxa are not known. The Onygenales have cleistothecial fruiting bodies, sometimes with complex appendages and often brightly colored asci. Several genera are strictly asexual. Eriksson and Winka [48] suggest three families within the Onygenales (Arthrodermataceae, Gymnoascaceae, and Onygenaceae). Additional families were accepted by Hawksworth et al. (Myxotrichaceae) [74] and Unterreiner et al. (Ajellomcetaceae) [219]. A total of approx. 90 species has been described. The Arthrodermataceae and Onygenaceae harbor two important groups of human pathogenic fungi: the dermatophytes and the dimorphic systemic fungi. Dermatophytes are keratinophilic fungi which are capable of invading the keratinous tissues of living mammals. They are grouped into three categories on the basis of host preference

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and natural habitat. Anthropophilic species almost exclusively infect humans, rarely animals. Zoophilic species are essentially pathogens of non-human mammals or birds, although animal-to-human transmission is not uncommon. Geophilic species are soil-associated organisms, and soil per se or soil-borne keratinous debris is a source of infection for humans as well as other animals. The dimorphic systemic fungi are phylogenetically closely related to the dermatophytes and can be included in the family Onygenaceae [28, 108, 146]. Species of Blastomyces and Histoplasma have a teleomorph in Ajellomyces, for the other genera (Coccidioides, Emmonsia, and Paracoccidioides) no teleomorph is known so far. The natural habitat of all species are warmblooded animals. Several species show a temperature–dependent yeasthyphae dimorphism. In the environment at room temperature, they all produce a filamentous mycelial form, at 37°C in the tissue they reproduce as yeasts. The phylogenetic position of the Chaetothyriales varies depending on which gene is used for analysis. rDNA trees show an independent lineage remote from the remaining Loculoascomycetes and relatively close to the Onygenales and Eurotiales (Fig. 2 [68]) or the lichenforming Lecanorales and Peltigerales [2, 162, 231]. In contrast, analysis of the RPB2-gene [111] places them close to the Loculoascomycetes. Two families, the Chaetothyriaceae with approx. 75 species, and the Herpotrichiellaceae with approx. 35 species, belong to the Chaetothyriales [231]. The diversity of anamorphs in Chaetothyriales is remarkable and often it is difficult to distinguish them from anamorphs of the Dothideales [68]. Many species are dimorphic and are able to grow in the yeast form (black yeasts). Like the fissitunicate ascomycetes, the black yeasts are also polyphyletic and occur within the Chaetothyriales, Dothideales, and Pleosporales. Calcium regulates in vitro dimorphism in chromoblastomycotic fungi [124]. Basic morphological differences are associated with variations in thallus structure and maturation [208], due to which anamorphs of Chaetothyriales are smaller and more homogeneously pigmented than those of Dothideales [68]. Whereas Capronia species, a teleomorphic genus of the Herpotrichiellaceae, were described from plants such as Ericaceae [10], most anamorphs, however, have been associated with a wide spectrum of human diseases. Several genera within the Herpotrichiellaceae appear to be polyphyletic based on the analysis of 18Sr RNA genes (for example, Cladophialophora, Exophiala, Phialophora and Rhinocladiella [68]).

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Pyrenomycetidae

The Hypocreales are pyrenomycetous ascomycetes with unitunicate asci produced within fleshy, lightly or brightly coloured, typically ostiolate perithecial ascocarps [167]. They include, however, some cleistothecial species within the Bionectriaceae too [167]. Eriksson and Winka [48] and Rossman et al. [172] accept five families within the Hypocreales (Bionectriaceae, Clavicipitaceae, Hypocreaceae, Nectriaceae, and Niessliaceae), whereas Hawksworth et al. [74] accept only three: Clavicipitaceae, Hypocreaceae, Niessliaceae. Approx. 900 species were described. Many widespread and economically important plant pathogenic genera belong to the Hypocreales, for example, Fusarium, Cylindrocarpon, and Nectria. Production of mycotoxins (for example, deoxynivalenol DON, and zearalenone ZON) is common among this group. Several species are highly allergic [211], some of which are opportunistic human pathogens. C. destructans (teleomorph: Nectria radicicola) and C. lichenicola are two rare opportunistic clinical fungi within the Hypocreales. Species of the genus Trichoderma (for example, T. viride) are potentially pathogenic, toxigenic, and implicated in allergy or hypersensitivity pneumonitis [41, 191]. Teleomorphs are known in different Hypocrea species [41, 99]. Trichoderma viride, T. longibrachiatum, T. pseudokoningii, T. koningii, and T. harzianum were reported in recent years as occurring in humans [66]. Like Fusarium, genotypic identification methods are necessary to identify these species unequivocally [96, 100, 218]. Trichoderma spp. and Gliocladium spp. are aggressive mycoparasites which can attack established fungal pathogens using extracellular cell wall degrading enzymes and antibiosis. Among the toxic metabolites is 6-n-pentyl-2Hpyran-2-one (PPT), produced by some Trichoderma strains, and gliotoxin, viridian, and gliovirin, produced by Gliocladium. Alternatively, both Trichoderma and Gliocladium are fast growing saprophytes which can compete ecologically with plant pathogens colonizing potential infection sites [79]. Several formulations of Trichoderma have been developed for biological control of different plant diseases, for example, the gray mold of grapevine, caused by Botrytis cinerea. Based on ribosomal DNA sequencing Acremonium was shown to be a highly polyphyletic genus with affiliation to at least three ascomycetous orders [61]. Teleomorphs of Acremonium are found in several genera of the Euascomycetes (Emericellopsis, Hapsidospora, Nectria, Nectriella,

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Neocosmospora, Pronectria, and Thielavia). A larger number of species, including the type species A. alternata and the human pathogenic A. kiliense, exhibit affinities to the Hypocreales. A. kiliense has been described to cause ulcerative, nodulose hyalohyphomycosis, mycetomes, and keratitis [41]. Stachybotrys chartarum is an anamorphic soil and indoor air, toxigenic fungus which especially degrades cellulose. It has been associated with a number of human and veterinary health problems. Most notable among these has been a cluster of idiopathic pulmonary hemorrhage cases that were observed in the Cleveland, Ohio (USA), area [220]. A teleomorph, Melanomma pomiformis, is known only in S. albipes. It is included in the Niessliaceae of the Hypocreales based on morphology. Beauveria bassiana and Metarhizium anisopliae are anamorphic soil fungi well-known as insect pathogens. Both genera have attracted a great deal of attention because of their biological control potential [1]. Both produce mycotoxins, and the destruxins, a group of secondary metabolites produced by M. anisopliae, are considered an important new generation of insecticides [210]. Although the proteinaceous insect cuticle is an effective barrier to many fungi, insect pathogens, including Beauveria and Metarhizium, have a series of extracellular proteolytic enzymes that degrade native insect cuticle [148]. B. bassiana and M. anisopliae are polyphage and attack a wide host range. B. brongniartii acts specifically against the European cockchafer (Melolontha melolontha) and is used as a biological control agent. After the fungus penetrates the cuticle and reaches the hemocoel, yeast-like blastospores are produced, most probably to overcome the host defense system. After the death of the host, the fungus grows filamentous again, producing abundant persistent conidia [235]. This dimorphism is similar to that of some human pathogens (for example, Histoplasma capsulatum, and Sporothrix schenckii) and plant pathogens (for example Ophiostoma novo-ulmi). Ribosomal DNA analyses places these anamorphic fungi within the Hypocreales [85, 167]. A teleomorphic stage of B. brongniartii, Cordyceps brongniartii, has been described from East Asia, but was never found in Europe [84, 190]. Verticillium is a further anamorph genus of many insect- and plantpathogenic species which is heterogeneous. Although Messner et al. [126] suggested a relationship of the common plant pathogen V. dahliae to the Hypocreales, partial sequences of the 28S rDNA [167] and a more comprehensive phylogenetic tree of complete 18S rDNA sequences

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(Fig. 2) include V. dahliae within the Phyllachorales. In contrast, the entomopathogenic V. lecanii clusters within the Hypocreales [167]. Yeast-like endosymbionts (YLSs) of plant hoppers (Homoptera:Delphacidae) and aphids (Homoptera:Aphididae) were placed within the filamentous entomopathogenic genus Cordyceps (Clavipitaceae) based on parsimony analysis of combined 18S- and 26S rRNA genes [202]. These unicellular organisms are not culturable outside their hosts; they reproduce by budding and lack pseudohyphae or true hyphae. They occur in the host body fat and are transmitted to the offspring through the ovary [135, 138]. YLSs play a role in sterol utilization and nitrogen recycling for the hosts [202]. The adaptation to an intracellular life form and yeast-like growth is unique among the Hypocreales. The Ophiostomatales are usually characterized by often long-necked perithecial ascocarps, but with cleistothecia in one genus (Europhium; [71, 72]) and evanescent asci. A yeast stage is known in many species [69, 159]. Two families are accepted: the Kathistaceae and Ophiostomataceae, with a total of approx. 120 species. The qualitative and quantitative monosaccharide pattern of purified yeast cell walls resembles Protomyces and Taphrina species containing rhamnose [159]. Based on cell wall sugars, sensitivity to cycloheximide, ribosomal DNA sequencing (Fig. 2), and some additional characteristics [1] species of Ophiostoma are phylogenetically distinct from morphologically similar Ceratocystis (Microascales; Fig. 2) species [193, 194]. Many species are associated with scolytide and platypodide bark beetles in woody tissue, where they occur as saprophytic or slightly aggressive blue stain fungi [86]. Molecular quantification using Real-time PCR showed that several thousand fungal spores can be vectored by a single insect [185]. O. ulmi and O. novo-ulmi are highly virulent plant pathogenic fungus causing Dutch elm disease [1]. Leptographium wageneri causes black stain root disease on conifers in Western North America. Sporothrix schenckii (Fig. 2) is an anamorphic species which is the agent of human sporotrichosis, in addition it is known as allergenic fungus [170]. Species with a Sporothrix-like anamorph are also known within the Hemiascomycetes (Stephanoascales: Stephanoascus ciferrii) and Basidiomycota (Cerinostereus cyanescens). The Diaphortales comprise more than 400 species in the two families Melanconidaceae and Valsaceae [74]. Usually long-necked perithecia are produced in pseudostromata, which can be well developed. Anamorphs

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often produce two types of conidia in pycnidia: unicellular small microconidia and filamentous, multicellular macroconidia, which usually are not able to germinate. Members of the Diaphortales are saprobes or— more often—pathogens on barks and twigs of wooden plants, producing toxins and/or blocking the vessels of the host. Cryphonectria parasitica (Fig. 2), the causal agent of the chestnut blight, was introduced to New York City from Eastern Asia in the early 20th century and virtually wiped out the American Chestnut (Castanea dentata) within a few decades, which originally was the most widespread tree species in the Eastern US. Other important plant pathogens are Diaphorte eres on elms, Diaphortella aristata on birch and Phomopsis viticola on grapevine. The characteristics of the perithecial order Phyllachorales are not clear-cut and await further molecular characterization and investigation of additional species. One family (Phyllochoraceae) with approx. 1,150 species is accepted [74]. Well-developed stromata which are immersed in plant tissue are common. Species are often biotrophic or sometimes necrotrophic on stems and leaves, with the main distribution in the tropics. Important wilting agents in temperate zones are the anamorphic species Verticillium dahliae (Fig. 2) and V. albo-atrum. Glomerella cingulata, more often encountered as the anamorph Colletotrichum gloeosporioides, has been reported as a parasite of over 100 angiosperms, causing anthracnose of citrus and banana, among others. Sutton [204] provided descriptions for almost 40 species of Colletotrichum, however, morphological criteria are of little use when information on a plant pathogenic isolate is needed. Colletotrichum coccodes, C. dematium, and C. gloeosporioides are known as rare opportunistic fungi causing keratitis in humans [41]. The Meliolales are obligate biotroph parasites of plant leaves, with a primarily tropical distribution. In temperate northern areas (Central Europe, Western North America) only one species, Meliola niessleana is known from Ericaceae. Similar to the Erysiphaceae, the Meliolaceae show a high degree of host specialization. The mycelium consists of thick brownish hypha growing epiphytically with two-celled hyphopodia, from which a thin hypha penetrates the host cell where it produces a lobed haustorium. The brownish, mainly spherical fruiting bodies, produce few asci with two (sometimes up to eight) ascospores. Phylogenetic analysis of the 18S rRNA gene integrates the Meliolales within the Pyrenomycetes [174], whereas the Erysiphaceae occupy a more basal position among the Euascomycetes [173].

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The Sordariales comprise approx. 700 species in eight families (Batistiaceae, Catabotrydaceae, Ceratostomataceae, Chaetomiaceae, Coniochaetaceae, Lasiosphaeriaceae, Nitschkiaceae, and Sordariaceae) [74]. Many species are saprophytic on rotten wood (for example, Neurospora spp. is very common in burned forests), coprophile and cellulolytic species also occur. The perithecial order Sordariales is less important from the economic point of view; however, it harbors genera well-known from experimental mycology (Neurospora, Podospora, and Sordaria; Fig. 2). Neurospora sitophila (Sordariaceae), the red bread mold, is known to infest bakeries and cause considerable contamination. In culture, the fungus literally lifts the lid of Petri dishes, it contaminates by rapid growth and the production of enormous numbers of pinkish airdispersed conidia. N. sitophila is also known as an allergenic fungus [170]. Species of the Chaetomiaceae differ from Sordariaceae by their usually globose or ovoid asci that lack an apical ring and deliquesce within the perithecium or cleistothecium. In addition, the best known species have conspicuous hyphal appendages on the ascocarp surface. Chaetomium atrobrunneum, C. funicola, and C. globosum are known as rare opportunistic human pathogens [41]. Members of the Chaetomiaceae are cellulolytic and occur naturally on paper, tapestries, and cotton fabrics. C. globosum is also known as allergenic fungus [170]. The genus Phaeoacremonium was introduced by Crous et al. [40] to distinguish Acremonium species with pigmented vegetative hyphae and conidiophores. P. parasiticum, originally described as Phialophora parasitica is known as the agent of phaeohyphomycoses or mycetomes [41]. P. aleophilum and P. inflatipes are plant pathogens associated with Esca, an economically important grapevine trunk diseases widespread in several wine-producing countries. The Microascales comprise the two families Chadefaudiellaceae and Microascaceae with approx. 80 species [74]. Members of this order are characterized by lack of stromata; they often produce perithecia, but cleistothecia are formed by some species. Previously, Microascaceae was placed among plectomycetes by some mycologists because of the mature condition of evanescent, scattered asci [19]. More recently, Barr [11] placed the family among the pyrenomycetes. The work of Berbee and Taylor [21, 22] and Spatafora and Blackwell [194] using DNA analysis clearly has placed the group within the perithecial ascomycetes (Pyrenomycetidae). Pseudallescheria boydii is a cleistiothecial species

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which is frequently encountered as a saprophyte in soil, manure, and polluted water. The species is reported worldwide as an agent of whitegrain mycetomes. In addition, the fungus causes systemic infections in immunocompromised hosts or occurs in the respiratory tract where it is the agent of allergic reactions, sinusitis, pneumonia or systemic pseudallescheriasis [41]. Several rare opportunistic human pathogens are found in the Microascales (for example, Scopulariopsis brevicaulis). Ceratocystis fimbriata (Fig. 2) is an aggressive primary pathogen with a worldwide distribution and causes diseases on a wide range of plants (sweet potato, rubber, coffee, quaking aspen, prune, apricot, etc. [230]). Its long-necked perithecia are morphologically closely similar to Ophiostoma species [1]. Different from Ophiostoma, no yeast stage is known for Ceratocystis species. The Xylariales comprise three families (Amphisphaeriaceae, Clypeosphaeriaceae, and Xylariaceae) and approx. 800 species [74]. In most cases, the ascocarp is a ± globose perithecium, rarely a cleistothecium, growing superficially on plant tissue or immersed in a well-developed stroma. Within the Xylariaceae, sometimes stipitate and branched stromata are present, forming conspicuous, often black, structures up to several cm high. Most species are saprobes, weak plant pathogens or endophytes on bark, wood and roots worldwide. Xylaria carpophila (Fig. 2) is a saprophyte on fruits of Fagus sylvatica; Hypoxylon spp. is a typical secondary invader of plants which are weakened by the attack of more aggressive pathogens. Rosellinia comprises a number of important root pathogens, for example, R. necatrix on grapevines and apple trees, R. quercina on oaks and R. arcuata on tea plants. The Erysiphales or powdery fungi are a monophyletic group of obligate biotrophic plant pathogens on a wide range of phanerogams. The group has a worldwide distribution, but is best known from temperate areas in the northern hemisphere. The host range covers mainly dicotyledons and—to a lesser extent —monocotyledons (mainly Gramineae); gymnosperms and pteridophytes are never attacked. Based on a narrow species concept used by most researchers in the field, most taxa are biologically specialized with a more or less narrow host range not exceeding the limits of a single host family, or - more often - one or a few species within a host genus [29]. The one family Erysiphaceae contains 18 teleomorphic and four anamorphic genera with over 400 species. The characteristic ± spherical sexual fruiting body is somewhat intermediate

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between a true cleistothecium and a perithecium. Lack of an ostiole and the irregular arrangement of the asci in the early stages of the development are characters shared with Plectomycetes. On the other hand, in ripe fruiting bodies of the Erysiphales the asci appear regularly arranged, and discharge of the asci does not occur by decay of the ascocarp, but by a special form of opening similar to the real perithecia of the Pyrenomycetes. Most genera can be distinguished easily by the number and morphology of the characteristic appendages of the fruiting bodies. One to numerous asci are produced in a single ascocarp, each containing two to eight ascospores. Among the economically most important parasitic Erysiphales, are Uncinula necator (A: Oidium tuckerii) on grapevine, Blumeria graminis on wheat and other grasses, Podosphaera leucotricha on apple, and Sphaerotheca humuli (syn. S. macularis) on hops. Many species are attacked naturally by the mycoparasite Ampelomyces quisqualis (Pleosporales). Commercial formulations of A. quisqualis are used for several field crops and in glass houses for the biological control of powdery mildew. Loculoascomytidae

Nannfeldt [134] first segregated the classical Loculoascomycetes (which he called ascoloculares) from the other filamentous ascomycetes. While the other filamentous ascomycetes usually have thin-walled asci with a single functional wall layer, the Loculoascomycetes have thick-walled asci with two separable wall layers (fissitunicate [1]). Based on ribosomal DNA sequencing, Berbee [20] and Sterflinger et al. [198] showed that the Loculoascomycetes are not monophyletic and can be separated in three distinct orders: the Chaetothyriales, the Dothideales, and the Pleosporales. Although the jack-in-the-box-type ascus is a good marker for large, monophyletic loculoascomycete orders (Fig. 2), it must have evolved at least twice or been lost at least once [20]. The large order of the Dothideales comprises 58 families with 711 genera and ca. 5,000 species [74]. Whereas human-pathogenic species of the genus Cladosporium show a phylogenetic relationship to the Chaetothyriales (Herpotrichellaceae) and were, therefore, assigned to the genus Cladophialophora by Masclaux et al. [120], endophytic, mycoparasitic, plant-pathogenic, and saprophytic species of Cladosporium can be included in the Dothideales. C. cladosporoides and C. herbarum were commonly found as endophytes of grapevine [181].

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The human pathogen Hortaea werneckii is a dimorphic black yeast which exclusively causes tinea nigra palmaris on one or both hands, or on the sole. It is restricted to tropical or subtropical areas [41]. H. werneckii is halotolerant, having its natural habitat in salty environments [234]. Aureobasidium pullulans, a common dimorphic endophyte of grapevine [181] and saprophyte on plant leaves, occurs in addition as allergenic fungus [86, 170] and rare opportunistic pathogen in humans, where it causes keratitis, pulmonary infection, systemic infection, cutaneous infection, peritonitis, and invasive mycosis in AIDS patients [41]. Complete 18S as well as partial 26S rDNA sequences corroborate a Dothideales relationship of Aureobasidium species (Fig. 2, [120, 182, 198]). A. pullulans is used for the production of the polysaccharide pullulan and has been tested as a biocontrol agent for the control of fire blight on apple trees, caused by Erwinia amylovora. Hormonema dematioides is a very similar dimorphic fungus but can be differentiated from A. pullulans by the absence of synchronous conidiation, by different physiological profiles, and genotypic approaches like RAPD-PCR (Random Amplified Polymorphic DNA-Polymerase Chain Reaction) [180]. It is also occasionally pathogenic to humans [66]. Nattrassia mangiferae (Synanamorph: Scytalidium dimidiatum) is known as a plant pathogen but is also commonly reported from human superficial infections in subtropical and tropical countries. In humans it causes expanded hyperkeratosis with scaling of the skin in extremeties, as well as onychomycoses [171]. Botryosphaeria spp. are pathogens of conifers and dicots. B. rhodina is known as a teleomorph of Lasiodiplodia theobromae. B. rhodina clusters with B. ribis (Fig. 2) in the Dothideales [20]. Cenococcum geophilum is a cosmopolitan and is the most widespread ectomycorrhizal fungus. It has an extremely broad host and habitat range. Parsimony and distance analyses positioned C. geophilum as a basal, intermediate lineage between the two Loculoascomycete orders, the Pleosporales and the Dothidiales [l14]. At least four independent lineages of mycorrhizal fungi have so far been identified among the Ascomycota. The Pleosporales form a monophyletic group with high bootstrap support (Fig. 2, [20]). In phylogenetic trees based on complete 18S rDNA sequences, they commonly appear as a sister group of the Dothideales (Fig. 2). Alternaria alternata is a saprophyte, opportunistic plant pathogen and a common endophyte [180], but may also cause skin

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lesions in humans after trauma and is known to cause allergic reactions. Teleomorphs of Alternaria species are known in Pleospora and Lewia [24], they can be included in the Pleosporales (Fig. 2, Pleosporaceae; [24]) based on complete sequences of 18S rDNA (Fig. 2). Eriksson and Winka [48] accept four families within the Pleosporales (Leptosphaeriaceae, Lophiostomataceae, Melanommataceae, and Pleosporaceae). Species of Phoma are widespread plant pathogens, some of which have teleomorphs in Leptosphaeria (for example, Phoma lingam) or chlamydospores with muriform septation, resembling the conidia of Alternaria (for example, Phoma glomerata). Partial sequences of the 18S rDNA of Phoma species exhibit a relationship to the teleomorph Cucurbitaria, which can be included in the Pleosporales (Leptosphaeriaceae [182]). Leptosphaeria senegalensis and L. thompkinsii cause mycetoma in Africa [41]. Coniothyrium fuckelii is the anamorph of L. coniothyrium. It is known as a plant-pathogen especially on Rosaceae and also from human infections [41]. The genus Cochliobolus harbors many plant- and human-pathogenic fungi [24, 41]. Based on ITS and glyceraldehyde-3-phosphate dehydrogenase gene sequences, Berbee et al. [24] support a suggestion from Tsuda and Ueyama [217] to separate the genus in two closely related genera Cochliobolus and Pseudocochliobolus. Additional molecular sequences, especially, from human-pathogenic species, however, are necessary to corroborate this concept further. Species of Cochliobolus exhibit a Bipolaris anamorph. Bipolaris hawaiiensis is a common saprophyte on plant material. Sinusitis, pulmonary and cerebral mycosis have been reported [41]. B. australiensis and B. papendorfii are rare opportunistic human pathogenic fungi [41]. Stemphylium is the anamorph which belongs to the teleomorphic Pleospora species (Fig. 2, [24]). S. macrosporoideum is known from a mixed infection in antromycosis. [14]. S. botryosum is together with Alternaria species considered one of the most important mold allergens in the United States [86]. There is some molecular evidence that the genus Pleospora (P. herbarum, P. rudis; [198]) is heterogeneous. Discomycetes

The Leotiales are the largest of the orders of inoperculate discomycetes. They are characterized by either cup- or disc-shaped apothecia, or asci that have more or less thickened apices. Although the apothecial

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discomycetes are not a monophyletic group based on molecular characters [58], there is support that the Leotiales, Lecanorales, and Pezizales are monophyletic orders within the apothecial Euascomycetes. Many members of the Leotiales live as soil saprophytes; some are parasitic on plants and belong to the worst fungal pathogens. Among these are Monilinia fructicola, the cause of brown rot of stone fruits and Sclerotinia sclerotiorum, the cause of lettuce drop and other vegetable diseases (Fig. 2). Moserella radicicola produces hypogeous apothecia [153]. Botrytis cinerea and its teleomorph Botryotinia fuckeliana is known as gray mold of grapevines and numerous other crops. The fungus is also known to induce “noble rot” which enhances the sweetness of the grapes used for dessert wines. In addition, B. cinerea is a common allergenic fungus [168]. Based on partial sequences of the 18S rDNA, B. cinerea can be assigned to the Sclerotiniaceae [80]. The Rhytismatales comprise more than 400 species and three families (Ascodichaenaceae, Cryptomycetaceae, and Rhytismataceae). The fruiting bodies are apothecia, often immersed in host tissue or on well-developed stroma. The main distribution is in temperate zones, most species are saprobes or necrotrophic parasites, often on leaves, but also on bark and wood [74]. Endophytic growth is widespread. A well-known member of the order is Rhytisma acerinum, the causal agent of tar spot on maple. The Pezizales are a large monophyletic order that contain the species commonly called operculate discomycetes as well as derived hypogeous forms that have evanescent asci with ascospores spread by mycophagy. O’Donnell et al. [141] recently investigated phylogenetic relationships among ascomycetous truffles and the true and false morels. The results indicate that the hypogeous ascomycetous truffle and truffle-like taxa studied represent at least 5 independent lineages within the Pezizales. The data also suggest that several epigeous and most hypogeous taxa have been misplaced taxonomically. There is strong support for a Tuberaceae-Helvellaceae clade which is a monophyletic sister group of a Morchellaceae-Discinaceae clade [162]. Members of the Morchellaceae, common morel (Morchella esculenta) and Tuberaceae (Tuber melanosporum) are well-known as food. There are only some poisonous species (for example, Gyromitra esculenta) of clinical importance. The Laboulbeniales comprise five families and approx. 2,000 species. Four families are obligate biotrophic ectoparasites of insects and -less often- mites and millipedes (Ceratomycetaceae; Euceraotomycetaceae,

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Herpomycetaceae, and Laboulbeniaceae). The fifth family, the Pyxidiophoraceae, is a group of morphologically distinct filamentous mycoparasites with an arthropod-associated phoretic stage. The Pyxidiophoraceae were placed within the Laboulbeniales based on common morphology of asci development and ascospores [26] and the phylogenetic analysis of 18S rRNA genes [25]. The Laboulbeniales rarely damage their host seriously and do not penetrate host tissue below the chitin layer. Most species show a high level of host specificity, infecting only hosts that belong to the same genus or group of closely related genera. Also widespread within the Laboulbeniales is a “position-specificity”, the often precise occurrence of parasitic thalli on restricted parts of the host. Position-specificity was explained in some species by the precise transmission of ascospores from one host to a new one during copulation. Due to an unusual morphology with very small stromata composed of a basal black haustorium and a dark cellular thallus [74], the systematic position of the Laboulbeniales remained unsolved till recently. Schwantes [179] erected the subclass Laboulbeniomycetidae as a link between the predominantly unicellular Protoascomycetidae (yeasts) and the filamentous Euascomycetidae, whereas Cavalier-Smith [37] excluded the Laboulbeniales from the Ascomycota and put them into a new class Zoomycetes together with a heterogeneous group of invertebrateassociated fungi. Recent phylogenetic analysis of partial sequences of 18S rRNA genes places the Laboulbeniales unambiguously within the Euascomycetes [227]. The robustness of that hypothesis was reinforced by parsimony analysis and maximum likelihood ratio tests applied to trees constructed using different topological constraints [227]. Symbiotaphrina buchneri and S. kochii (Fig. 2) are yeast-like endosymbionts (YLSs) of anobiid beetles which inhabit the mycetome between the foregut and midgut of the host. These symbionts are transmitted vertically via colonization of the insect egg, newly hatched larvae eat the fungi on the egg case [139]. In contrast to the YLSs belonging to the Hypocreales, Sympiotaphrina species are culturable on artificial media. The YLSs provide nutrients to their host and detoxify substances, which are deleterious to the insects. Due to morphological characters, Sympiotaphrina spp. were integrated originally in the family Taphrinaceae (Archiascomycetes). Phylogenetic analysis of the 18S rRNA gene, however, showed them to be members of the Euascomycetes, close to the class Discomycetes [139]. The genus

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Symbiotaphrina appears to be a monophyletic group distinct from the other major lineages within the Euascomycetes, which underwent a morphological adaptation from filamentous to unicellular growth form due to the endosymbiontic life form. Acknowledgments

Sincere thanks to Prof. H. Prillinger, University of Agricultural Sciences, Vienna, Austria and Prof. M. Garbelotto, University of California, Berkeley, CA, for their support and advice. For financial support during the last few years. I thank the Austrian Ministry for Agriculture, the European Commission, and the Department of Forestry and Fire Protection, State of California. References [1] Alexopoulos CJ, Mims CW, Blackwell M. Introductory mycology, ed 4, revised. New York: John Wiley & Sons, 1996. [2] Anonymous. Homepage of the project “Deep Hypha”. http://ocid.nacse.org/research/ deephyphae/ [3] Anonymous. Homepage of the project “Genolevures”. http://cbi.labri.fr/Genolevures/ index.php [4] Anonymous. Homepage of MIPS (Munich Information Center for Protein Sequences). http://websvr.mips.biochem.mpg.de/proj/eurofan.index.html [5] Anonymous. Homepage of the National Center for Biotechnology Information (NCBI) http://www.ncbi.nlm.nih.gov/ [6] Appel DJ, Gordon TR. Relationships among pathogenic and nonpathogenic isolates of Fusarium oxysporum based on the partial sequence of the intergenic spacer region of the ribosomal DNA. Mol Plant-Microbe Interact 1996; 9: 125 –138. [7] Ashby SF, Nowell W. The fungi of stigmatomycosis. Ann Bot 1926; 40: 69–84. [8] Baldauf SL, Palmer JD. Animals and fungi are each other’s closest relatives: Congruent evidence from multiple proteins. Proc Natl Acad Sci USA 1993; 90: 11558–11562. [9] Baldauf SL, Roger AJ, Wenk-Seifert I, Doolittle WF. A kingdom-level phylogeny of eukaryotes based on combined protein data. Science 2000; 290: 972–977. [10] Barr ME. Prodromus to class Loculoascomycetes. Amherst, Mass: Newell 1987. [11] Barr ME. Prodromus to nonlichenized, pyrenomycetous members of class Hymenoascomycetes. Mycotaxon 1990; 39: 43–184. [12] Bartnicki-Garcia S. Cell wall composition and other biochemical markers in fungal phylogeny. In: Harborne JB, ed. Phytochemical phylogeny. London: Academic Press, 1970: 81–103. [13] Bartnicki-Garcia S. The cell wall: a crucial structure in fungal evolution. In: Rayner ADM, Brasier CM, Moore D, eds. Evolutionary biology of the fungi. Cambridge, UK: Cambridge Univ Press, 1987: 389–403.

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[14] Bassiouny A, Maher A, Bucci TJ, Moawad MK, Hendawy DS. Non invasive antromycosis (diagnosis and treatment). J Laringol Otol 1982; 96: 215–228. [15] Batra LR. Taxonomy and systematics of the Hemiascomycetes (Hemiascomycetidae). In: Subramanian CV, ed. Taxonomy of fungi. Chennai, India: Univ of Madras, 1973: 187–214. [16] Beadle G, Tatum E. Genetic control of biochemical reactions in Neurospora. Proc Nat Acad Sci USA 1941; 27: 499–506. [17] Begerow D, Bauer R, Oberwinkler F. Phylogenetic studies on nuclear large subunit ribosomal DNA sequences of smut fungi and related taxa. Can J Bot 1997; 75: 2045–2056. [18] Bellemere A. Asci and ascospores in ascomycete systematics. In: Hawksworth D. ed. Ascomycete systematics: problems and perspectives in the nineties. New York: Plenum Press; 1994: 111–126. [19] Benny GL, Kimbrough JW. A synopsis of the orders and families of Plectomycetes with keys to genera. Mycotaxon 1980; 12: 1–91. [20] Berbee ML. Loculoascomycete origins and evolution of filamentous ascomycete morphology based on 18S rRNA gene sequence data. Mol Biol Evol 1996; 13: 462–470. [21] Berbee ML, Taylor JW. Convergence in ascospore discharge mechanism among pyrenomycete fungi based on 18S ribosomal RNA gene sequence. Mol Phylo Evol 1992a; 1: 59–71. [22] Berbee ML, Taylor JW. Two Ascomycetes classes based on fruiting–body characters and ribosomal DNA sequence. Mol Biol Evol 1992b; 9: 278–284. [23] Berbee ML, Yoshimura A, Sugiyama J, Taylor JW. Is Penicillium monophyletic? An evolution in the family Trichocomaceae from 18S, 5.8S and ITS DNA sequence data. Mycologia 1995; 87: 210–222. [24] Berbee ML, Pirseyedi M, Hubbard S. Cochliobolus phylogenetics and the origin of known, highly virulent pathogens, inferred from ITS and glyceraldehyde-3-phosphate dehydrogenase gene sequences. Mycologia 1999; 91: 954–977. [25] Blackwell M. Minute mycological mysteries: the influence of arthropods on the lives of fungi. Mycologia 1994; 86: 1–17. [26] Blackwell M, Malloch D. Pyxidiophora. A link between the Laboulbeniales and hyphal ascomycetes. Memoirs of the New York Botanical Garden (Rogerson Festschrift) 1989; 49: 23–32. [27] Bon E, Casaregola S, Blandin G, Llorente B, Neuvéglise C, Munsterkotter M, Guldener U, Mewes HW, Van Helden J, Dujon B, Gaillardin C. Molecular Evolution of Eukaryotic Genomes: Hemiascomycetous Yeast Spliceosomal Introns. Nucleic Acids Res 2003; 31: 1121–1135. [28] Bowman BH, Taylor JW. Molecular phylogeny of pathogenic and non-pathogenic Onygenales. In: Reynolds DR, Taylor JW, eds. The fungal holomorph: mitotic, meiotic and pleomorphic speciation in fungal systematics. Wallingford: CAB International 1993: 169–178. [29] Braun U. The Erysiphales (powdery mildews) of Europe. Jena: Fischer Verlag, 1996. [30] Braun EL, Halpern AL, Nelson MA, Natvig DO. Large-scale comparison of fungal sequence information: mechanisms of innovation in Neurospora crassa and gene loss in Saccharomyces cerevisiae. Genome Res. 2000; 10: 416–430.

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Yeast Biodiversity and Evolution VU NGUYEN THANH Department of Microbiology, Food Industries Research Institute, Hanoi, Vietnam

ABSTRACT Yeast has been in existence in civilization for thousands of years and the study of this group of microorganisms has almost two centuries of history. Vast majority of the studies, thus far, were focused on the model yeast Saccharomyces cerevisiae. Yeasts in nature are much more diverse than what is presented in scientific literature or the common perception about yeast(s) as a sugar-fermenting microorganism. The diversity and evolution of yeasts were revealed by morphological, biochemical characteristics, sexual behavior, nucleic acid sequence, and ecological habitat. Around 700 of known yeast species are now placed under three classes of Ascomycota (Archiascomycetes, Euascomycetes, and Hemiascomycetes) and three classes of Basidiomycota (Ustilaginomycetes, Urediniomycetes, and Hymenomycetes). The biotechnology potential of yeasts is immense. This chapter gives an overview of the diverse world of yeasts through taxonomic and classification approach. Key Words: Yeast, biodiversity, evolution, taxonomy, classification

INTRODUCTION

Yeast has been in existence in our civilization for thousands of years and its study has roughly two centuries of history. The discovery of yeast as a living organism dats back to the very beginning of the birth of microbiology when Antonie van Leeuwenhoek observed the yeast cells from a beer precipitate. Being one of the most important organisms in the fermentation industry and biotechnology, as well as the model for Address for correspondence: Dr. Vu Nguyen Thanh, Department of Microbiology, Food Industries Research Institute, 301-Nguyen Trai, Thanh Xuan, Hanoi, Vietnam. Tel: (844)5589004; Fax: (844)-8584554; E-mail: [email protected].

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eukaryotic cells, yeast has been a subject of intensive study for many decades in biochemistry, cell biology, classical genetics, and molecular biology. In a unamimous global effort, yeast was the first eukaryotic organism whose genome was fully sequenced and which triggered an unprecedented research activity of “post-genomic era”. The knowledge accumulated on yeast is enormous. Vast majority of the studies have so far been focused on the model yeast Saccharomyces cerevisiae, and to a lesser extent, to a few other yeasts that are important to biotechnology like Schizosaccharomyces pombe, Kluyveromyces marxianus, Pichia pastoris, Phaffia rhodozyma or human pathogens like Candida albicans, Cryptococcus neoformans. Yeasts in nature are much more diverse than that presented in the scientific literature. This chapter gives an overview of the diverse world of yeasts through a taxonomic and classification approach. DEFINITION OF YEAST

Yeast has long been in use since human history. The earliest evidence of established winemaking practice dates back to 5400-5000 BC [67, 69] (Fig. 1). A recent study on the trace of mixed fermented beverage in

Fig. 1. Archeological records of early winemaking practice. Left - This narrow necked jar was one of the six excavated in the “kitchen” of a Neolithic residence at Hajji Firuz Tepe (Iran), dating to 5400-5000 BC. The reddish residue inside the jar was found to contain calcium tartrate and terebinth tree resin. The jar originally contained a liquid, judging by its relatively long, narrow neck, and the residue being confined to its bottom half. Tartaric acid is present in abundance in Middle East, only in grapes. Thus, it was assumed that the jar had contained wine. The terebinth tree resin could be used as a preservative against further acidification by acetic acid bacteria. Anaerobic condition could be ensured by clay stoppers that were found nearby. This was the earliest direct indication of the winemaking practice by mankind [from 67] (the image was kindly provided by University of Pennsylvania Museum of Archaeology and Anthropology). Right - Vintage scene on the tomb of Nakht, Thebes, 1500 BC [2] (photo courtesy of the Metropolitan Museum of Art).

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China has even pushed the date to 7000 BC [68]. A precipitate under wine in a sample which dated to 3400 BC was proved to contain Saccharomyces cerevisiae DNA [16]. In Chinese and Japanese the word for yeast is , it consists of two elements “fermentation” ( ) and “mother” ( ), which literally means mother, or the cause of fermentation. Similarly, in other languages, the closest equivalents for “yeast” are gist (Dutch), levure (French), Hefe (German), (Russian). All these refer to the process or agents causing lifting, frothing, and foaming of organic substrates. The similarity in meanings of words from distant cultures and languages evidences the ancient and constant contact of mankind with yeast and its activity. The word “yeast” is commonly understood in daily life. However, the usage of this word as a terminology has caused continuous discussions and ambiguity. Biologists in the field of molecular research define “yeast” as Saccharomyces cerevisiae. Meanwhile, for the brewing industry, “yeast” is often restricted to the group of technological species termed Saccharomyces sensu stricto which consists of S. cerevisiae, S. pastorianus, S. bayanus, and the natural relative S. paradoxus. The environmental yeasts are often referred to as “wild (unwanted) yeasts”. In reality, yeast is extremely complex and diverse in nature. The latest yeast identification manual [53] includes 689 species. To date, more than 300 species of yeasts have been described. The terminology “yeast” has its own evolution, reflecting our understanding of the nature and diversity of this exciting group of fungi. One of the best definitions of yeasts in general is “fungi, basidiomycetes or ascomycetes, whose vegetative stage is unicellular, and multiplies by budding or fission”. MORPHOLOGICAL DIVERSITY

Classical characterization of fungal diversity was mainly based on morphological characteristics from class to species level. It is surprising that the development of chemotaxonomic, and molecular techniques during last few decades rather modified but did not make too much change in the established system. In other words, the classical system developed on the basis of morphological observation retains its value. As a part of fungal kingdom, the system of yeast classification shares a similar tendency. Even though yeasts are often regarded as a simplified form of fungi, the morphology of yeasts is complex enough and could be used as a good

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A

B

Fig. 2. Transmission electron micrographs showing: (A) enteroblastic budding in Rhodotorula acuta, and (B) holoblastic budding in Saccharomyces cerevisiae [9](photo courtesy of W.H. Batenburg-Van der Vegte, Centraalbureau voor Schimmelcultures, The Netherlands).

Monopolar budding

Bipolar budding

Multilateral budding

Conidia on stalk

Fission

Pseudomycelium

Arthrospores

Endospores

Chlamydospores

Ballistoconidia

Fig. 3. Mode of vegetative reproduction in yeasts (adapted from [101] with permission from Elsevier).

tool for diversity assessment and taxonomic identification. The form of yeast cells varies from spheroidal, oval, elongated to lemon shaped, reniform and could be so characteristic that it is possible to utilize it for a precise identification. The colony appearance of yeasts ranges from the well-known white-cream in Saccharomyces cerevisiae to slimy hyaline (Lipomyces, Cryptococcus), orange (Cystofilobasidium), red (Rhodotorula, Sporobolomyces and Phaffia), brown (Metschnikowia pulcherima), greenish (Trichosporonoides) and black (Aureobasidium).

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Important morphological characteristics of yeasts include the mode of vegetative and sexual reproduction. Vegetative or asexual reproduction occurs in yeasts as budding, fission or a combination of these two processes. Budding in basidiomycetous yeasts is enteroblastic, as opposed to holoblastic in most ascomycetous yeasts [102] where all layers of the mother cell or condiogenous cell are involved in the formation of the bud. After separation of the bud, a scar remains through which no further budding occurs. During enteroblastic budding, the growing bud emerges through the mother cell wall and leaves a scar. Percurrent enteroblastic budding may result in annellations, eventually forming a collarette [101]. Both enteroblastic and holoblastic modes of budding could be judged by observation under a light microscope, but the best results could be obtained with with the use of an electron microscope (Fig. 2). The position of the budding site is also an important feature. If budding is restricted to one pole of the mother cell, it is referred to as monopolar budding and may occur on a rather broad base, for example, in Malassezia. If buds are formed at the distal poles of the mother cell, it is referred to as bipolar budding (Fig. 3). Bipolar budding is characteristic of apiculate yeasts like Hanseniaspora and Kloeckera. In most basidiomycetous yeasts, budding is limited to a restricted area at the poles of the cells, and frequently occurs on short denticles, but the cells do not become lemon-shaped. In sympodial budding mode, new buds appear just behind and adjacent to the previous budding site. Sympodial budding occurs in Sympodiomycopsis and some species of Bensingtonia and Malassezia [1, 14, 93]. Reproduction by fission implies the duplication of a vegetative cell by means of the ingrowth from cell wall of a transverse septum which bisects the long axis of the cell, being characteristic of the genus Schizosaccharomyces. Some yeasts reproduce by the formation of conidia borne on a stalk-like, tubular structure (Fig. 3). Species of Fellomyces, Kockovaella, and Kurtzmanomyces form conidia on elongated stalks with septa at the distal end. Septa in species of Sterigmatomyces and Tsuchiyaea are located at the mid-region or at varying positions, as in the case of Ballistosporomyces [72, 73, 112, 113, 114]. Some basidiomycetous yeasts are able to produce actively discharged blastoconidia (Fig. 3), often referred to as ballistospore or more precisely ballistoconidia. They are produced on sterigmata that protrude from vegetative cells and are discharged into the air by the so-called Buller’s

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drop mechanism, similar to the discharged basidiospores of certain Basidiomycetes. The difference is that ballistoconidia are produced mitotically, whereas basidiospores are meiotically formed. Ballistoconidia may be bilaterally symmetrical, for example, in the genera Kockovaella, Sporidiobolus, Sporobolomyces, and Udeniomyces, or rotationally symmetrical in Bullera and the teleomorph Bulleromyces. The presence or absence of ballistoconidia has been used to differentiate Sporobolomyces from Rhodotorula, and Bullera from Cryptococcus [30]. However, molecular data suggests that ballistoconidium-forming yeasts are closely related to nonballistoconidium-forming yeasts [74]. In addition, the formation of ballistoconidia is an easily lost characteristic and this may complicate identifications. Reniforma represents a unique mode of conidiogenesis and cell morphology. The yeast has kidney-shaped cells with a flat base circumscribed by a brim. Budding is enteroblastic and produces miniature kidney-shaped cells distinguishable from all other known yeasts [86]. Although yeasts do not reproduce by the exclusive formation of hyphae, numerous taxa under suitable conditions produce pseudohyphae or true hyphae. Pseudohyphae are cells produced by a series of buddings, each daughter cell remaining attached to its mother cell, thus forming a chain which may be branched, the cross-walls being formed by each bud that stays connected to its mother cell. By contrast, the cross-walls of true hyphae are formed centripetally in an already elongated cell [101] (Fig. 3). Pseudohyphae are not always easy to distinguish from true hyphae by casual microscopical examination. Hyphal growth is an ecological adaptation and mostly occurs when the nutrient in the medium is exhausted. For pathogenic species, filamentous growth allows yeast to invade the tissue and maintains a foothold for reinvasion. In predacious yeasts, short filamentous growths of yeasts may be used to attack other yeast species, in order to obtain supplies of organic sulphur compounds, such as methionine [65]. For basidiomycetous yeasts, hyphal growth often associates with sexual process. Successful mating triggers mycelial growth in all known basidiomycetous yeasts (Fig. 7). Hyphae may also form a clamp connection arising from outgrowths, which at cell division make a connection between two cells by fusion with the lower one. Such clamp connections are characteristic of the dikaryotic phase of basidiomycetous taxa [101]. Hyphae may break apart or disarticulate and form one-celled arthrospores or arthroconidia (Fig. 3). Frequently the arthrospores

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formed in this way on solid media are arranged in a characteristic zig-zag fashion. Arthrospores are characteristic for genera Arxula, Dipodascus, Galactomyces, Saccharomycopsis, Geothrichum, Moniliella, Trichosporon, and Trichosporonoides. In old culture, some yeasts may form asexual endospores. These endospores are vegetative cells which are delimited within cells or hyphae (Fig. 3). The formation of endospores is relatively rare and with low repeatability, and could be observed in old cultures of Trichosporon, Cryptococcus and Oosporidium. Another type of asexual specialized spore that occurs in yeasts is chlamydospore, defined as a thick-walled, non-deciduous, intercalary or terminal, asexual spore formed by the rounding off of a cell or cells (Fig. 3) [101]. Chlamydospore formation is a characteristic of Candida albicans, C. dubliniensis, some species of Cryptococcus and Metschnikowia. In Metschnikowia, chlamydospores, however, play a dual function. The germination of chlamydospores in Metschnikowia may yield a diploid yeast phase or lead to the meiotic reduction resulting in haploid ascospores (Fig. 6). ULTRASTRUCTURE OF THE CELL WALL AND HYPHAL SEPTA

The use of electron microscopy in microbiology immediately demonstrated a distinct difference between ascomycetous and basidiomycetous yeast cell wall structure [71]. With standard methods of sample preparation, it is observed that the cell wall of ascomycetous yeasts consists of the inner thick, electron transparent (bright) and thin, electron dense (dark) outer layers. On the other hand, the cell wall of basidiomycetous yeasts is laminated with many different layers (Fig. 1). Although the link between the ultrastructure of the cell wall and the reaction to diazonium blue B (DBB) is not well understood [118], it is known that all basidiomycetous yeasts give positive reaction to the dye, while ascomycetous yeasts do not. The reaction was used as an alternative to TEM observation for differentiating between these two groups of yeasts [36]. Different groups of yeasts have remarkable dissimilarities in the ultrastructure of the hyphal septa (Fig. 4). These dissimilarities relate well with other characteristics and are supported by rDNA phylogenetic

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analysis. The following types of septa are known in ascomycetous yeasts (Fig. 4-A) [101]: A1. Septa with plugged dolipore: pore with thickened edge and endoplasmic reticulum through the pore with dilations at both ends to form plugs. This type of septa is found in Ambrosiozyma (the only yeast in which dolipore could be seen under a light microscope). A2. Septa with plasmodesmata: septa with narrow connections between the protoplast of adjacent cells, as found in the yeast Saccharomycopsis capsularis. A3. Septa with central connection: it is formed by the two plasma membranes being retained in the middle of the septum when it is closed by centripetal growth, found in Arthroascus javanensis. The following types of septa are known in basidiomycetous yeasts (Fig. 4-B) [101]: B1 & B2. Septa with a dolipore. Endoplasmic reticulum may form a cap or parenthesomes at both ends of the pore. The cap may be primitive or vesicular. Dolipores with or without a cap are found in the Filobasidiaceae. B3. Septa with a simple pore with a tapered edge, as found in the teliospore forming yeasts.

A 1

2

3

1

2

3

B

Fig. 4. Schematic drawing of the septal ultrastructure of ascomycetous (A) and basidiomycetous (B) yeasts. (adapted from [101] with permission from Elsevier). A1 – plugged dolipore; A2 – plasmodesmata; A3 – closer line. B1 – dolipore with cap; B2 – dolipore with vesicular cap; B3 – simple pore.

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SEXUAL CYCLE

Characteristics of sexual life play a fundamental role in the classification of natural diversity wherever it is applicable. The separation of true fungi into Zygomycetes, Ascomycetes and Basidiomycetes was initially based on their sexual behavior and now is well supported by chemotaxonomic and phylogenetic data. Yeasts are polyphyletic with their members scattered within Ascomycetes and Basidiomycetes. In each class, yeasts’ sexual life is also dissimilar and could be used as a good tool for diversity assessment and classification. Life Cycle of Ascomycetous Yeasts

Ascosporogenous yeasts during vegetative reproduction stabilize in haploid or diploid states, or represent a mixture of both states. In homothallic haploid yeasts, the diploid state is short and exists only in the form of a zygote. Plasmogamy, karyogamy and meiosis take place in the zygote and as a result the zygote transforms into ascus containing ascospores. The ascospores germinate and the next cycle of haploid growth commences. Yeasts with this type of life cycle are referred to as haploids [51]. Formation of zygote in these yeasts could happen in different ways. The most typical way is the fusion of two independent cells, resulting in a conjugative ascus with amoeboid forms, for example, species of Schizosaccharomyces (Fig. 5-1). In another case, cells initially form outgrowths or conjugation tubes, and are fused with formation of dumbbell-shaped asci. This type of asci is characteristic for species of Torulaspora and Zygosaccharomyces. In another method of diploidization, the haploid cell first carries out mitosis and produces a special bud without separating from the mother cell. Subsequently, both nuclei of the mother cell migrate into the bud, where karyogamy and meiosis take place. Then the nuclei move back into the mother cell, which eventually transforms into an ascus containing 1 to 4 ascospores. This type of life cycle is known for species of Debaryomyces and some of Pichia (Figure 52). Species of Nadsonia and Saccharomycopsis have another characteristic method of conjugation. The dikaryotic cell in this case does not turn into ascus, instead both nuclei move into a specialized bud at the opposite side of the first bud. Here the two undergo karyogamy, meiosis and, as a result, an ascus containing one ascospore is formed (Fig. 5-3).

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1

4 K

M

A

2

M

A

M

A

K

5 K

M

A

3

K

6 K

Haploid

M

A

M

A

K

Diploid

Fig. 5. Life cycle of haploid and diploid ascomycetous yeasts (drawing courtesy of Prof. I.P. Babjeva). [119] K – karyogamy; M – meiosis; A – ascospore.

In homothallic yeasts with a stable diploid phase, the diploid vegetative cells undergo meiosis and turn into one conjugated asci. Four types of life cycles were described for these yeasts and all share very fast establishment of the diploid state after a short haploid phase. In Saccharomycodes ludwigii, ascospores conjugate directly while still being in the ascus, and the very first bud from the zygote is diploid (Fig. 5-4). Auto-diploidization is known for Hanseniaspora. Ascospore of the yeast undergoes mitosis with the formation of two nuclei and is followed by karyogamy, resulting in diploid generation (Fig. 5-5). In Saccharomyces cerevisiae, conjugation occurs shortly after a few generations of haploid growth (Fig. 5-6). In heterothallic haploid yeasts, the diploid phase can be either restricted to zygote, or prolonged with many generations of diploid growth. Heterothallic yeasts with a stable diploid state are usually heterozygous in mating type and their vegetative cells are bisexual. The life cycle of these yeasts is similar to the life cycle of homothallic yeasts with a stable diploid phase. In either case, they produce non-conjugated asci. Ascospores of compatible mating types fuse while still inside the

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Conjugation

a

Diploid phase

Karyogamy

Haploid phase a

Chlamydospore Meiosis

Ascus

Germination of ascospores Ascospores

Fig. 6. Life cycle of Metschnikowia pulcherima. (Drawing courtesy of Prof. I.P. Babjeva [119]). Yeast of the genus Metschnikowia is well-known for the formation of needle-shaped ascospores. This group of yeasts is pathogenic to invertebrates and the needle-shaped ascospores in some species could be discharged forcefully allowing active predation [64].

ascus, or shortly after some generations of haploid growth. Heterothallic cells tend to be capable of sexual agglutination. Ascospores are diverse in morphology. They can be spherical, oval, cylindrical, reniform, curved, or needle-shaped. The presence of ridges on the surface of spherical ascospores makes them Saturn or hat shaped. Ascospores may be covered with a membrane. The asci can persist or bust easily. All these characteristics are of taxonomical value. Life Cycle of Basidiomycetous Yeasts

The relationship of some non-fermenting yeasts and Basidiomycetes was first suggested by Kluyver and van Niel in 1925 when they noted the similarity of Sporobolomyces ballistoconidia to basidiospores [8]. The first direct mating proof of the basidiomycetous nature of Rhodotorula glutinis and the description of its teleomorphic state as Rhodosporidium was done by Banno in 1967 [6]. In the following years, Leucosporidium scottii (perfect state of Candida scottii) [31], Filobasidiella neoformans (perfect state of Cryptococcus neoformans) [62] were described. Most species of basidiomycetous yeasts are heterothallic with bipolar or tetrapolar mating

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Fig. 7. Sexual dimorphism in Bannoa hahajimensis [39]. After mating of two compatible strains (OK52 and OK173), the yeast changes from haploid unicellular growth to mycelial dikaryotic growth and results in the formation of pro-basidia containing diploid nucleus. Conj – conjugation between individual cells; Clamp – clamp connection; Nc – haploid nucleus; KG – nucleus after karyogamy.

types. The unicellular phase is restricted to the haploid state (Fig. 7). Most of basidiomycetous yeasts are plant parasites. Their complete life cycle requires host plants and a shift in environmental conditions. Due to the extreme difficulty in finding compatible mating types and correct conditions favoring sexual development, knowledge on the life cycle of basidiomycetous yeasts is still meager. The classification of teleomorphic basidiomycetous yeasts is mainly based on the morphology of basidia. With some generalizations, basidiomycetous yeasts could be divided into three groups: the teliospore-forming yeasts, the Filobasidiales group, and the Tremellales group. In teliospore-forming yeasts, after conjugation of haploid cells with compatible mating types, a dikaryotic (two nuclei) septed mycelium with clamp connection is formed. After some period of dikaryotic growth, teliospores are formed intercalary or terminally on the mycelia. Teliospores could be colorless or dark colored, usually rich in lipid and with thickened cell-wall. Karyogamy takes place in the teliospores.

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Species of the Filobasidiales with genus Filobasidiella, as an example, do not produce teliospores. Filobasidiella basidia are formed on septed hyphae with clamp connections. Karyogamy and meiosis take place in the basidia. Basidiospores in chain are formed at four points on the apex of basidia (Fig. 8). Species of Tremellales group also lack teliospores, but produce fruit bodies with a hymenium in which basidia are formed. Tremella – the typical genus of the group, forms gelatinous basidiocarps on wood in association (parasitic) with other basidiomycetes, for example, dacrymycetaceous fungi. The basidiocarps could have the size of several centimeters (Fig. 9). Some species of Tremella are cultivated commercially for food and medicine [5]. In culture, isolated basidiospores of Tremella give rise to the yeast state, with mycelia typically developing only when compatible strains are paired. Pairing of suitable cells yields a rapid response to complementary pheromones. As a result budding ceases, Homothallic isolates

Heterothallic isolates a

aa

Germinate into yest cells

a

Haploid budding

Conjugation

Dikaryotic mycelium

Basidial formation and sporulation Meiosis and spore formation

Basidial formation and karyogamy

Fig. 8. Life cycle of human pathogenic yeast Filobasidiella neoformans (from [61], reproduced with permission from Elsevier).

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Fig. 9. Fruiting body of Tremella mesenterica on wood (in association with Dacrymycetes). The fruiting body could have the size of several to tens of centimeters. Basidiospores collected from the fruiting body will yield haploid unicellular growth on common cultivation media for yeasts.

conjugation tubes develop and fuse, from which dikaryotic hyphal growth is initiated. Under conditions suitable for fruiting, basidiome formation starts soon after dikaryon initiation. The basidiocarp is gelatinous and 24 celled basidia are produced in an amphigenous surface hymenium. Each basidial cell produces a tubular epibasidium tipped by a sterigma and ballistospore. Ballistospore can germinate either by production of ballistoconidia or by budding [4]. CELL WALL COMPOSITION

The use of carbohydrate composition in characterization of fungi started nearly 200 years ago by the French chemist Henri Braconnot, who isolated the slimy substance of Agaricus volvaceus and compared it with that obtained from lower animals [from 106]. The comparison of the compositions of polysaccharide that make up the cell wall has great impact on the classification of yeasts [85]. Characteristics of the cell wall

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Table 1. Cell wall categories for yeast and yeast-like fungi (adapted from [106] with permission from Elsevier) Component

Family

Class

Mannan dominant Chitin low No galactose

Saccharomycetaceae Candidaceae

Hemiascomycetes

Mannan dominant Chitin intermediate Galactose

Dipodascaceae Saccharomycodaceae Candidaceae (‘chitinous yeasts’)

Hemiascomycetes

Mannan variable Chitin intermediate Fucose or rhamnose

Sporidiobolaceae, Sporobolomycetaceae

Mannan low Chitin

Ustilaginaceae

Ustilaginomycetes

Mannan low Chitin Xylose

Tremellaceae Filobasidiaceae Cryptococcaceae

Hymenomycetes

Mannan low Chitin Rhamnose

Ophiostomataceae

Euascomycetes

Urediniomycetes

composition could separate taxa at the generic and above generic levels (Table 1). Budding ascomycetous yeasts are characterized by the presence of predominant b-1,3-glucan and low proportion of highly branched b1,6-glucan. The remaining major component of the cell wall of these yeasts is a glucomannan-protein complex [85]. The cell wall composition of fission yeast Schizosaccharomyces differs significantly from that of budding ascomycetous yeasts. In addition to the b-glucans, the fission yeast cell wall contains a-1,3-glucan, which does not occur in budding ascomycetous yeasts [2]. The mannan component of the fission yeast also differs from that of the budding yeasts, i.e. it has a single galactose unit attached to the main backbone rather than the oligomannose chains [3]. The uniqueness of Schizosaccharomyces amongst ascomycetous yeasts was also confirmed by ribosomal RNA analysis [56]. The content of chitin in multilaterally budding ascomycetous yeasts usually does not exceed 2%. Chitin is confined to the bud scar areas and thus the chitin

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content is proportional to the number of buds produced. A small chitin amount (about 0.1%) is located throughout the lateral wall [70]. In yeasts of basidiomycetous affinity like Cryptococcus, Rhodotorula, and Sporobolomyces, the chitin content is significantly higher (up to 10%), where it does not concentrate in the bud scars, but is more or less evenly distributed throughout the cell wall in the form of microfibrils [10]. The presence or absence of xylose in the cell wall or whole cell hydrolysate plays an important role in discriminating basidiomycetous yeasts at generic and above generic levels. According to von Arx and Weijiman, species of heterobasidio-mycetous affinity could be divided into two families based on the presence or absence of xylose. Family Filobasidiaceae, characterized by the presence of xylose, contains: Bullera, Cryptococcus, Filobasidium, Phaffia, and Trichosporon. On the other hand, the family Sporobolomycetaceae, characterized by the absence of xylose in the cells, contains genera Rhodotorula, Rhodosporidium, and Sporobolomyces [85]. Later, Takase and his colleagues demonstrated the usefulness of cell wall composition, especially the presence or absence of xylose in combination with other characteristics like CoQ type, and septal structure for the classification of the highly heterogenous ballistospore-forming yeasts [74, 76]. G+C CONTENT

Regularities in DNA base composition were noted fifty years ago by Erwin Chargaff and his colleagues [17, 18]. The parity rule A=T, G=C played a vital role leading to the discovery of the Watson and Crick double-helix model [105]. Another important observation was that the ratio of G+C to the total bases (A+T+G+C) tends to be constant for a given species, but varies between species [18]. The base composition is rather uniform not only among organisms of the same species, but also with respect to different parts of a given DNA molecule. There are more codons than amino acids, so that most amino acids can correspond to more than one triplet codon. This gives some flexibility to the nucleic acid sequence. DNA base composition is a reflection of phylogenetic relationship. The wide range of G+C contents in different groups of yeasts thus reflects the diversity. The G+C contents of 689 yeast species included in the recent identification manual [53] ranged from 27 to 70 Mol%, which is close to the range for the fungal kingdom as a whole. With a few exceptions, yeast species with ascomycetes affinity have G+C

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value less than 50%, while species of basidiomycetes affinity have the G+C value above 50%. The strains with G+C contents differ from each other by 2-2.5% and usually belong to a separate species. The identity in G+C values, however, does not necessarily indicate a close relationship [58]. The range of G+C content of species within a genus is often 10% or less, as found in Debaryomyces, Hanseniaspora, Issatchenkia, Kluyveromyces, Metschnikowia and several other genera. Genera showing G+C range among species of greater than 10% may be polyphyletic, but a narrow range does not ensure monophyly [58]. In some groups of yeasts, the G+C contents are not evenly distributed throughout the genome. It was shown that some parts of genomic DNA in Geothrichum clavatum differ from other parts by roughly 12% [91]. So far there is no explanation for this phenomenon. The classical methods for determination of G+C contents are CsCl equilibrium density generated by ultra-centrifugation [32], enzymatic degradation and subsequent HPLC analysis [38], and spectroscopic methods [66]. A simple and rapid method to determine the G+C content, based on the calculation of melting temperature using a real-time thermocycler, utilizes the intercalating dye Sybr Green I, which binds to the double stranded DNA and gets released during the melting course, and thus changes the fluorescence intensity [109]. CoQ TYPE

The ubiquinones, (coenzyme Q) first described and named by Crane et al. [22, 23], are important carriers in the electron transport chain of the respiratory systems. The number of isoprene units attached to the quinine nucleus and saturation degree of the isoprene chain varies, but in each species only one type of ubiquinone dominates. Usually the domination is over 90% of the total ubiquinonologous and the abundant compound is regarded as the major ubiquinone type. The biochemical background of the phenomenon remains largely unresolved, but it is assumed that the chain lengths depend on the product specificity of trans-long-chain prenyl diphosphate synthases that yield their precursors and the specificity of the trans-membrane transporter [41, 98]. The variation in ubiquinone type was utilized extensively for the circumscription and identification of genera in bacterial taxonomy [21, 46, 82, 110]. For yeasts, ubiquinone systems have been adopted as a

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Yeast genera and coenzyme Q type presence

Genera

Coenzyme type

Arxiozyma, Cyniclomyces, Hanseniaspora, Kluyveromyces, Nadsonia, Saccharomyces, Saccharomycodes, Torulaspora, Zygosaccharomyces

6

Ambrosiozyma, Issatchenkia, Saturnispora, Williopsis

7

Citeromyces, Clavispora, Cystofilobasidium, Mrakia, Pachysolen, Saccharomycopsis

8

Aciculoconidium, Arxula, Babjevia, Bensingtonia, Brettanomyces, Debaryomyces, Dekkera, Dipodascopsis, Dipodascus, Hyalodendron, Itersonilia, Lodderomyces, Metschnikowia, Sporopachydermia, Stephanoascus, Sterigmatomyces, Sympodiomyces, Trichosporonoides, Tsuchiyaea, Wickerhamia, Yarrowia, Zygoascus

9

Bullera, Bulleromyces, Chionosphaera, Fellomyces, Filobasidiella, Kockovaella, Kurtzmanomyces, Leucosporidium, Protomyces, Pseudozyma, Saitoella, Sporidiobolus, Sterigmatosporidium, Sympodiomycopsis, Tilletiaria, Tilletiopsis, Ustilago, Xanthophylomyces

10

Aureobasidium, Erythrobasidium

10(H2)

Myxozyma, Zygozyma

8, 9

Filobasidium, Lipomyces, Rhodosporidium, Rhodotorula, Schizosaccharomyces, Trichosporon

9, 10

Eremothecium Sporobolomyces

6, 7, 9 10, 10(H2)

Pichia

7, 8, 9

Cryptococcus

8, 9, 10

Candida

6, 7, 8, 9

useful taxonomic criterion at generic level [75, 92, 111, 117]. To date, the following coenzyme types are known in yeasts: CoQ6, CoQ7, CoQ8, CoQ9, CoQ10, CoQ10(H2) (see Table 2). Ubiquinone type is genericspecific and its variation within genus often indicates a cumbersome nature, and the genus may require revision. Typical examples are genera like Pichia, Cryptococcus, and Candida. The heterogeneity of these genera was confirmed by morphological, biochemical, and rDNA sequencing data. The method for determining ubiquinone type for yeasts, described in detail by Yamada and Kondo [117] is relatively simple and accessible by most laboratories.

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YEAST DIVERSITY AND EVOLUTION AS REVEALED BY RNA/DNA SEQUENCING

Sequences of ribosomal RNA (rRNA) and its coding regions (rDNA) have been used extensively during the last decade for assessment of diversity and relationship among organisms. The usefulness of rRNA/ rDNA sequence comparison comes from two important properties: (1) ribosomes are present in all cellular organisms and appear to share a common evolutionary origin, thus providing molecular history shared by all organisms; (2) some rRNA/rDNA sequence are sufficiently conserved for all organisms, thus allowing alignment for comparison of less conserved areas used to measure evolutionary relationship [52]. Ribosomal RNAs in eukaryotes have several size classes. The genes coding for large (25S to 28S), small (18S) and 5.8S rRNAs are located in tandem repeats as many as 100 to 200 copies. The separately transcribed 5S rRNA gene may also be included in the repeat [52]. The sequence comparison of gene coding for the above mentioned classes of rRNA plays a fundamental and leading role in modern classification of yeasts. 5S rRNA

In early phylogenetic studies, 5S rRNA was chosen for its conserved nature and small size (about 120 nucleotides). The small size allows easy sequence determination and the conserved nature favors assessment of broad phylogenetic relationships [42, 52]. In basidiomycetous yeasts and filamentous fungi, the similarity in 5S rRNA correlates with the septal structure of these organisms [52]. Also, based on the 5S rRNA, the relatedness of Rhodosporidium malvinellum and Rhodosporidium toruloides was questioned and with supports by other phylogenetic data, a separate genus Kondoa was installed for R. malvinellum [116]. Analyzing 5S rRNA sequences prompted the division of the ascomycetes among three groups: (1) Schizosaccharomyces and Protomyces, (2) budding yeasts, and (3) filamentous fungi [103, 104]. Despite the impact of the 5S rRNA phylogenetic studies on the earlier understanding of the fungal diversity, its role is now replaced by the informationally richer 18S and 26S rDNA sequences [52].

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5.8S rRNA

Direct sequencing of 5.8S rRNA was more difficult than that of the 5S rRNA due to the presence of modified nucleotides. With the size of about 160 nucleotides, the information provided by the 5.8S sequence is not much more than that by the 5S rRNA. For those reasons, there were not many phylogenetic studies based on the 5.8S sequences [52]. Recently, a rather large database of 5.8S rDNA sequence was established as the flanking ITS regions were of interest for species identification. So far, the use of 5.8S rDNA coding region for phylogenetic assessment remains limited. 18S rDNA

With the advancement of sequencing technology and the demand for greater resolving power, taxonomists began to use 18S rDNA sequence for phylogenetic assessment. The 18S rRNA coding region is large enough and contains very conserved regions as well as divergent parts. Sequencing of 18S rDNA allows estimation of the relationships of organisms from phylum down to the species level. Possibility of aligning makes it possible to put all living organisms in one phylogenetic tree [83]. In 1992, Berbee and Taylor analyzed a large set of 18S rDNA sequences from Ascomycetes, Basidiomycetes, Zygomycetes, and Chytridiomycetes, in combination with data of fungal fossil records and deduced the timescaled evolutionary history of the true fungi [12]. According to this, Ascomycetes fall phylogenetically into three groups (Archiascomycetes, Hemiascomycetes, and Euascomycetes) established during the Carboniferous era, from 330 to 310 million years ago (Ma). The “true yeast” (Hemiascomycetes) diverged from filamentous Ascomycetes about 310 Ma. The first true yeasts were filamentous, giving rise to Dipodascopsis uninucleata and Endomyces geotrichum. The lineage of unicellular yeasts, represented by Kluyveromyces lactis and Saccharomyces cerevisiae, diverged from the filamentous type roughly 240 Ma. In turn, K. lactis and S. cerevisiae were derived from a common ancestor around 80 Ma during the Cretaceous period. The first comprehensive phylogenetic placement of yeasts among higher fungi was inferred from 18S rDNA sequences by Wilmotte and associates [107]. The separation of Ascomycetes into three lineages was supported, and the xylose negative basidiomycetous yeasts formed a distinctive branch apart from

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Fig. 10. Placement of representative yeasts among higher fungi as inferred from 18S rDNA sequences (adapted from [107] with permission from Elsevier). Ascomycetous yeasts are presented in blue color while basidiomycetous yeasts are displayed in red.

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xylose containing basidiomycetes (Fig. 10). The sequencing of 18S rDNA was used widely for assessment of relationships among ascomycetous yeasts [15, 44, 77] as well as basidiomycetous yeasts [35, 96, 115]. It was especially useful in combination with other characteristics like GC content, CoQ type and cell wall composition for the classification of ballistoconidium-forming yeasts [37, 76, 94]. The usefulness of 18S rDNA sequences, however, is hampered by the incompleteness of available database. With current sequencing technology, the complete 18S rDNA of around 1,800 nucleotides still remains relatively large to be used for routine species identification. 26S rDNA

A recent achievement in studying biodiversity of yeasts was the establishment of a database of 26S rDNA D1/D2 sequences for all known species of yeasts. The completeness of the database makes it possible to compare any new isolate with all known species, and thus any potentially new species could be detected easily. This has accelerated the pace of biodiversity exploration, as witnessed during the last few years. Major part of this database was contributed by Kurtzman and Robnett [54, 55] for ascomycetous yeasts, and of Scorzetti et al. [87] for basidiomycetous yeasts. The D1/D2 part of the 26S rDNA was chosen for it’s relatively short (about 600 nucleotides) and divergent nature, which is suitable for generic assessment and routine species identification. These extensive sequencing projects substantially restructured the system of yeast taxonomy by pointing out the potentially synonymous species, the relationship between teleomorphic and anamorphic taxa, and the heterogeneity of some genera. In combination with previous nDNA reassociation, mating compatibility studies, Kurtzman and Robnett drew up following conclusions: (1) strains showing greater than 1% substitutions in the ca. 600-nucleotide D1/D2 domain are likely to be different species, and (2) strains with 0–3 nucleotide differences are either conspecific or sister species [54]. While these conclusions are helpful, their usage should be accompanied with precaution. Recently, Lachance et al. [63] examined interbreeding members of the yeast species Clavispora lusitaniae and found that mating compatible strains might be different for more than 6% in the 26S rDNA D1/D2.

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Other House-keeping Genes

Besides rDNA, other genes also were shown to be useful in characterization of yeast diversity. Daniel et al. [24] employed the actin gene as a phylogenetic marker for selected members of the anamorphic genus Candida and seven related teleomorphic genera. The major clustering inferred from the actin gene sequences corresponded to the one by rDNA analysis. The actin gene offers a particular advantage of an unambiguous alignment. In some cases where higher resolution power is required, DNA topoisomerase II gene (known as DNA gyrase gene, which is utilized extensively for characterization of prokaryotes) could be exploited. The gene was useful for delimitation species and strains of pathogenic Candida [47]. Divergent histone promoter regions flanked by highly conserved sequences could also be utilized for species identification [11]. Recently Kurtzman and Robnett analyzed 75 species of “Saccharomyces complex” using an extensive list of genes including rDNA repeat (18S, 26S, ITS), nuclear genes (translation elongation factor 1a, actin-1, RNA polymerase II) and mitochondrially encoded genes (small subunit rDNA, cytochrome oxidase II) and showed that the multi-gene approach enables a better resolution of previously poorly supported clades [57]. As a result, species of the historically long-standing genera like Saccharomyces, Kluyveromyces and some other members of the Saccharomycetaceae were circumscribed into 5 new genera Lachancea, Nakaseomyces, Naumovia, Vanderwaltozyma, and Zygotorulaspora [60]. This is seemingly a biologically correct rearrangement but it has been met with confusion and criticism due to the importance of the revised genera in the accumulated literature [9]. Sequencing of the whole genome

Ideally, the best way to characterize yeast is to sequence its whole genome. In 1989, Andre Goffeau set up a European consortium to sequence the genome of the budding yeast Saccharomyces cerevisiae. The project was historical since it was the first eukaryotic organism to be sequenced and the sequence was approximately 60 times larger than any sequence previously attempted. Sanger’s “shotgun” method was utilized as standard for S. cerevisiae genome sequencing. In 1996, after six years of intensive work with the involvement of 600 scientists in Europe,

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North America, and Japan, the project was completed [34]. The genetic information of S. cerevisiae is distributed in 16 distinct chromosomes with a total of 13.4 megabases (Mb). A total of 5,855 open reading frames (ORFs) were predicted and almost 70% of the total sequence consists of ORFs. The yeast genome contains approximately. 140 ribosomal RNA genes in a large tandem array on chromosome XII and 40 genes encoding small nuclear RNAs (snRNAs) scattered throughout the 16 chromosomes. About one-third of the sequenced genes were of unknown function since the translated product lacked significant homology to any other protein with known function. The genes are regarded as ‘orphan’ genes. Genetic redundancy was noted in S. cerevisiae, and it was considered as the source for functional diversity and adaptation [40]. Fission yeast Schizosaccharomyces pombe was the next yeast whose genome was fully sequenced [108]. Fission yeast is the second in term of biotechnological importance after S. cerevisiae. It is widely used for fermentation in countries with warm weather, due to its optimum growth at relatively high temperature (30-42°C). The biology and genetic background of fission yeast are distant from all Hemiascomycetes in general and S. cerevisiae in particular. The whole genome of S. pombe is 13.8 Mb, slightly larger than that of S. cerevisiae. Substantial differences exist between budding and fission yeast. Genome of S. pombe consists of three chromosomes (instead of 16 as in S. cerevisiae), whereas only 5% of the S. cerevisiae genes have predicted introns, in S. pombe 4,730 introns have been characterized and confirmed in 43% of the genes. With 60.2% genome codifies protein, the genome of S. pombe is less compact than the one of S. cerevisiae (70% genome codifies protein). Only two-thirds of predicted proteins in S. pombe have homologues in S. cerevisiae [40]. Genome sequencing of an important human pathogen Candida albicans was completed in 2004 [45]. Because C. albicans has no known haploid or homozygous form, sequencing was performed as a wholegenome shotgun of the heterozygous diploid genome in a clinical isolate. The known diploid form of C. albicans has eight sets of chromosome pairs. Its genome size is about 16 Mb (haploid), about 30% greater than S. cerevisiae. The reference haploid genome contains 6,419 ORFs [45]. Recently the Génolevures Consortium consisting of 14 French institutes and laboratories has engaged in an ambitious project of sequencing 14 Hemiascomycetes yeasts. Until January 2004, the project has completed genome sequencing for four yeasts: Candida glabrata (13

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chromosomes, total 12.3Mb without the rDNA, 5,283 ORFs), Kluyveromyces lactis (6 chromosomes, total 10.6 Mb, 5,329 ORFs), Debaryomyces hansenii (7 chromosomes, 12.2 Mb, 6,906 ORFs), and Yarrowia lipolytica (6 chromosomes, 20.5 Mb, 6,703 ORFs) [28]. This effort has totaled the number of all fully sequenced yeasts to seven. The sequencing of following yeasts is underway: S. bayanus, S. servazzii, Zygosaccharomyces rouxii, S. kluyveri, Kluyveromyces thermotolerans, K. lactis, K. marxianus, Pichia angusta, P. sorbitophila, and Candida tropicalis. Besides that, partially completed genome sequencing of six yeasts (S. paradoxus, S. mikatae [49], K. walti [48], S. kudriavzevi, S. castellii [20], and Ashbya gossypii [27]) was done by various other laboratories. Genome sequencing and the theory of whole genome duplication

Perhaps the biggest impact of yeast genome sequencing on understanding the evolution of eukaryotes is the ultimate proof of the theory of whole genome duplication. According to Ohno [81] if evolution had been entirely dependent upon natural selection, from bacterium only numerous forms of bacteria would have emerged. Big leaps in evolution would require a large gene pool with previously no existing functions. He suggested that evolution is supported by tandem gene duplication, and more importantly—by whole genome duplication. The newly created gene pool freed from previous tasks could evolve for acquisition of new functions, which would eventually facilitate the host in the process of adaptation and occupation of a new ecological niche. Indeed, most genes are not unique, but are part of larger families of related genes. These gene families are a result of duplication of an ancestral gene and this could happen many times in the past. While the idea on local duplication was generally accepted, whole genome duplication was a subject of long standing debate. Logically, the proof for the whole genome duplication could be found by tracing the “fossilized” patterns of the genome sequences obtained from organisms of ancestral lineage and the lineage where the duplication event took place. In 2004, convincing evidence of genome duplication was found by comparing the genome sequences of Kluyveromyces waltii [48], Ashbya gossypii [27] to the one of S. cerevisiae. In both cases, the genome sequences were computed and divided into blocks with homology in gene sequence and gene order on the chromosome syntenies. Genomes of different organisms were compared

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based on these blocks (Fig. 11). The evidence was striking. Genomes of K. waltii and S. cerevisiae are related by a 1:2 mapping, with each region of K. waltii corresponding to two regions of S. cerevisiae, as expected for whole-genome duplication. When the duplicated genes were compared with the ancestral gene, it appeared that 95% of accelerated evolution involves only one member of a gene pair, and thus exactly as predicted by the initial idea of whole genome duplication. After duplication, rearrangement of genes plays an important role in the process of evolution and speciation. It is known that species in the Saccharomyces sensu stricto are closely related but genetically isolated. The mating among them generally yields sterile hybrids. The presence of reciprocal chromosomal translocations in different species was thought to play a role in reproductive isolation and speciation. Recently Delneri et al. [26] have attempted to “play back” evolution by reconfiguring the Saccharomyces cerevisiae genome so that it is colinear with that of S. mikatae, and demonstrated that this imposed genomic colinearity allows the generation of sexually fertile interspecific hybrids. Thus, it is now generally accepted that genome duplication, massive gene loss, and chromosome rearrangement played an important role in the evolution of eukaryotes in general and yeasts in particular. It is unknown, however, how many times the whole genome duplication actually took place and how large was the impact of those events on the course of evolution. From a biological comparative point of view, the difference between K. waltii and S. cerevisiae may be not that large, at least, not as large as one would expect for 100 million years of evolution after the duplication event. OVERVIEW OF IMPORTANT GROUPS OF YEASTS

According to the recently accepted system [53], yeasts are placed under Ascomycota and Basidiomycota. The hierarchy range varies in different systems. An overview of important groups of yeasts is given below: Ascomycetous yeasts

Ascomycetous yeast group consists of representatives from three classes: Archiascomycetes, Euascomycetes, and Hemiascomycetes (Table 3). The class Archiascomycetes was proposed by Nishida and Sugiyama in 1994 [78] to accommodate genera Schizosaccharomyces, Taphrina, Protomyces,

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Fig. 11. Schematic presentation of whole genome duplication event and supported evidence [from 48]. After duplication event (a) and massive gene loss (b) only some fractions of duplicated genes retained (c) the homology of which could be detected when looking at the genome of S. cerevisiae (d). By comparing genomes of K. waltii and S. cerevisiae the duplication evidence could be seen through the retained groups of genes with homology in sequence and order (e).

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Table 3. Classification of the ascomycetous yeasts according to Kurtzman C.P. [59] (reproduced with permission from Elsevier) Class Order Family Genus Phylum: Ascomycota “Archiascomycetes” Schizosaccharomycetales Schizosaccharomycetaceae Schizosaccharomyces Taphrinales Taphrinaceae Taphrina Lalaria Protomycetales Protomycetaceae Protomyces Saitoella Pneumocystidales Pneumocystidaceae Pneumocystis Euascomycetes Endomyces Oosporidium Hemiascomycetes Saccharomycetales (syn Endomycetales) Ascoideaceae Ascoidea Cephaloascaceae Cephaloascus Dipodascaceae Dipodascus Galactomyces Sporopachydermia Stephanoascus Wickerhamiella Yarrowia

Family Genus Zygoascus Endomycetaceae Endomyces Helicogonium Myriogonium Phialoascus Trichomonascus Eremotheciaceae Eremothecium Coccidiascus Lipomycetaceae Babjevia Dipodascopsis Lipomyces Zygozyma Metschnikowiaceae Clavispora Metschnikowia Saccharomycetaceae Arxiozyma Citeromyces Cyniclomyces Debaryomyces Dekkera Issatchenkia Kluyveromyces Lodderomyces Pachysolen Pichia Saccharomyces

Table 3 Contd.

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Table 3 Contd. Saturnispora Torulaspora Williopsis Zygosaccharomyces Saccharomycodaceae Hanseniaspora Nadsonia Saccharomycodes Wickerhamia Saccharomycopsidaceae Ambrosiozyma Saccharomycopsis

Candidaceae (anamorphic) Aciculoconidium Arxula Blastobotrys Botryozyma Brettanomyces Candida Geotrichum Kloeckera Myxozyma Schizoblastosporion Sympodiomyces Trigonopsis

Saitoella, and Pneumocystis. This major lineage represents the earliest diverging ascomycete lineage prior to the separation of two other major lineages Hemiascomycetes and Euascomycetes. Archiascomycete genera share some principal characteristics with both ascomycetous and basidiomycetous yeasts, such as the cell wall ultrastructure, biochemical characteristics, mode of conidium ontogeny and major ubiquinone system. Members of Archiascomycetes lack ascogenous hyphae and ascomata. They are morphologically so diverse that it is impossible to give common morphological characteristics for the class [88]. The most economically important member of the class is Schizosaccharomyces pombe. This yeast was isolated in 1890 from the East African millet beer, called “pombe”. Pombe was made and widely consumed in Africa [7]. S. pombe can only very rarely be isolated from the natural environment and was thus thought to be domesticated by man during the history of preparation of alcoholic drinks in tropical countries, similar to that of S. cerevisiae in temperate and subtropical zone. Species of the genus Taphrina are parasitic on a wide variety of vascular plants, primarily on ferns, the Rosales, and Fagales [88]. In parasitic phase, Taphrina forms intra- or subcuticular dikaryotic mycelia and produces naked asci. They form yeast growth in saprobic, haploid phase. The anamorphic state of Taphrina is Lalaria. Protomyces is also a plant pathogen with a more restricted host range than Taphrina. Infection by Protomyces is often associated with gall and lesion formation and color change in the host.

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Pneumocystis carinii is a major causal agent of pneumonia in immunocompromised patients [88]. Saitoella is an anamorphic genus and formerly placed under Basidiomycota. It was recognized only recently as a member of Ascomycota, mainly by the analysis of 18S rDNA [77]. The placement of Endomyces under Euascomycetes or Hemiascomycetes is uncertain. All known species of the genus are obligate parasites on mushroom. On a natural substrate, they produce white, floccose colonies with pseudo-mycelium or true hyphae. They form asci containing 2-12 helmet-shaped ascospores and open by rupture at the apex [25]. Another genus in the Euascomycetes is Oosporidium. This genus with single species O. margaritiferum is unusual for ascomycetous yeasts by the presence of orange to red pigments, which is a usual feature of basidiomycetous yeasts. The pigments are, however, not of carotenoid nature. This anamorphic genus is also characterized by intense production of endospores in old culture [90]. An intriguing group of symbiotic yeasts from aphids (Homoptera: Aphididae) and planthoppers (Homoptera: Delphacidae) could also be placed under Euascomycetes. It is ascertained that most of the 4,400 known species of aphids harbor prokaryotic intracellular symbionts in the cytoplasm of mycetocytes, huge cells in the abdomen specialized for this purpose. However, some Cerataphidini aphids do not harbor them but possess yeastlike extracellular symbionts in the abdominal hemocoel [33]. The rice planthoppers, Nilaparvata lugens, Sogatella furcifera, and Laodelphax striatellus, on other hand harbor yeast-like endosymbionts in the fat body, and transmit them to the next generation through the ovary [19, 79, 80]. In both cases the yeasts cannot be cultivated and were characterized based on rDNA. The phylogenetic study indicated that the yeasts are closely related and belong to the entomopathogenic group of pyrenomycetes fungi [33, 80, 95]. Hemiascomycetes are often regarded as “true yeasts”. Yeasts of Hemiascomycetes are characterized by the absence of ascocarps and asci developed directly from the cell or mycelia. The class contains 11 families with 43 teleomorphic and 12 anamorphic genera with approximately 460 species. The largest teleomorphic genus is Pichia with 91 species. This genus characterized for the production of hat-shaped ascospores, is, however, considered to be polyphyletic but awaits revision. Relatively large and well-known teleomorphic genera of Hemiascomycetes include Debaryomyces (15 species), Kluyveromyces (15 species), Saccharomyces

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(14 species), Dipodascus (13 species), Metschnikowia (10 species), Saccharomycopsis (10 species), and Zygosaccharomyces (9 species). Among anamorphic genera, Candida is the largest and well-known to be polyphyletic with 163 species. The next relatively large genus is Geotrichum with 11 species. Belonging to Hemiascomycetes are the yeast species of most economic and scientific interest—the baker’s yeast S. cerevisiae, the species widely used in fermentation—Kluyveromyces marxianus, the human pathogen Candida albicans, and Pichia pastoris - the yeast nowadays commonly used host for expression of heterologous protein. Most species of the Hemiascomycetes are vectored or associated with animals, mainly insects. Recently a new species has been discovered Debaryomyces mycophilus, from the gut of a fungus eating wood louse. The yeast was so adapted to this association that it could not grow in culture. Later, it was confirmed that D. mycophilus is depleted in iron and needs exogenous sideropheres produced by fungi for iron acquisition [99]. Species Cyniclomyces guttulatus is specialized with the living environment inside the gut of rabbits and can be cultured only in the presence of increased level of CO2 and temperature between 30-40°C [84]. Species in the genus Metschnikowia are characterized by the formation of needle-shaped ascospores; the latter are believed to help the yeast cross through epithelial cells of insect and thus facilitate parasitism. Similar to the endosymbiotic yeasts of Euascomycetes, it is believed that many yeasts of Hemiascomycetes are also obligate symbionts and thus escape detection. So far only two unculturable yeasts of Hemiascomycetes are known: Coccidiascus legeri, an intracellular symbiont in the epithelial cells of fruit fly [29], and Macrorhabdus ornithogaster, the yeast that colonizes the gastric isthmus of many species of birds [100]. Coccidiascus legeri was discovered in 1913, however its placement in Hemiascomycetes was based solely on morphological characteristics and should be verified with more reliable phylogenetic analysis. The yeast Macrorhabdus ornithogaster was mistakenly referred to as unculturable “megabacterium” and its true nature was revealed only recently. Phylogenetic analysis based on 18S and 26S rDNA sequences indicates that M. ornithogaster is close to Dipodascus and Metschnikowia [100]. Yeasts of Hemiascomycetes are also known to be parasitic on plants as in the case of Eremothecium. E. ashbyi and E. gossypii are often used for industrial production of riboflavin. Recently an interesting genus

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Ascobotryozyma [50] (with anamorphic state known as Botryozyma [89]) was described. It is vectored by nematodes through a haustoria-like structure that helps them to attach to the host cuticles. A wide perception is that yeasts occur only in sugar-rich environments. However, some yeasts are known to live only in soil. Species of the genus Lipomyces are typical soil inhabitants. The yeast was isolated by Starkey in 1946 by chance when he was trying to isolate Azotobacter on a nitrogen-free medium. The yeast could utilize trace amounts of nitrogen in the medium or ammonium from the air for growth, and thus was mistakenly assigned as capable of nitrogen fixation, the property which is unique for the prokaryote world. Lipomycetes yeasts produce large amounts of intracellular lipids, various amylases, and are of biotechnological and bio-medical application potential. Indeed, vast majority of known yeasts from Hemiascomycetes are free living saprophytes on sugar-rich environments. These yeasts are often vectored or associated with insects. The typical genera are Saccharomyces, Zygosaccharomyces, Kluyveromyces, Pichia, and Debaryomyces. It is most likely that the Hemiascomycetes was formed as the result of co-evolution with insects and the class flourished during the Cretaceous era along with the appearance of flowering plants and the diversification of insect world. Basidiomycetous yeasts

The biology of basidiomycetous yeasts is very different from that of ascomycetous yeasts. If the term “yeast” often associates with fermentation activity, then it is worth noting that among approx. 220 species of basidiomycetous yeasts, only a few are able to ferment sugars. It is believed that only about 1% of the species of basidiomycetous yeasts in nature have been collected and described [87]. The classification of Basidiomycetous yeasts into higher taxonomic ranks is mainly based on characteristics like septal structure, basidium morphology and cell wall composition. These morpho-chemical characteristics are generally in agreement with phylogenetic analysis based on rDNA sequencing. In some systems of classification, all basidiomycetous yeasts are placed under three classes: Ustilaginomycetes, Urediniomycetes, and Hymenomycetes [97]. Class Ustilaginomycetes is characterized by the presence of a septal structure with a primitive dolipore, the presence of glucose as a major component of cell wall, and the absence of xylose in

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the cell wall. Species in the class Urediniomycetes have septa with simple pore structure and cell wall with mannose as a major component. Xylose is absent in the cell wall of Urediniomycetes. Species of Hymenomycetes differ from the Ustilaginomycetes and Urediniomycetes by the presence of dolipore and the presence of xylose in the cell wall. In the classification system of basidiomycetous yeasts presented in a recent identification manual [53], weights were given mainly on morphological characteristics (Table 4.). The highest taxonomic ranks are separated solely based on septal pore structure and the morphology of basidia. The treatment of lower taxonomic ranks, however, is similar in different systems. The classification of basidiomycetous yeasts is further complicated since many of them share a very short stage of unicellular growth. These organisms are better known in multicellular forms. One important aspect of basidiomycetous yeast biology is parasitism on plants, fungi, and animals. The most distinct among them is Cryptococcus neoformans, the teleomorphic state of which is known as Filobasidiella neoformans. C. neoformans is the causative agent of cryptococcosis. Given the neurotropic nature of the fungus, the most common clinical form of cryptococcosis is meningoencephalitis. The course of the infection is usually subacute or chronic. Cryptococcosis may also involve the skin, lungs, prostate gland, urinary tract, eyes, myocardium, bones, and joints. In AIDS, cryptococcosis is the fourth most important cause of death due to infectious diseases [43]. C. neoformans has a worldwide distribution, and two varieties of the fungus are known, namely C. neoformans var. neoformans and C. neoformans var. gattii. C. neoformans var. neoformans is found in both temperate and tropical climates in association with avian habitats, especially pigeons. C. neoformans var. gattii is most often found in tropical or subtropical locations and frequently isolated from debris of Eucalyptus. Malassezia spp. are lipophilic yeasts found on skin and body surfaces of humans and animals. Malassezia is a member of the normal skin flora and may occasionally cause superficial and deep mycoses. The teleomorphic state of Malassezia is not known. Malassezia furfur and Malassezia pachydermatis are the most common and well-studied. Malassezia infections are mostly endogenous and originate from the colonized skin. They may occur in otherwise healthy individuals as well as immunocompromised hosts, such as bone marrow transplant recipients and patients with cancer or AIDS [43].

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Table 4. Classification of the basidiomycetous yeasts according to T. Boekhout et al. [13] (reproduced with permission from Elsevier) Characteristic Order Family Genus

Characteristic Order Family Genus

Teleomorphic taxa Auriculoscypha 1. With “simple” septal pores Coccidiodictyon A. Basidia cylindric, transversely septate Ordonia Ustilaginales Septobasidium Ustilaginaceae Atractiellales Microbotryum Chionosphaeraceae Schizonella Chionosphaera Sorosporium Stilbum Sphacelotheca Atractogloeaceae Sporisorium Atractogloea Ustilago Agaricostilbales Ustilentyloma Agaricostilbaceae Sporidiales Agaricostilbum Sporidiobolaceae B. Basidia globose, none septate Leucosporidium Cryptobasidiales Rhodosporidium Cryptobasidiaceae Sporidiobolus Coniodictyum Erythrobasidium Cryptobasidium Kondoa Microstromaceae Sakaguchia Microstroma Platygloeales Exobasidiales Cystobasidiaceae Exobasidiaceae Colacogloea Brachybasidium Cystobasidium Dicellomyces Kriegeria Exobasidiellum Mycogloea Exobasidium Occultifer Laurobasidium Tijbodasia Septobasidiales Septobasidiaceae Table 4 Contd.

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Table 4 Contd. 2. With dolipore septa, parenthesomes cupulate A. Basidia “cruciate- septate” Tremellales Sirobasidiaceae Fibulobasidium Sirobasidium Tremellaceae Bulleromyces Itersonilia Holtermannia Phyllogloea Sirotrema Tremella Trimorphomyces B. Basidia aseptate Filobasidiales Filobasidiaceae Cystofilobasidium Filobasidiella Mrakia

Xanthophylomyces Syzygosporaceae Christiansenia Syzygospora Anamorphic taxa Sporobolomycetaceae Bensingtonia Kurtzmanomyces Rhodotorula Sporobolomyces Sterigmatomyces Cryptococcaceae Bullera Cryptococcus Fellomyces Kockovaella Phaffia Trichosporon Tsuchiyaea Udeniomyces Hyalodendron Moniliella

Trichosporon beigelii is also considered to be a human pathogen. This species is primarily responsible for a disease of hair shafts called white piedra. T. beigelii could be found widely in soil, plant litter and various other substrates. T. beigelii is known to cause a number of conditions in addition to white piedra, which include summer-type hypersensitivity pneumonitis, mucosal infections, and deeply invasive infections including fungemia, single organ infection, and widely disseminated infections. Invasive infections occur in immunosuppressed patients [43]. Although the majority of basidiomycetous yeasts are thought to be plant pathogens, the direct involvement of yeasts in plant diseases remains largely unknown. Ballistoconidium-forming yeasts are found basically on the surface of any plant but rarely associate with any obvious symptom. The lack of knowledge in this field may be due to the dimorphic nature of basidiomycetous yeasts. It is believed that plant

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parasitism is associated with dikaryotic growth and during this phase the requirement for the host could be of obligate nature. Some fungal associated species are assumed to be mycoparasitic since the formation of haustoria and colacosomes is common for them. Many basidiomycetous yeasts are of biotechnological importance and of great potential. Phaffia rhodozyma (teleomorph Xanthophyllomyces dendrorhous) is a biological source for astaxanthin, an economically important pigment used in aquaculture. The yeast could be found in tree exudates in countries with temperate climate. P. rhodozyma is among the rare basidiomycetous yeasts that can ferment sugars. It is well known that some species of Cryptococcus, Rhodotorula, and Trichosporon could accumulate a large amount of lipid (up to 65% of dry weight) and are considered as good candidates for microbial production of lipids. Basidiomycetous yeasts in general have wider assimilation profile for exogenous carbon sources than ascomycetous yeasts. Many are known to produce xylanases (Cryptococcus albidus), cellulases (Cryptococcus cellulolyticus) and amylases (Cryptococcus spp., Trichosporon spp.). Species of Pseudozyma produce mannosylerythritol lipids and beta-lipase for the synthesis of glucoside esters exhibiting surfactant properties [30]. Species of Rhodotorula, Cryptococcus, and Trichosporon are capable of growing on terpenoids, and aromatic compounds, and produce a range of intermediate products. Due to the high stereo- and regio-selectivity, these biotransformation reactions could be successfully utilized for synthesis of otherwise difficult-to-obtain organic compounds. CONCLUSION

Currently, around 700 species of yeasts are known. The list is expanding rapidly with approximately. 50 new species being added every year. Recent advances in understanding of the diverse world of yeasts are characterized by a polyphasic approach that combines traditional morpho-physiological with chemotaxonomic and phylogenetic studies. The establishment of the reference rDNA database for all known species has accelerated the pace of discovering natural diversity. Enormous amounts of information derived from genomic sequencing activities triggered unprecedented growth of bioinformatics and molecular evolution studies. Bioinformatics has changed the nature of evolution studies from mainly speculative science into solid fact-based science. The ultimate proof of whole genome duplication theory is an example.

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Most of our knowledge about yeast biology is so far restricted to the behavior of pure cultures in artificial media. Little is known about the life of yeasts in natural environment. Living forms exist in nature in interaction and they co-evolve. Many species are obligated to the life in interaction and thus cannot be cultivated in isolation from their natural partners and would, therefore, easily escape our detection. It is estimated that we know only about 1%, or at max 5%, of yeasts that do exist in nature. Moreover, among 700 known species, only less than 10 are being employed in different biotechnological processes. The vast potential of yeasts still awaits further exploration. References [1] Ahearn DG, Simmons RB. Malassezia Baillon. In: Kurtzman CP, Fell JW, eds. The yeasts, a taxonomic study. Amsterdam, Netherlands: Elsevier, 1998: 782–784. [2] Bacon JS, Jones D, Farmer VC, Webley DM. The occurrence of alpha (1-3) glucan in Cryptococcus, Schizosaccharomyces and Polyporus species, and its hydrolysis by a Streptomyces culture filtrate lysing cell walls of Cryptococcus. Biochim Biophys Acta 1968; 158(2): 313–315. [3] Ballou C. Structure and biosynthesis of the mannan component of the yeast cell envelope. Adv Microb Physiol 1976; 14(11): 93–158. [4] Bandoni RJ, Boekhout T. Tremelloid genera with yeast phases. In: Kurtzman CP, Fell JW, eds. The yeasts, a taxonomic study. Amsterdam, Netherlands: Elsevier, 1998: 705–717. [5] Bandoni RJ, Zang Mu. On undescribed Tremella from China. Mycologia 1990; 82: 270–273. [6] Banno I. Studies on the sexuality of Rhodotorula. J Gen Appl Microbiol 1967; 13:167– 196. [7] Barnett JA, Lichtenthaler FW. A history of research on yeasts 3: Emil Fischer, Eduard Buchner and their contemporaries, 1880-1900. Yeast 2001; 18(4): 363–388. [8] Barnett JA, Robinow CF. A history of research on yeasts 4: cytology part I, 1890– 1950. Yeast 2002; 19(2): 151–182. [9] Barnett JA. A history of research on yeasts 8: taxonomy. Yeast 2004; 21(14): 1141– 1193. [10] Bartnicki-Garcia S. Cell Wall Chemistry, Morphogenesis, and Taxonomy of Fungi. Ann Rev Microbiol 1968; 22: 87–108. [11] Bell PJ. Yeast differentiation using histone promoter sequences. Lett Appl Microbiol. 2004; 38(5): 388–392. [12] Berbee ML, Taylor JW. Dating the evolutionary radiations of the true fungi. Canadian J Botany 1993; 71(8): 1114–1127. [13] Boekhout T, Bandoni RJ, Fell JW, Kwon-Chung KJ. Discussion of teleomorphic and anamorphic genera of heterobasidiomycetous yeasts. In: Kurtzman CP, Fell JW, eds. The yeasts, a taxonomic study. Amsterdam, Netherlands: Elsevier, 1998: 609–625.

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[32] Flossdorf J. A rapid method for the determination of the base composition of bacterial DNA. J Microbiol Methods 1983; 1: 305–311. [33] Fukatsu T, Ishikawa H. Phylogenetic position of yeast-like symbiont of Hamiltonaphis styraci (Homoptera, Aphididae) based on 18S rDNA sequence. Insect Biochem Mol Biol 1996; 26(4): 383–388. [34] Goffeau A, Barrell BG, Bussey H, Davis RW, Dujon B, Feldmann H, Galibert F, Hoheisel JD, Jacq C, Johnston M, Louis EJ, Mewes HW, Murakami Y, Philippsen P, Tettelin H, Oliver SG. Life with 6000 genes. Science 1996; 274(5287): 546–567. [35] Gueho E, Kurtzman CP, Peterson SW. Phylogenetic relationships among species of Sterigmatomyces and Fellomyces as determined from partial rRNA sequences. Int J Syst Bacteriol 1990; 40(1): 60–65. [36] Hagler AN, Ahearn DG. Rapid diazonium blue B test to detect basidiomycetous yeasts. Int J Syst Bacteriol 1981; 31: 204–208. [37] Hamamoto M, Nakase T. Phylogenetic analysis of the ballistoconidium-forming yeast genus Sporobolomyces based on 18S rDNA sequences. Int J Syst Evol Microbiol 2000; 50(3): 1373–1380. [38] Hamamoto M, Sugiyama J, Komagata K. DNA base composition of strains in the genera Rhodosporidium, Cystofilobasidium, and Rhodotorula determined by reversedphase high-performance liquid chromatography. J Gen Appl Microbiol 1986; 32: 215– 223. [39] Hamamoto M, Thanh VN, Nakase T. Bannoa hahajimensis gen. nov., sp. nov., and three related anamorphs, Sporobolomyces bischofiae sp. nov., Sporobolomyces ogasawarensis sp. nov. and Sporobolomyces syzygii sp. nov., yeasts isolated from plants in Japan. Int J System Evol Microbiol 2002; 52: 1023–1032. [40] Herrero E, de la Torre MA, Valentin E. Comparative genomics of yeast species: new insights into their biology. Int Microbiol. 2003; 6(3): 183–190. [41] Hirooka K, Bamba T, Fukusaki E, Kobayashi A. Cloning and kinetic characterization of Arabidopsis thaliana solanesyl diphosphate synthase. Biochem J 2003; 370(Pt 2): 679–686. [42] Hori H, Osawa S. Evolutionary change in 5S RNA secondary structure and a phylogenic tree of 54 5S RNA species. Proc Natl Acad Sci USA 1979; 76(1): 381– 385. [43] Howard DH, Kwon-Chung KJ. Zoopathogenic basidiomycetous yeasts. Stud Mycol 1995; 38: 59–66. [44] James SA, Cai J, Roberts IN, Collins MD. A phylogenetic analysis of the genus Saccharomyces based on 18S rRNA gene sequences: description of Saccharomyces kunashirensis sp. nov. and Saccharomyces martiniae sp. nov. Int J Syst Bacteriol 1997; 47(2): 453–460. [45] Jones T, Federspiel NA, Chibana H, Dungan J, Kalman S, Magee BB, Newport G, Thorstenson YR, Agabian N, Magee PT, Davis RW, Scherer S. The diploid genome sequence of Candida albicans. Proc Natl Acad Sci USA 2004; 101(19): 7329–7334. [46] Katayama-Fujimura Y, Tsuzaki N, Kuraishi H. Ubiquinone, fatty acids and DNA base composition determination as a guide to taxonomy of the genus Thiobacillus. J Gen Microbiol 1982; 128: 1599–1611. [47] Kato M, Ozeki M, Kikuchi A, Kanbe T. Phylogenetic relationship and mode of evolution of yeast DNA topoisomerase II gene in the pathogenic Candida species. Gene 2001; 272(1-2): 275–281.

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[84] Phaff HJ, Miller MW. Cyniclomyces van der Walt & D.B. Scott. In: Kurtzman CP, Fell JW, eds. The yeasts, a taxonomic study. Amsterdam, Netherlands: Elsevier, 1998: 154–156. [85] Phaff HJ. Chemotaxonomy based on the polysaccharide composition of the cell walls and capsules. In: Kurtzman CP, Fell JW, eds. The yeasts, a taxonomic study. Amsterdam, Netherlands: Elsevier, 1998: 45–47. [86] Pore RS, Sorenson WG. Reniforma strues, a new yeast from wastewater. Mycologia 1990; 82: 549–553. [87] Scorzetti G, Fell JW, Fonseca A, Statzell-Tallman A. Systematics of basidiomycetous yeasts: a comparison of large subunit D1/D2 and internal transcribed spacer rDNA regions. FEMS Yeast Res. 2002; 2(4): 495–517. [88] Sjamsuridzal W, Tajiri Y, Nishida H, Thuan TB, Kawasaki H, Hirata A,Yokota A, Sugiyama J. Evolutionary relationships of members of the genera Taphrina, Protomyces, Schizosaccharomyces, and related taxa within the archiascomycetes: integrated analysis of genotypic and phenotypic characters. Mycoscience 1997; 38: 267–280. [89] Smith MTh, Shann C, Batenburg-van der Vegte WH, Schmitt R, Wehrli E, Roeijmans HJ, van Eijk GW. Botryozyma nematodophila gen. nov., spec. nov. (Candidaceae). Antonie van Leeuwenhoek. 1992; 61(4): 277–284. [90] Smith MTh, Oosporidium Stautz. In: Kurtzman CP, Fell JW, eds. The yeasts, a taxonomic study. Amsterdam, Netherlands: Elsevier, 1998: 598–599. [91] Smith MTh, Poot GA. Genomic characterization of Dipodascus capitatus, Dipodascus spicifer and Geotrichum clavatum. Antonie van Leeuwenhoek 1998; 74: 229–235. [92] Sugiyama J, Fukagawa M, Chiu SW, Komagata K. Cellular carbohydrate composition, DNA base composition, ubiquinone systems, diazonium blue B color test in the genera Rhodosporidium, Leucosporidium, Rhodotorula and related basidiomycetous yeasts. J Gen Appl Microbiol 1985; 31: 519–550. [93] Sugiyama J, Suh SO. Sympodiomycopsis Sugiyama, Tokuoka & Komogata. In: Kurtzman CP, Fell JW, eds. The yeasts, a taxonomic study. Amsterdam, Netherlands: Elsevier, 1998: 846–847. [94] Suh SO, Nakase T. Phylogenetic analysis of the ballistosporous anamorphic genera Udeniomyces and Bullera, and related basidiomycetous yeasts, based on 18S rDNA sequence. Microbiol 1995; 141(Pt4): 901–906. [95] Suh SO, Noda H, Blackwell M. Insect symbiosis: derivation of yeast-like endosymbionts within an entomopathogenic filamentous lineage. Mol Biol Evol 2001;18(6): 995–1000. [96] Suh SO, Sugiyama J. Phylogeny among the basidiomycetous yeasts inferred from small subunit ribosomal DNA sequence. J Gen Microbiol 1993;139(Pt7): 1595–1598. [97] Swann EC, Taylor JW. Phylogenetic perspectives on basidiomycete systematics: evidence from the 18S rRNA gene. Canadian J Bot 1995; 73: S862–S868. [98] Swiezewska E, Dallner G, Andersson B, Ernster L. Biosynthesis of ubiquinone and plastoquinone in the endoplasmic reticulum-Golgi membranes of spinach leaves. J Biol Chem 1993; 268(2): 1494–1499. [99] Thanh VN, Van Dyk MS, Wingfield MJ. Debaryomyces mycophilus sp. nov., a siderophore-dependent yeast isolated from woodlice. FEMS Yeast Res 2002; 2(3): 415–427. [100] Tomaszewski EK, Logan KS, Snowden KF, Kurtzman CP, Phalen DN. Phylogenetic analysis identifies the ‚megabacterium‘ of birds as a novel anamorphic ascomycetous

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[115] Yamada Y, Maeda K, Mikata K. The phylogenetic relationships of Rhodosporidium dacryoidum Fell, Hunter et Tallman based on the partial sequences of 18S and 26S ribosomal RNAs: the proposal of Sakaguchia gen. nov., a heterobasidiomycetous yeast genus. Biosci Biotechnol Biochem 1994; 58(1): 99–103. [116] Yamada Y, Nakagawa Y, Banno I. The phylogenetic relationhip of the Q9-equipped species of the heterobasidiomycetous yeast genera Rhodosporidium and Leucosporidium based on partial sequences of 18S and 26S ribosomal ribonucleic acids: the proposal of a new genus Kondoa. J Gen Appl Microbiol 1989; 35: 377–385. [117] Yamada Y, Kondo K. Coenzyme Q system in the classification of the yeast genera Rhodotorula and Cryptococcus and the yeast-like genera Sporobolomyces and Rhodosporidium. J Gen Appl Microbiol 1973; 19: 59–77. [118] Yarrow D. Methods for the isolation, maintenance and identification of yeasts. In: Kurtzman CP, Fell JW, eds. The yeasts, a taxonomic study. Amsterdam Netherlands: Elsevier, 1998: 77–100. [119] Babjeva I.P. Family Saccharomycetaceae. Tachtadjanyana AL. (ed). Plants’ World, Vol. 2, Moscow, Prosveshenie, 1991 (in Russian).

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Evolutionary Relationships among Economically Important Species of Aspergillus Subgenera Aspergillus and Fumigati JÁNOS VARGA1, KRISZTINA RIGÓ 1, SÁNDOR KOCSUBÉ 1, KÁROLY PÁL1, BEÁTA TÓTH 2, ROBERT A. SAMSON 3 and ZOTIA KOZAKI EWICZ 4 1

Department of Microbiology, Faculty of Sciences, University of Szeged, Szeged, Hungary 2 Cereal Research non-Profit Company, Szeged, Hungary 3 Centraalbureau voor Schimmelcultures, Utrecht, The Netherlands 4 CABI Bioscience UK Centre, Bakeham Lane, Egham, Surrey TW 20 9TY, UK

ABSTRACT The genus Aspergillus is one of the most important filamentous fungal genera. Aspergillus species are used for the production of several enzymes, drugs and other metabolites useful for mankind. On the other hand, Aspergillus species produce a range of mycotoxins harmful to humans and animals, and can also cause several human and animal diseases. Molecular techniques enabled mycologists to get an insight into the phylogenetics of Aspergilli, and the evolution of mycotoxin production with special emphasis on aflatoxin biosynthesis. Phylogenetically unrelated species were found to produce the same mycotoxins. For example, aflatoxins have been produced under laboratory conditions by species belonging to three different sections, while ochratoxin A and patulin are produced by a variety of unrelated species. Based on this observation, mycotoxin-producing abilities of the isolates were lost (or

Address for correspondence: Dr. János Varga, Department of Microbiology, Faculty of Sciences, University of Szeged, P.O. Box 533, H-6701 Szeged, Hungary. Tel: 36-62-544-515 Fax: 36-62-544-823, E-mail: [email protected].

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gained) several times during the evolution of the genus. In this review, attempts have been made to summarize the current knowledge on the taxonomy of economically important Aspergilli belonging to Aspergillus subgenera Aspergillus and Fumigati, and on the evolution of mycotoxin production. Key Words: Aflatoxin, Eurotium, Neosartorya, Petromyces, ochratoxin, phylogenetics, taxonomy

INTRODUCTION

The name Aspergillus was first applied by Micheli in Nova Plantarum Genera of 1729 to molds bearing conidial heads and stalks [128]. However, these fungi began to be recognized as active agents of decomposition processes and causes of animal and human diseases only in the middle of the nineteenth century [128]. Today, Aspergillus is among the most economically important of the fungal genera. Aspergillus species can be both harmful and beneficial for mankind. They are responsible for a number of plant and human diseases and produce several toxins. At the same time, Aspergilli are widely used in the food and pharmaceutical industry for the production of various acids, enzymes and other compounds useful for humans [16, 26, 97, 115]. Aspergillus Species as Pathogens of Plants and Animals

Although not considered to be major causes of plant disease, Aspergillus species are responsible for several disorders in various plants. For example, chlorosis of almonds, albinism of citrus, black rot of onions, crown rot of peanuts, and vine canker of grapes are caused by Aspergilli [103, 164]. The most common plant pathogens are Aspergillus niger and A. flavus [164]. In contrast with specialized plant pathogens such as powdery mildews, rusts and some Fusarium species, Aspergillus species are opportunistic pathogens without host specialization [145]. While only a limited number of Aspergillus species can invade living plant tissues, several are encountered as storage molds on plant products [83, 128]. Several Aspergillus species are considered as opportunistic human and animal pathogens. Diseases caused by Aspergillus species are increasing in importance, especially among immunocompromised hosts. Clinical manifestations are variable, ranging from allergic to invasive disease, largely depending on the status of the host’s immune system. Aspergillus fumigatus is of prevailing importance. This species may cause several

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diseases, such as allergic bronchopulmonary aspergillosis, aspergilloma, and invasive aspergillosis, a usually fatal disease particularly in immunocompromised patients [19]. Today, A. fumigatus is responsible for more than 90% of invasive aspergilloses in humans [89]. Other species like A. niger, A. terreus and A. flavus can also be the causative agents of different diseases like sinusitis, osteomyelitis, keratitis, bronchiectasis, otomycosis and ophthalmitis [98]. Mycotoxins Produced by Aspergilli

Aspergillus species can contaminate foods and feeds at different stages including pre- and postharvest stages, processing and handling. The most important aspect is the formation of mycotoxins, which may have harmful effects on human and animal health. Several Aspergillus mycotoxins have been identified as contaminants in foods and feeds, the economically most important of which are the aflatoxins, ochratoxins and patulin (Table 1; Fig. 1). Recent studies indicate that these compounds can be produced by a number of Aspergillus species [164, 173]. However, only a few of these mycotoxin producers can be regarded as potential health hazards either because they produce only traces of the given mycotoxin, or are encountered rarely, if at all, in food products (for example, aflatoxin producing A. ochraceoroseus, Emericella venezuelensis and E. astellata isolates) [124]. However, new data indicate that some species recently reported to be mycotoxin producers can be regarded as main sources of mycotoxin contamination in various food products [1, 15, 124]. For example, although ochratoxin-producing abilities of Table 1. Some economically important mycotoxins produced by Aspergillus species in various agricultural products [15, 87, 164, 172] Mycotoxins

Agricultural product

Species

Aflatoxins

Peanuts, Corn, Cotton

A. flavus, A. parasiticus, A. nomius

Ochratoxins

Cereals Meats, Cheese Grapes, Wine Coffee, Spices Figs

P. verrucosum P. nordicum A. niger, A. carbonarius A. ochraceus, A. niger, A. carbonarius A. alliaceus

Patulin

Cereals, Malt Apples, Pears

P. expansum, A. clavatus P. expansum

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aflatoxin B1

O

Fig. 1.

O

O

O

HOOC

O

OH

NH

OCH3

ochratoxin A

O

O

O O

Cl

O OH patulin

Structures of the most common mycotoxins produced by Aspergilli.

black Aspergilli have only recently been discovered, these fungi are now considered as major sources of ochratoxin contamination in wine, raisins and coffee [124]. Aspergillus Species in the Food and Pharmaceutical Industries

Aspergillus species are used in oriental fermentation processes for the production of soy sauce, miso, sake, mirin, shochu and katsuobushi [64, 108, 185]. Aspergillus species have the capacity to secrete large amounts of proteins, metabolites and organic acids into their culture medium. This property has been widely exploited by the food and beverage industries where compounds secreted by Aspergilli have been used for decades. This has led to the GRAS (generally regarded as safe) status for some of their products. Filamentous fungi like A. awamori, A. niger and A. oryzae are, therefore, suitable organisms for the production of commercially interesting homologous and heterologous proteins. Consequently, Aspergilli are frequently used in the fermentation industry for the production of various organic acids including citric acid, gluconic acid, itaconic acid, malic acid, tartaric acid [18], and enzymes like =- and C-amylases, catalase, cellulase, =- and >-galactosidases, >-glucanase, glucose oxidase, hemicellulase, inulinase, invertase, lipase, peroxidase, pectinases, acid and alkaline proteases, tannase, urate oxidase and xylanase [95]. A. oryzae and black Aspergilli are used as hosts for heterologous expression of a number of economically important proteins like antibody fragments, interleukin 6, manganese peroxidase, prochymosin, glucoamylase and others [61, 75, 126]. Besides, genetically modified black Aspergillus isolates are used in the fermentation industry for the production of various enzymes including arabinofuranosidase, chymosin, glucose oxidase, pectin esterase, phospholipase, phytase and xylanase [115].

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Various Aspergillus species are used in the pharmaceutical industry in biotransformations [79]. The species most frequently involved include A. alliaceus, A. flavus, A. niger, A. ochraceus, A. oryzae and A. terreus. Besides, A. terreus and some other Aspergillus species produce lovastatin, a cholesterol-lowering compound [7, 77]. TAXONOMIC OUTLINE OF THE ASPERGILLUS GENUS

Partly because of its economic importance, the genus Aspergillus has one of the better described taxonomies among filamentous fungi. Aspergillus teleomorphs belong to different genera all of which are members of family Trichocomaceae of the Eurotiales order (Table 2)[8]. This order is monophyletic as proved by several molecular studies [56]. Raper and Fennell [128] described 18 species groups within the Aspergillus (form) genus based mainly on cultural and morphological features, which were treated as sections belonging to six subgenera by Gams et al. [53]. Phylogenetic studies of ribosomal RNA gene sequences led to the acceptance of three subgenera with a total of 15 sections and the socalled ‘Warcupiella group’, a treatment currently accepted by most Aspergillus researchers (Table 2)[121]. Our studies focused on the phylogenetic analysis of sequences of the intergenic spacer region and the 5.8 S rRNA gene (ITS region) and the D1-D2 region of the 28 S rRNA gene in various Aspergilli (Fig. 2). This review deals with Aspergillus subgenera Aspergillus and Fumigati (Table 2). Aspergillus subgenus Aspergillus Aspergillus section Aspergillus

Members of this section are characterized by uniseriate blue-green conidial heads, and cleistothecia in shades of yellow, orange or orange-

Fig. 2. Organization of the rRNA gene cluster in Aspergilli, and location of the primers used to amplify the ITS region and the D1-D2 region of the 28 S rDNA.

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Classification of Aspergillus species based on rDNA sequences Section

Assignment to subgenus Gams et al. [53]

Peterson [121]

Teleomorph

Aspergillus

Aspergillus Restricti Cervini Terreia Flavipedesa Nigri Circumdati Flavi Cremei Candidi Wentii

Aspergillus Aspergillus Fumigati Nidulantes Nidulantes Circumdati Circumdati Circumdati Circumdati Circumdati Circumdati

Aspergillus Aspergillus Aspergillus Aspergillus Aspergillus Aspergillus Aspergillus Aspergillus Aspergillus Aspergillus –

Eurotium – – Fennellia Fennellia – Neopetromyces Petromyces Chaetosartorya – –

Fumigati

Fumigati Clavati

Fumigati Clavati

Fumigati Fumigati

Neosartorya Neocarpenteles

Nidulantes Nidulantes Nidulantes Circumdati Ornati Ornati

Nidulantes – – Nidulantes Nidulantes Warcupiella gr.

Emericella – – – Sclerocleista Warcupiella

–

–

–

Nidulantes Nidulantes Versicolores Usti Sparsi Ornati Warcupiella group Ochraceoroseib a

Molecular data indicate that sections Terrei and Flavipedes should be merged [Varga et al. submitted] b This section has been proposed recently to accommodate A. ochraceoroseus and A. rambellii [50]

red. All members are osmophilic and require high concentrations of sugar or salt for optimal growth. Aspergillus species with Eurotium teleomorphs together with an asexual species, A. proliferans have originally been included in the A. glaucus species group [128]. This group has been renamed section Aspergillus by Gams et al. [53]. Raper and Fennell [128] involved 17 species in this group, while Kozakiewicz [83] recognized 26 species. Based on 28S rDNA sequences, Peterson [121] suggested merging sections Aspergillus and Restricti. In the modified section

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Aspergillus, he recognized at least 18 species, although some species (for example, E. appendiculatum, E. spiculosum or E. glabrum) have not been examined. Members of section Aspergillus are widely distributed in nature. These species occur on organic materials at low moisture levels, and can be isolated from stored cereals, from food products containing high concentrations of sugar or salt or from leather. Since the ascospores are heat resistant, Eurotium species can also contaminate canned products [144]. Xerophilic fungi also frequently cause spoilage of intermediate moisture and low water activity (aw) baked goods [59]. Eurotium species, Wallemia sebi and the xerophilic Aspergillus species, such as Aspergillus restrictus and A. penicillioides, are widely distributed fungi that are common contaminants of stored grains, nuts, spices and cereal products [122]. Eurotium species are also used for the fermentation of katsuobushi, a Japanese fermented fish product [108]. Eurotium species also produce a range of secondary metabolites which are harmful to animals or humans, including the nephrotoxic ochratoxin A [27, 43], kotanin derivatives [20], echinulin, which causes feed refusal in swine [180], erythroglaucin, auroglaucin and flavoglaucin which have been implicated in the toxicity of molded barley to rabbits [106, 107], the antiviral compound xanthocillin [10], and the anthraquinone derivative physcion [11]. Peterson [121] recognized 18 species in the revised section Aspergillus based on sequence analysis of the D1-D2 region of the 28 S rRNA gene (Table 3. Fig. 3a). The species previously classified into section Restricti formed one clade together with E. halophilicum. Considerable genetic variability was observed within these asexual species indicating that at least six asexual species should be retained in this section [121, 150, 151]. Peterson [121] ranked E. montevidensis, E. cristatum, E. athecium, E. medium, E. umbrosum and E. pseudoglaucum as variants of E. amstelodami, E. amstelodami, E. rubrum, E. echinulatum, E. herbariorum and E. repens, respectively. The ITS region of isolates belonging to section Aspergillus (Fig. 3b) has been examined and the analysis indicated that in most species recognized earlier in this section form two groups, one was characterized by E. amstelodami, while the other one included E. herbariorum (Fig. 3b). Due to resolution of the phylogenetic tree within these groups, it was not possible to distinguish between the species included unambiguously. Further regions of the genome of these isolates should be analyzed to draw more accurate taxonomic conclusions.

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b.

Fig. 3. Neighbor-joining trees of Aspergillus section Aspergillus based on ribosomal RNA gene sequences; a. tree based on sequences of the D1-D2 region of the 28 S rDNA gene; b. tree based on ITS sequences. Numbers above branches indicate bootstrip values; only values > 50% are shown. The bar represents genetic distance.

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Table 3. Species assigned to Aspergillus sections Aspergillus, Candidi, Nigri and Cremei (associated teleomorph is in brackets; modified after Pitt et al [123]) Section Aspergillus (Eurotium): Teleomorph unknown: A. caesiellus Saito 1904 A. conicus Blochwitz 1914 A. gracilis Bainier 1907 A. penicillioides Spegazzini 1896 A. proliferans Smith 1943 A. restrictus Smith 1931 A. vitricola Ohtsuki 1962 Teleomorph known: E. amstelodami Mangin 1909 E. appendiculatum Blaser 1976 E. athecium (Raper & Fennell) Arx 1974 E. carnoyi Malloch & Cain 1972 E. chevalieri Mangin 1909 E. cristatum (Raper & Fennell) Malloch & Cain 1972 E. echinulatum Delacroix 1893 E. glabrum Blaser 1976 E. halophilicum Christensen et al. 1961 E. herbariorum Link 1809 E. intermedium Blaser 1976 E. leucocarpum Hadlok & Stolk 1969 E. medium Meissner 1897 E. niveoglaucum (Thom & Raper) Malloch & Cain 1972 E. pseudoglaucum (Blochwitz) Malloch & Cain 1972 E. repens de Bary 1870 E. rubrum König et al. 1901 E. tonophilum Ohtsuki 1962 E. umbrosum (Bainier & Sartory) Malloch & Cain 1972 E. xerophilum Samson & Mouchacca 1975 Section Cervini (teleomorph unknown): A. bisporus Kwon-Chung & Fennell 1971

A. A. A. A.

cervinus Massee 1914 kanagawaensis Nehira 1951 nutans McLennan & Ducker 1954 parvulus Smith 1961

Section Candidi (teleomorph unknown): A. campestris Christensen 1982 A. candidus Link 1809 A. taichungensis Yaguchi et al. 1995 Section Nigri (teleomorph unknown): Biseriate species: “A. niger aggregate”: A. costaricaensis Samson & Frisvad 2004 A. foetidus Thom & Raper 1945 A. homomorphus Steiman et al. ex Samson & Frisvad 2004 A. lactocoffeatus Frisvad & Samson 2004 A. niger Tieghem 1867 A. piperis Samson & Frisvad 2004 A. sclerotioniger Samson & Frisvad 2004 A. tubingensis (Schober) Mosseray 1934 A. vadensis Samson, de Vries et al. 2004 “A. brasiliensis” Others: A. carbonarius (Bainier) Thom 1916 A. ellipticus Raper & Fennell 1965 A. heteromorphus Batista & Maia 1957 “A. ibericus” “A. keveiensis” (Aspergillus sp. IN7) Uniseriate species: A. aculeatus Iizuka 1953 A. japonicus Saito 1906 Section Cremei (Chaetosartorya): A. dimorphicus Mehrota & Prasad 1969 A. flaschentraegeri Stolk 1964 A. gorakhpurensis Kamal & Bhagrava 1969 A. itaconicus Kinoshita 1931 Table 3 Contd.

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Table 3 Contd.

A. pulvinus Kwon-Chung & Fennell 1965 A. sepultus Tuthill & Christensen 1986 A. wentii Wehmer 1896 C. chrysella (Kwon-Chung & Fennell) Subramanian 1972

[A. chryseides Samson & Gams 1985] C. cremea (Kwon-Chung & Fennell) Subramanian 1972 [A. cremeoflavus Samson & Gams 1985] C. stromatoides Wiley & Simmons 1973 [A. stromatoides Raper & Fennell 1965]

Aspergillus section Nigri

Black Aspergilli (Aspergillus niger species group [128]) have a significant impact on modern society. Many species cause food spoilage, and several are used in the fermentation industry to produce hydrolytic enzymes, such as amylases or lipases, and organic acids, such as citric acid and gluconic acid [16, 83, 115, 128]. They are also candidates for genetic manipulation in the biotechnology industries since A. niger has been granted the GRAS status by the Food and Drug Administration of the US government. Although the main source of black Aspergilli is soil, members of this section have been isolated from various other sources [164]. Black Aspergilli are one of the more problematic groups for identification. Raper and Fennell [128] described 12 species of the black Aspergilli. Al-Musallam [9] revised the taxonomy of the A. niger group by taking mainly morphological features into account. She recognized seven species within this group (A. japonicus, A. carbonarius, A. ellipticus, A. helicothrix, A. heteromorphus, A. foetidus and A. niger), and described A. niger itself as an aggregate consisting of seven varieties and two formae. Kozakiewicz [83] distinguished A. ellipticus, A. heteromorphus, A. japonicus, A. helicothrix, A. atroviolaceus (treated as A. aculeatus or A. japonicus var. aculeatus in other classifications) and A. carbonarius, a species exhibiting echinulate conidial ornamentations, which distinguished it from the rest of black Aspergillus strains, displaying verrucose conidia. Within the verrucose category, A. fonsecaeus, A. acidus (A. foetidus var. acidus), A. niger var. niger, A. niger var. phoenicis, A. niger var. ficuum, A. niger var. tubingensis, A. niger var. pulverulentus, A. niger var. awamori, A. citricus (A. foetidus) and A. citricus var. pallidus (A. foetidus var. pallidus) were recognised. In recent years, different phenotypic and genotypic markers have been applied for clarifying the taxonomy of black Aspergilli. Among the genotypic approaches, nuclear

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and mitochondrial DNA (mtDNA) polymorphisms and PCR-based techniques led to the recognition of four species within the A. niger species complex (A. niger, A. tubingensis, A. brasiliensis, A. foetidus) [6, 86, 102, 168-170]. Sequence comparisons of nuclear genes encoding various extracellular enzymes supported these results [22, 36, 58]. Yokoyama et al. [190] also distinguished two clusters within the A. niger species complex based on phylogenetic analyses of sequences of the mitochondrial cytochrome b gene (Fig. 4). Several well-known species’ names such as A. awamori, A. usamii, A. phoenicis and A. ficuum have been reduced to synonymy. Regarding other black Aspergillus species, phylogenetic analyses of sequences of the intergenic spacer region and the 5.8 S rRNA gene (ITS region), and the D1-D2 region of the 28 S rRNA gene indicated that apart from those mentioned earlier, at least five other species belong to this section: A. heteromorphus, A. ellipticus, A. carbonarius, A. japonicus and A. aculeatus (Table 3; Fig. 4) [63, 78, 114, 170]. The uniseriate species A. japonicus, A. aculeatus and isolate CBS 114.80, which is considered to represent a new species, form one welldefined clade, while the biseriate species are on a separate branch [114] (Fig. 4). Although some toxins and other toxic agents, for example, oxalic acid, nigragillin, malformins and naphtho-g-pyrones [140], have been reported to be produced by black Aspergilli, the production of ochratoxins is of real economic importance. Until recently, ochratoxin production has been observed only in A. niger and A. carbonarius [5, 6, 72, 110, 153, 161, 184]. These species are now considered as major sources of ochratoxin contamination in tropical and subtropical foods including dried vine fruits, wines and coffee [2, 23, 124]. Aspergillus vadensis has recently been described as a new biseriate black Aspergillus species which was isolated from dead plant debris [3739]. RFLP patterns of the rRNA, pkiA and pelA genes and secondary metabolite profiles of this species were different from all other black Aspergilli justifying the treatment of this single isolate as a new species [38]. In a recent paper, Samson et al. [133] described several new species belonging to this section. A. costaricaensis was isolated from soil in Costa Rica and produces large pink to grayish brown sclerotia. Aspergillus lacticoffeatus was isolated on coffee beans in Venezuela and Indonesia, and is an efficient producer of ochratoxin A. Aspergillus piperis was isolated from black grounded pepper and produces large yellow to pink

b.

c.

Fig. 4. Neighbor-joining trees of Aspergillus section Nigri based on ribosomal RNA and cytochrome b gene sequences; a. tree based on sequences of the D1-D2 region of 28 S rDNA gene; b. tree based on mitochondrial cytochrome b sequences; c. tree based on ITS sequences. Numbers above branches indicate bootstrap values; only values >50% are shown. The bar represents genetic distance.

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brown sclerotia. A. sclerotioniger was isolated from green coffee beans, and produces large yellow to red brown sclerotia and large amounts of ochratoxin A. A. homomorphus which was incorrectly described earlier, is also accepted. At this time, 15 species are accepted in Aspergillus section Nigri, and at least four of these produce ochratoxin A: A. carbonarius, A. niger, A. lacticoffeatus and A. sclerotioniger. Other black Aspergilli including A. tubingensis and A. japonicus isolates also have the capacity to produce ochratoxins [3, 14, 35, Varga et al., unpublished observations]. These observations should be confirmed in other laboratories. Korean authors also described a new black Aspergillus species named A. coreanus from traditional Korean Nuruk (malt) [191]. The ITS sequence of this species differs from that of A. tubingensis by a single nucleotide, questioning the species status of this isolate (data not shown). Recently, two presumably new black Aspergillus species related to A. carbonarius have also been identified. “Aspergillus ibericus” was isolated from Portuguese grapes. This species is similar to A. carbonarius, but does not produce ochratoxin A, and has smaller conidia than typical A. carbonarius isolates. Sequence analysis of the ITS region and the calmodulin gene, and AFLP analysis justified its treatment as a new species [24]. Another black Aspergillus isolate resembling A. carbonarius, IN7 isolated from Indian soil and characterized previously exhibited different mtDNA, RAPD and isoenzyme profiles than the other A. carbonarius isolates examined, indicating that this isolate, tentatively called “A. keveionsis”, may also represent a new species in section Nigri [78]. Sequencing of the ITS region of this isolate is in progress. The currently accepted species in section Nigri is listed in Table 3 (species not yet described formally are in quotation marks). A synoptic key to species according to Samson et al. [133] is also included (Table 4). Aspergillus section Flavi

Aspergillus section Flavi historically includes species with conidial heads in shades of yellow-green to brown, and dark sclerotia. Isolates of the socalled domesticated species, such as A. oryzae, A. sojae and A. tamarii, are used in oriental food fermentation processes and as hosts for heterologous gene expression [26]. Genetically modified A. oryzae strains are used for the production of enzymes including lactase, pectin esterase, lipase, protease and xylanase [115]. Several species of section Flavi

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Table 4. 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14.

Provisional synoptic key to species in Aspergillus section Nigri

Species list: A. aculeatus A. brasiliensis ined A. carbonarius A. costaricaensis A. ellipticus A. japonicus A. foetidus A. heteromorphus A. homomorphus A. lacticoffeatus A. niger A. piperis A. sclerotioniger A. tubingensis

15. A. vadensis Conidia more than 6 mm diam: Conidia spinulose: Conidia strongly ellipsoidal: Metulae not produced: Metulae less than 15 mm in length: Production of sclerotia: Sclerotia yellow to orange: Sclerotia yellow to pinkish brown: Sclerotia pink to grayish yellow: Colony diameter at 25 °C on CYA, 7 d, less than 30 mm: Colony diameter at 37 °C on CYA, 7 d, larger than 70 mm: Colony diameter at 37 °C on CYA 7.d, between 55 and 65 mm: Colony diameter at 37 °C on CYA 7 d, less than 40 mm: Colony diameter at 37 °C on CYA, 7 d, 0 mm: Acid production on CREA agar weak or not present:

3, (5) (3), 5, 6, 8, 9 (1), 5, (6) 1, 6 (7), (8), 9, (10), (11), 13, (14), (15) (1), (3), 4, (5), (6), 12, 13, (14) 13 12 3 15 2, 7, 10, 11, 12, 14 4, 15 1, 3, 5, 6, 8, 9, 13 (5), 8 (1), (7), 8, 9 Table 4 Contd.

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Table 4 Contd.

Conidium colour en masse light brown to dark blonde: Conidium colour en masse greenish–olive: Production of ochratoxin A: Production of pyranonigrin A: Production of one or more naphtha-g-pyrones: Production of asperazine: Production of secalonic acid D: Production of aflavinines: Production of antafumicins: Production of corymbiferan lactone/funalenone-like compounds: Production of kotanin, desmethylkotanin and/or orlandin: Production of austdiol: Production of neoxaline:

10, 15 8, 3, 3, 2,

(15) 10, (11), 13 7, 10, 11, 12, 13, 14 3, 4, 7, 11, 12, 13, 14, 15

7, 1, 4, 7 4,

14, 15 9 12, (14) 13

10, (11) 5 (1)

(Numbers in parentheses: feature not always present)

produce aflatoxins, among which aflatoxin B1 is the most toxic of the many naturally occurring secondary metabolites produced by fungi. Aflatoxins are mainly produced by A. flavus and A. parasiticus, which coexist with and grow on almost any crop or food. Regarding the intra- and interspecific variability of Aspergillus section Flavi, ITS, 28 S rDNA and mitochondrial cytochrome b sequences of type strains or representative isolates of the species and subspecies currently assigned to this section have been analyzed recently [121, 129, 182]. Phylogenetic analysis of sequence data indicated that species of Aspergillus section Flavi form distinct clades (Fig. 5). The main clades identified based on sequence data could also be distinguished based on colony color and their ubiquinone systems. The ‘A. flavus’ clade includes species characterized with Q-10(H2) as their main ubiquinone, and conidial colors in shades of green, bearing dark sclerotia. Studies on the genetic variability of A. flavus indicated that the name is currently applied to a paraphyletic group of isolates that may produce aflatoxins B or G, and have large or small sclerotia [54]. It was suggested that isolates with small sclerotia, able to produce both aflatoxins B and G (group II),

b.

Fig. 5. Neighbor-joining trees of species of Aspergillus section Flavi based on ribosomal RNA gene sequences. a. tree based on sequences of the D1-D2 region of the 28 S rDNA gene; b. tree based on ITS sequences. Numbers above branches indicate bootstrap values; only values >50% are shown. The bar represents genetic distance.

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deserve recognition as a new species [42, 54]. This species has been described recently as A. parvisclerotigenus [50]. The other group (group I) includes isolates producing only aflatoxin B, and having large or small sclerotia. This group also includes isolates of A. oryzae, and has previously been described as having a recombining population structure [57]. Although several lines of evidence suggest that A. oryzae and A. sojae are morphological variants of A. flavus and A. parasiticus, respectively, it was suggested that these taxa should be retained as separate species because of the regulatory confusion that conspecificity might generate in the food industry [57]. The ‘A. tamarii’ clade contains species with ubiquinone system Q-10(H2), and conidia in shades of olive to brown, while the ‘A. alliaceus’ clade consists of species with the Q-10 ubiquinone system, and conidia in shades of ochre [85, 129]. Two species of this clade, Petromyces alliaceus and P. albertensis, produce high amounts of ochratoxin (50-300 mg ml-1), and are considered to be responsible for ochratoxin contamination of figs [15, 32, 166]. Petromyces alliaceus may also cause chronic otorrhoea [82]. Besides their harmful properties, Petromyces species also have beneficial effects on mankind. P. alliaceus strains produce asperlicins, potent cyclic peptide antagonists of cholecystokinin, which is a neurotransmitter involved in the hormonal regulation of pancreatic and gastric secretion, gallbladder contraction, and gut motility [90]. P. alliaceus strains are also used for steroid and alkaloid transformations [21], and for the production of pectin degrading enzyme preparations [104]. The recently described aflatoxin-producing species A. pseudotamarii and A. bombycis are closely related to A. caelatus and A. nomius, respectively [73, 117]. Physiological properties and mycotoxin-producing abilities of these taxa justify their treatment as separate species (Table 5)[73, 117]. Recent data indicate that A. nomius is a paraphyletic group likely to contain several other species [34, 41, 42, 44]. Two other species, A. avenaceus and A. leporis, were found to form separate lineages not closely related to any of the main clades identified. Aspergillus clavatoflavus and A. zonatus have been proposed to be excluded from Aspergillus section Flavi, in accordance with previous suggestions [83]. Phylogenetic analysis of partial 28 S rRNA gene sequences supported the findings (Fig. 5). The currently accepted species of section Flavi are listed in Table 6.

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Table 5. Mycotoxin producing abilities of the economically important species of Aspergillus section Flavi, and other aflatoxin-producing Aspergilli [48, 50, 51, 138, 147, 172] Species

Aflatoxins

CPAa

Kojic

B

G

Aspergillus section Flavi A. avenaceus – A. bombycis + A. caelatus – A. flavus group I +

– + – –

– – – ±

– + + +

A. parvisclerotigenus

+

±

+

+

A. lanosus A. leporis

– –

– –

– –

+ +

A. nomius

+

+

–

+

A. oryzae A. parasiticus

– +

– +

± –

+ +

A. pseudotamarii A. tamarii-like sp. A. sojae A. tamarii A. zhaoquingensis P. alliaceus

+ + – – + –

– + – – – –

+ + – + – –

+ + + + + +

Aspergillus section Nidulantes (Emericella) E. astellata

+

–

–

–

E. sp IBT 21903 E. venezuelensis

+ +

– –

– –

– –

Others

acid avenaciolide – – aspergillic acid, paspaline, nominine aspergillic acid, paspaline, nominine griseofulvin leporine, pseurotin, antibiotic Y aspergillic acid, pseurotin, nominine – aspergillic acid, parasiticol, parasiticolide A – ND – fumigaclavine A tenauzonic acid b ochratoxins, asperlicine, nominine, kotanins, paspaline

sterigmatocystin, terrein, asperthecin sterigmatocystin sterigmatocystin, terrein, shamixanthone, asperthecin Table 5 Contd.

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Table 5 Contd.

Aspergillus section Ochraceorosei A. ochraceoroseus A. rambellii a b

+ +

– –

– –

– –

sterigmatocystin sterigmatocystin, versicolorins, averufin

CPA, cyclopiazonic acid; ND, not determined this species produces only aflatoxin B2

Table 6. Species assigned to Aspergillus sections Flavi and Circumdati (associated teleomorph is in brackets; modified after Pitt et al. [123]) Section Flavi (Petromyces):

Section Circumdati (Neopetromyces):

A. avenaceus Smith 1943 A. bombycis Peterson et al. 2001 A. caelatus Horn 1997 A. coremiiformis Bartoli & Maggi 1978 A. flavus Link 1809 A. lanosus Kamal & Bhargava 1969 A. leporis States & Christensen 1966 A. nomius Kurtzman et al. 1987 A. oryzae (Ahlburg) Cohn 1884 A. parasiticus Speare 1912 A. parvisclerotigenus Frisvad and Samson 2004 A. pseudotamarii Ito et al. 2001 A. sojae Sakaguchi & Yamada ex. Murakami 1971 A. tamarii Kita 1913 P. albertensis Tewari 1985 [A. albertensis Tewari 1985] P. alliaceus Malloch & Cain 1973 [A. alliaceus Thom & Church 1926]

A. auricomus (Gueguen) Saito 1939 A. bridgeri Christensen 1982 A. cretensis Frisvad & Samson 2004 A. elegans Gasperini 1887 A. flocculosus Frisvad & Samson 2004 A. insulicola Montemayor & Santiago 1975 A. melleus Yukawa 1911 A. neobridgeri Frisvad & Samson 2004 A. ochraceus Wilhelm 1877 A. ochraceopetaliformis Batista & Maia 1957 A. ostianus Wehmer 1899 A. persii Zotti & Corte 2002 A. pseudoelegans Frisvad & Samson 2004 A. roseoglobulosus Frisvad & Samson 2004 A. sclerotiorum Huber 1933 A. steynii Frisvad & Samson 2004 A. sulphureus (Fresen.) Wehmer 1901 A. westerdijkiae Frisvad & Samson 2004 N. muricatus (Udagawa et al.) Frisvad & Samson 2000 [A. muricatus Udagawa et al. 1994]

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In the past few years, isolates with unusual mycotoxin profiles have been encountered in some food products. Kumeda et al. [84] identified a new genotype related to A. nomius predominating in Japanese and Vietnamese sugarcane field soils by heteroduplex panel analysis. Isolates resembling A. flavus, but producing aflatoxins B, G and cyclopiazonic acid were found in peanut fields in Argentina [162], while A. tamarii-like isolates producing the same pattern were found in black pepper and black tea samples in the Czech Republic [112]. These latter isolates were found to be closely related to A. tamarii based on ITS sequence data (data not shown). Further molecular analysis of these isolates is in progress. Aflatoxin-producing species (A. flavus, A. parasiticus, A. nomius, A. bombycis and A. pseudotamarii) were scattered throughout the dendrogram indicating that aflatoxin-producing ability was lost (or gained) several times during evolution. Another aflatoxin-producing species, A. ochraceoroseus was found to be unrelated to any of the species belonging to either sections Flavi or Circumdati [80, 81]. This species accumulates aflatoxin B1 and sterigmatocystin simultaneously, and has previously been assigned either to Aspergillus sections Wentii, Cremei, Cervini or Circumdati [31, 83, 135]. However, ITS and 28 S rDNA data indicated that this species is closely related to section Nidulantes [80]. Its distant relationship to Aspergillus section Flavi is supported by the observation that A. ochraceoroseus DNA hybridized only weakly if at all to the A. flavus and A. parasiticus aflatoxin biosynthetic gene probes [81]. Additionally, the order of genes of the aflatoxin biosynthetic gene cluster of A. ochraceoroseus was more similar to that of the sterigmatocystin gene cluster of A. nidulans than to that of A. parasiticus [81]. These data indicate that the aflatoxin and sterigmatocystin biosynthetic pathway genes in A. ochraceoroseus are different from known pathway genes. Another A. ochraceoroseus isolate was found to differ from the type strain sufficiently to be described as a new species, A. rambellii in a forthcoming paper [50]. These two isolates have been assigned to a new section, section Ochraceorosei which now includes species not able to grow at 37°C, producing yellow ellipsoidal conidia, biseriate conidial heads, long conidiophore stipes that are smooth and producing aflatoxin and sterigmatocystin. Additionally, aflatoxin production has also been observed recently in Emericella venezuelense and E. acristata isolates and in an undescribed Emericella isolate, IBT 21903, which are members of Aspergillus section Nidulantes [48, 51].

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Aspergillus section Circumdati

Aspergillus section Circumdati historically includes species with biseriate conidial heads in shades of yellow to ochre. Species of Aspergillus section Circumdati are economically important on account of a number of aspects. Several species produce diverse mycotoxins harmful for animals and humans including ochratoxin A, xanthomegnin and viomellein [13, 40, 163, 166]. Isolates of A. ochraceus or A. sclerotiorum are useful for biochemical transformation of steroids, alkaloids or phenazines [30]. Some species, for example, A. sclerotiorum and A. melleus, are important sources of proteolytic enzymes [96], and other metabolites [100], while sclerotia of several species contain antiinsectan compounds [111, 183]. The most frequently encountered species of the section, A. ochraceus is predominantly isolated from desert and cultivated soil, but is also a postharvest pathogen of several agricultural products including cereals, coffee beans, corn and pecans [83]. A. ochraceus was also found to cause allergic bronchopulmonary aspergillosis [109]. Interspecific variability of species assigned to this section was examined using phenotypic features, sequences of the ITS region and the D1-D2 region of the 28S rRNA gene (Fig. 6). Phylogenetic analysis of sequence data indicated that Aspergillus campestris, A. lanosus, A. dimorphicus and A. sepultus belong to Aspergillus sections Candidi, Flavi and Cremei, respectively (Fig. 6) [120, 175]. Two teleomorphic species previously assigned to this section, Petromyces alliaceus and P. albertensis, together with the asexual A. lanosus were found to belong to Aspergillus section Flavi [165, 175]. These results were also supported by phenotypic data, and by the main ubiquinones observed in these species [85]. Species of the revised Aspergillus section Circumdati formed two main clades, which could also be distinguished based on phenotypic methods (Fig. 6). A sexually reproducing ochratoxin-producing species, Neopetromyces muricatus was also found to belong to this section [49, 159, 165]. All these species are characterized by the Q-10(H2) ubiquinone system. Aspergillus auricomus and A. elegans did not belong to any of these clades. Ochratoxin-producing abilities of the isolates examined did not correlate with their taxonomic relationships based on ITS sequence data. However, a clade found not to produce ochratoxins during an earlier study of genetic variability of A. ochraceus was also identified in this study [167, 175].

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Recently, Peterson [118] also analyzed the relationships among species of this section using sequences of the ITS region, the calmodulin gene, the mitochondrial small subunit rRNA and translation elongation factor 1a. He could not distinguish A. ochraceus isolates from A. petrakii isolates, but could differentiate between A. ostianus, A. melleus and A. ochraceus. A. insulicola and A. ochraceopetaliformis isolates exhibited very small sequence divergence, however, they were retained in separate species. A. bridgeri and A. sclerotiorum were also found to be related, but were distinct species. He also identified one atypical A. petrakii and two atypical A. bridgeri isolates which possibly represent new species. Recently another new species, A. persii has also been assigned to this section [192]. In a recent paper, seven new species have been described in section Circumdati: A. cretensis, A. flocculosus, A. neobridgeri, A. pseudoelegans, A. roseoglobulosus, A. steynii, and A. westerdijkiae [46]. Twelve species of the accepted 20 produce mellein, 17 produce penicillic acid and 17 produce xanthomegnins. Nine species produce large amounts of ochratoxin A: A. cretensis, A. flocculosus, Neopetromyces muricatus, A. pseudoelegans, A. roseoglobulosus, A. westerdijkiae, A. sulphureus, A. ochraceus and A. sclerotiorum. Ochratoxin production in these species has been confirmed using HPLC with diode array detection and comparison to authentic standards. The currently accepted species of this section are listed in Table 6. Aspergillus section Terrei

This section historically includes isolates with strongly columnar, cinnamon to orange-brown biseriate conidial heads. Aspergillus terreus Thom is a ubiquitous fungus in our environment. Strains of this cosmopolitan species are frequently isolated from desert and grassland soils and compost heaps, and as contaminants of plant products like stored corn, barley and peanuts [83]. A. terreus is an economically important species used in the fermentation industry for the production of itaconic acid and itatartaric acid and for enzyme production [18, 95]. Aspergillus terreus isolates produce a range of secondary metabolites including lovastatin (mevinolin), a cholesterol lowering drug [7, 77, 97], the antitumor metabolites quadrone and asterriquinone [25, 76], acetylcholinesterase inhibitors such as territrem B and terreulactone [28, 29], or kodaistatins [179]. Other secondary metabolites produced by A. terreus isolates are considered as mycotoxins, including patulin,

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b.

Fig. 6. Neighbor-joining trees of species of Aspergillus section Circumdati based on ribosomal RNA gene sequences; a. tree based on sequences of the D1-D2 region of the 28 S rDNA gene; b. tree based on ITS sequences. Type strains are set in bold type, and the sections are indicated on the right. Numbers above branches indicate bootstrap values; only values >50% are shown. The bar represents genetic distance.

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ochratoxins, citrinin, emodin and sulochrine [131, 137, 139, 156]. This species is also an important human pathogen [88]. Aspergillus terreus was the only species belonging to section Terrei of the Aspergillus genus until recently [53, 128]. Molecular studies have since indicated that this section should be expanded to include a number of other species [121]. Two regions of the ribosomal RNA gene cluster, namely the ITS region and the D1-D2 region of the 26 S rRNA gene were analyzed (Fig. 7) [121, Varga et al., unpublished data]. Phylogenetic analysis of sequence data enabled us to classify the isolates into different clades which mostly correspond to the species A. terreus, A. flavipes, A. niveus, A. carneus and A. janus/A. janus var. brevis. A. allahabadii, A. terreus var. aureus and A. niveus var. indicus belonged to the A. niveus clade, while an Aspergillus isolate previously classified as A. niveus was most closely related to A. flavipes isolates. A. anthodesmis formed a distinct branch on the tree. Although it was previously suggested, based on 28S rDNA sequence data, that Aspergillus section Terrei should include A. carneus and A. niveus isolates, phylogenetic analysis of ITS sequences indicate that A. flavipes isolates are more closely related to A. terreus than A. carneus isolates. Based on these observations, it was suggested merging sections Terrei and Flavipedes. The high bootstrap values supported these findings. As seen, while the D1-D2 sequences of most A. carneus and A. niveus isolates were identical to those of A. terreus isolates, their ITS sequences are more informative, i.e. they allowed the separation of A. terreus, A. carneus and A. niveus isolates (Fig. 7). Lovastatin production of the isolates did not correlate with their positions on any trees based on sequence data, indicating that lovastatin production was lost (or gained) several times during evolution. The species accepted in this section are listed in Table 7. Aspergillus sections Cervini, Candidi and Cremei

These sections mostly include economically less important species, with the exception of A. candidus that is an important spoilage fungus, and A. wentii which is a source of enzyme preparations, and frequently included in ‘koji’ [95, 122, 128]. Section Cervini includes five species, two of which (A. parvulus and A. nutans) are possibly synonyms based on 28 S rDNA sequences (Table 3) [121]. Aspergillus section Candidi was suggested to include three species based on molecular and chemotaxonamic evidence (Table 3) [121, 127, 175]. The three species

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belonging to this section include A. candidus, A. campestris and A. taichungensis [189]. Section Cremei has been studied in detail by Peterson [120]. He transferred several species to this section based on molecular evidence. For example, A. wentii (from section Wentii), A. dimorphicus (section Circumdati), and A. gorakhpurensis (section Versicolores) all now belong to section Cremei. Later, Varga et al. [175] also transferred A. sepultus to this section. The teleomorphic state (Chaetosartorya) is known in three species of this section (Table 3). Aspergillus subgenus Fumigati Aspergillus section Fumigati

Aspergillus section Fumigati includes species characterized by uniseriate, columnar conidial heads in shades of green and flask shaped vesicles [128]. The genus Neosartorya (family Trichocomaceae) was established to accommodate teleomorphs of species belonging to the “Aspergillus fischeri series” of the A. fumigatus species group [128]. This section now involves the anamorphs of more than 20 sexual Neosartorya species, and 5 asexual Aspergilli (Table 7) [136]. Japanese authors are especially active in describing new Neosartorya species from tropical or subtropical soils and herbs [67-71, 143, 148, 149, 157, 158, 188]. The most important species among them is Aspergillus fumigatus Fresenius, which is a ubiquitous filamentous fungus in the environment, and also an important human pathogen [128]. Several Neosartorya species have also been described as causative agents of human diseases including invasive aspergillosis, osteomyelitis, endocarditis and mycotic keratitis [33, 62, 74, 92, 93, 113, 146]. All of the examined Neosartorya species produce heatresistant ascospores which are frequently encountered from different food products [60, 132, 155]. In addition, several mycotoxins are produced by these species, many of which may cause serious health hazards [47, 52, 134]. Although it was suggested that A. fumigatus is able to produce ochratoxins, this observation was not confirmed by other authors [4]. Some species also have valuable properties for mankind; for example, N. fischeri strains produce fiscalins which effectively inhibit the binding of substance P to the human neurokinin receptor [186], while A. fumigatus strains produce pyripyropenes, potent inhibitors of acyl-CoA:cholesterol acyltransferase [154], the immunosuppressant restrictocins [105], ribotoxins [91] and fumagillin, which exhibits amebicidal activity [101]. Phylogenetic analyses of sequences of part of the b-tubulin gene and the D1-D2 region of the 28 S rRNA gene have recently been carried out

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Fig. 7. Neighbor-joining trees of species of Aspergillus section Terrei based on ribosomal RNA gene sequences. a. tree based on sequences of the D1-D2 region of the 28 S rDNA gene; b. tree based on ITS sequences. Numbers above branches indicate bootstrap values; only values >50% are shown. The bar represents genetic distance.

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Table 7. Species assigned to Aspergillus sections Terrei, Flavipedes, Clavati and Fumigati (associated teleomorph is in brackets; modified after Pitt et al. [123]) Section Terrei (Fennellia): A. allahabadii Mehrota & Agnihotri 1963 A. ambiguus Sappa 1955 A. carneus Blochwitz 1933 A. janus Raper & Thom 1944 A. janus var. brevis Raper & Thom 1944 A. microcysticus Sappa 1955 A. niveus var. indicus=A. terreus var. aureus Thom & Raper 1945 A. terreus Thom 1918 F. nivea (Wiley & Simmons) Samson 1979 [A. niveus Blochwitz 1929] Section Flavipedes (Fennellia): A. iizukae Sugiyama 1963 F. flavipes (Bainier & Sartory) Thom & Church 1926 [A. flavipes (Bainier & Sartory) Thom & Church] Fennellia monodii Locquin-Linard 1990 Section Clavati (Neocarpenteles): A. clavatonanicus Batista et al. 1955 A. clavatus Desmazieres 1834 A. giganteus Wehmer 1901 A. ingratus Yaguchi et al. 1993 A. longivesica Huang & Raper 1971 A. rhizopodus Rai et al. 1975 N. acanthosporus Udagawa & Takada 1971 [A. acanthosporus Udagawa & Takada 1971] Section Fumigati (Neosartorya): Teleomorph unknown: A. brevipes Smith 1952 A. brunneouniseriatus Singh & Bakshi 1961 A. duricaulis Raper & Fennell 1965

A. fumigatus Fresenius 1863 A. neofumigatus Hong, Frisvad & Samson 2004 A. unilateralis Thrower 1954 A. viridinutans Ducker & Thrower 1954 Heterothallic: N. fennelliae Kwon-Chung & Kim 1974 N. nishimurae Takada et al. 2001 N. otanii Takada et al. 2001 N. spathulata Takada & Udagawa 1985 N. udagawae Horie et al. 1995 Homothallic: N. aurata (Warcup) Malloch & Cain 1973 N. aureola (Fennell & Raper) Malloch & Cain 1973 N. botucatensis Horie et al. 1995 N. delicata Kong 1997 N. fischeri (Wehmer) Malloch & Cain 1973 N. glabra (Fennell & Raper) Kozakiewicz 1989 N. hiratsukae Udagawa et al. 1991 N. indohii Horie et al. 2003 N. multiplicata Yaguchi et al. 1994 N. paulistensis Horie et al. 1995 N. primulina Udagawa et al. 1993 N. pseudofischeri Peterson 1992 N. quadricincta (Yuill) Malloch & Cain 1973 N. spinosa (Raper & Fennell) Kozakiewicz 1989 N. stramenia (Novak & Raper) Malloch & Cain 1973 N. sublevispora Someya et al. 1999 N. takakii Horie et al. 2001 N. tatenoi Horie et al. 1992 N. tsurutae Horie et al. 2003 “Neosartorya sp.” NRRL 4179

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on this group (Fig. 8). The recently described Neosartorya pseudofischeri was found to be basal to all other species of section Fumigati [55, 119, 177]. Phylogenetic analyses supported the hypothesis that heterothallism is a derived character, and that sexuality was lost several times during the evolution of Aspergillus section Fumigati [55, 177]. The heterothallic N. fennelliae and N. udagawae strains were closely related to the homothallic Neosartorya sp. NRRL 4179 and N. aureola, respectively. Strain FRR 1266, which was earlier classified as a highly divergent A. fumigatus isolate, was found to belong to the A. viridinutans species [174]. An A. viridinutans strain IMI 306135 was most closely related to an asexual strain isolated from cocoa beans (JV3, Fig. 8). These two latter strains were more closely related to A. fumigatus and N. fischeri than to any A. viridinutans strains. These observations were also supported by carbon source utilisation data and restriction analysis of the mitochondrial and nuclear DNA of the strains [130]. Phylogenetic analysis of b-tubulin, calmodulin and ITS sequences led to the description of these isolates together with several Korean strains as a new species, A. neofumigatus [66]. Phylogenetic analysis of b-tubulin, ITS and hydrophobin led to the assignment of these isolates into the recently described new species. A. lentulus [66, Varga et al., unpublished observations]. This species also differs from A. fumigatus in producing auranthine, cyclopiazonic acid, terrein, secalonic acid, and not producing fumitremorgins, fumagillin, fumigaclavins and fumigatin, in contrast with typical A. fumigatus isolates [66]. The results indicate that the presence or absence of nodding conidial heads is not an unequivocal morphological character for the identification of species within Aspergillus section Fumigati. Recently, Hong et al. [66] identified two new species. N. latiniosa and N. coreana related to N. spinosa from soil samples which came from various parts of the world. Besides, several other Neosartorya isolates have been found previously which have characteristic ascospore ornamentations [119, 134]. Molecular analysis of these isolates is needed to clarify their position. Wang et al. [181] examined mitochondrial cytochrome b sequences of isolates belonging to section Fumigati. A neighbour-joining tree based on sequences available in the Genbank database is presented in Fig. 9. The topology of the tree underlines the close relationship between sections Fumigati and Clavati as they do not form separate (monophyletic) clades (A. clavatus was more closely related to section Fumigati species than to those belonging to section Clavati). This

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Fig. 8. Neighbor-joining trees of Aspergillus section Fumigati based on ribosomal RNA gene and >-tubulin gene sequences. a. tree based on sequences of the D1-D2 region of the 28 S rDNA gene; b. tree based on >-tubulin sequences. Numbers above branches indicate bootstrap values; only values >50% are shown. The bar represents genetic distance.

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Fig. 9. Neighbor-joining tree of Aspergillus section Fumigati and Clavati based on mitochondrial cytochrome b sequences. Numbers above branches indicate bootstrap values; only values >50% are shown. The bar represents genetic distance.

phenomenon was also observed by Varga et al. [173] who also found that A. clavatus is closely related to section Fumigati isolates based on isoenzyme and carbon source utilization data.

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Phylogenetic analyses can be misleading in intraspecific comparisons if the examined population is able to recombine [152]. Recently the reproductive mode of A. fumigatus has been examined [176]. Using statistical tests (the index of association test and the parsimony tree length permutation test) on isoenzyme and sequence specific DNA primer analysis data, the results obtained supported the premise that recombination played an important role in A. fumigatus populations. Network methods were also used successfully to visualize the recombining structures of A. fumigatus populations. Inspite of the applied techniques, it was not possible to distinguish between the alternative hypotheses of whether past meiotic exchanges, parasexuality or a cryptic sexual stage were responsible for the recombining population structure of A. fumigatus. The identification of a mating type gene homologue in an A. fumigatus genomic database indicates that past meiotic exchanges could be responsible for the observed nonstructured population [125, 178]. Aspergillus section Clavati

Species belonging to Aspergillus section Clavati are characterized by clavate-shaped vesicles and large blue-green uniseriate conidial heads. Aspergillus clavatus is the most economically important species of the section, which can be isolated mainly from cultivated soil and dung, and also occurs on stored products (mainly cereals) with high moisture content [94]. A. clavatus and its relatives produce a number of mycotoxins including patulin, kojic acid, cytochalasins, and tremorgenic mycotoxins [45]. The genotoxic mycotoxin patulin is receiving worldwide attention due to its frequent occurrence in apple juice [17, 65]. A. clavatus was found to be responsible for an extrinsic allergic alveolitis known as malt worker’s lung, and in cases of mycotoxicoses of animals fed with by-products of malting [45, 94]. The toxic syndromes observed in animals were suggested to result from the synergistic action of these mycotoxins [45]. A. clavatus and A. giganteus isolates also produce ribotoxins, which are promising tools for immunotherapy of cancer [99]. Mycotoxin-producing abilities and phylogenetic relationships among isolates representing the six species currently assigned to this section have been examined [171]. A phylogenetic analysis of ITS sequence data indicated that most isolates belong to two main clades, which have also been identified previously using 28 S rDNA sequence data (Fig. 10)

b.

Fig. 10. Neighbor-joining trees of Aspergillus section Clavati based on ribosomal RNA gene sequences. a. tree based on sequences of the D1D2 region of the 28 S rDNA gene; b. tree based on ITS sequences. Numbers above branches indicate bootstrap values; only values >50% are shown. The bar represents genetic distance.

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[121]. Aspergillus pallidus isolates clustered together with strains of A. clavatus indicating that A. pallidus isolates should be reclassified as A. clavatus. The anamorph of Hemicarpenteles acanthosporus was also found to belong to this section. A new genus, Neocarpenteles was proposed to accommodate this species as Neocarpenteles acanthosporum [160]. Correlation was not observed between patulin production and the taxonomic position of the isolates tested, indicating that patulinproducing abilities were lost several times during the evolution of Aspergillus section Clavati. A new species, A. ingratus suggested to belong to section Clavati, was not examined because the isolate received from the authors could not be cultivated [187]. Patulin-producing abilities of Aspergillus species were examined using analytical (HPLC, TLC) and molecular detection methods. For the latter, a primer pair developed based on iso-epoxydon dehydrogenase (IDH) gene sequences of Penicillium expansum was used [116]. This gene encodes for a key enzyme of patulin biosynthesis [141, 142]. A good correlation was observed between patulin production and the presence of an IDH gene fragment in the isolates. Aspergillus longivesica was found for the first time to produce large amounts of patulin [171]. CONCLUSION AND FUTURE PROSPECTS

In this review, efforts have been made to give an overview of recent developments in the taxonomy of economically important aspergilli. Phylogenetic analysis of sequences of the ribosomal RNA gene cluster and the b-tubulin gene were found to be useful for clarifying the taxonomic relationships of toxigenic Aspergilli causing pre- and postharvest contamination of agricultural products. Molecular data has helped to clarify, at least partially, the taxonomy of black Aspergilli, A. flavus and its relatives, sections Circumdati and Terrei including ochratoxin and patulin-producing species, and the uniseriate sections Aspergillus, Fumigati and Clavati, respectively. Phylogenetically unrelated species were found to be able to produce the same mycotoxins. Further studies are needed for clarifying the taxonomy of sections Aspergillus, Terrei and Flavipedes, and especially that of Aspergillus subgenus Nidulantes not covered in this review. Furthermore, there is an urgent need for an exhaustive compilation of the current knowledge on Aspergillus taxonomy comparable to the work of Raper and Fennell published in 1965. Since then, no such attempts have been made.

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Acknowledgments

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[175] Varga J, Tóth B, Rigó K, Téren J, Hoekstra RF, Kozakiewicz Z. Phylogenetic analysis of Aspergillus section Circumdati based on sequences of the internal transcribed spacer regions and the 5.8 S rRNA gene. Fungal Genet Biol 2000; 30: 71–80. [176] Varga J, Tóth B. Genetic variability and reproductive mode of Aspergillus fumigatus: a review. Infect Genet Evol 2003; 3: 3–17. [177] Varga J, Vida Z, Tóth B, Debets F, Horie Y. Phylogenetic analysis of newly described Neosartorya species. Antonie van Leeuwenhoek 2000; 77: 235–239. [178] Varga J. Mating type gene homologues in Aspergillus fumigatus. Microbiology 2003; 149: 816–819. [179] Vertesy L, Burger HJ, Kenja J, Knauf M, Kogler H, Paulus EF, Ramakrishna NV, Swamy KH, Vijayakumar EK, Hammann P. Kodaistatins, novel inhibitors of glucose6-phosphate translocase T1 from Aspergillus terreus Thom DSM 11247. Isolation and structural elucidation. J Antibiot 2000; 53: 677–686. [180] Vesonder RF, Lambert R, Wicklow DT, Biehl ML. Eurotium spp. and echinulin in feed refused by swine. Appl Environ Microbiol 1988; 54: 830–831. [181] Wang L, Yokoyama K, Miyaji M, Nishimura K. Mitochondrial cytochrome b gene analysis of Aspergillus fumigatus and related species. J Clin Microbiol 2000; 38: 1352–1358. [182] Wang L, Yokoyama K, Takahasi H, Kase N, Hanya Y, Yashiro K, Miyaji M, Nishimura K. Identification of species in Aspergillus section Flavi based on sequencing of the mitochondrial cytochrome b gene. Int J Food Microbiol 2001; 71: 75–86. [183] Whyte AC, Gloer JB, Wicklow DT, Dowd PF. Sclerotiamide: a new member of the paraherquamide class with potent antiinsectan activity from the sclerotia of Aspergillus sclerotiorum. J Nat Prod 1996; 59: 1093–1095. [184] Wicklow DT, Dowd PF, Alfatafta AA, Gloer JB. Ochratoxin A: an antiinsectan metabolite from the sclerotia of Aspergillus carbonarius NRRL 369. Can J Microbiol 1996; 42: 1100–1103. [185] Wolf G. Traditional fermented food. In: Anke T, ed. Fungal biotechnology. Weinheim, Germany: Chapman & Hall, 1997: 3–13. [186] Wong SM, Musza LL, Kydd GC, Kullnig R, Gillum AM, Cooper R. Fiscalins: new substance P inhibitors produced by the fungus Neosartorya fischeri. Taxonomy, fermentation, structures, and biological properties. J Antibiot 1993; 46: 545–553. [187] Yaguchi T, Someya A, Miyadoh S, Udagawa SI. Aspergillus ingratus, a new species in Aspergillus section Clavati. Trans Mycol Soc Japan 1993; 34: 305–310. [188] Yaguchi T, Someya A, Udagawa S. A new species of Neosartorya from Taiwan soil. Mycoscience 1994; 35: 309–313. [189] Yaguchi T, Someya A, Udagawa SI. Aspergillus taichungensis, a new species from Taiwan. Mycoscience 1995; 36: 421–424. [190] Yokoyama K, Wang L, Miyaji M, Nishimura K. Identification, classification and phylogeny of the Aspergillus section Nigri inferred from mitochondrial cytochrome b gene. FEMS Microbiol Lett 2001; 200: 241–246. [191] Yu TS, Yeo SH, Kim HS. A new species of Hyphomycetes, Aspergillus coreanus sp. nov. isolated from traditional Korean Nuruk. J Microbiol Biotechnol 2004; 14: 182–187. [192] Zotti M, Corte AM. Aspergillus persii: a new species in section Circumdati. Mycotaxon 2002; 83: 269–278.

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Polymerase Chain Reaction-based Methods in Fusarium Taxonomy YOUSSUF A.M.H. GHERBAWY1, MOHAMED A. EL-NAGHY2 and ABDULLAH ALTALHI 3 1

Botany Department, Faculty of Science, South Valley University, Qena, Egypt 2 Botany Department, Faculty of Science, El Minia University, El Minia, Egypt 3 Biology Department, Faculty of Science, Taif University, Taif, Saudi Arabia

ABSTRACT Fusarium species are ubiquitous fungi, infecting a wide range of crop plants causing diseases which result in significant quantitative and qualitative losses to global agriculture. Some fusaria are also known to produce deleterious mycotoxins resulting in toxin contamination of crop products. Outbreaks of diseases caused by these fungi pose a great problem for the agricultural industry and there is need for rapid and accurate identification methods for implementation of the control measures of diseases caused by these fungi. The accurate identification of Fusarium species has always been problematic because of the contradictory classification systems proposed by various researchers, based primarily on cultural and morphological characters that could be highly variable depending on the media and cultural conditions used. In addition, degeneration of cultures, as well as the production of mutants, may further add to the problems of fungal identification and diagnosis. In recent years, the increasing use of molecular methods in fungal diagnostics has emerged as a possible means to overcome the problems associated with the existing phenotypic identification systems. This chapter focuses on the most 1 Address for correspondence: Dr. Youssuf A.M.H. Gherbawy, Botany Department, Faculty of Science, South Valley University, Qena 83523, Egypt. Tel: +0020965335352, Fax: +0020965211279, E-mail: [email protected].

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famous molecular tools that are used by researchers for Fusarium diagnostic, taxonomic and phylogenetic studies. Key Words: AFLP, DGGE, microsatellites, rRNA sequencing, RAPD, RFLP, ITS regions

INTRODUCTION

Fusarium is a large cosmopolitan genus of pleoanamorphic hyphomycetes whose members are responsible for a wide range of plant diseases [41], production of certain mycotoxins, that cause mycotic infections of humans and other animals [104]. Collectively the fusaria represent the most important phytopathogen and mycotoxigenic genus of the filamentous fungi [86]. In addition to producing a wide range of toxins, including trichothecenes and fumonisins, the fusaria are noted for their production of other secondary metabolites such as gibberellin plant growth hormones, named after Gibberella fujikuroi from which they were first discovered. TAXONOMY

The high level of biodiversity or variation within the genus has led to considerable difficulties in the taxonomic treatment of the genus. The ideal taxonomic system should reflect the genetic relationships among taxa. Some believe that it should also recognize, at an appropriate level, taxa which are distinguished practically by significant aspects of their pathogenicity, toxigenicity or ecological characters without unnecessary splitting of genetically unified populations. The history of Fusarium systematics has shown marked swings between excessively narrow species concepts and those which are so broad that practically useful information on pathogenicity and toxigenicity has been lost. Contemporary studies of biodiversity in Fusarium, which have led to the description of several new specific and infraspecific taxa, are based on the examination of large populations of isolates in which traditional morphological criteria are integrated with detailed data on pathogenic specialization, toxin production and ecology, and more recently with information derived from molecular taxonomic studies. Fusarium is one of the most challenging genera for morphological systematists [44]. Traditional classification and identification schemes are based exclusively on morphological species concept derived from cultural characteristics of single-spore isolates grown on special media, shared morphological traits of the anamorph, host range, and to a lesser extent,

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teleomorph micromorphology [17]. Given the paucity and plasticity of phenotypic traits, it is not surprising that conflicting morphological species concepts are employed in taxonomic treatments of this genus [17, 51, 105], For this reason, the systematics of Fusarium remain controversial and confusing. Gerlach and Nirenberg’s system [51] is the most differentiated, including 73 species and 26 varieties; Booth [17] recognized 44 species and 7 varieties and Nelson et al. [105] only 30 species. Most Fusarium isolates studied by mycotoxicologists and plant pathologists in the first three quarters of the 20th century were initially identified incorrectly using one of several over-simplified morphological systems. For example, the name of the toxin nivalenol was based on the misidentification of an isolate producing it as Fusarium nivale. Since then, F. nivale has been moved to a new genus, Microdochium, and the nivalenol producer was reidentified first as F. tricinctum and F. sporotrichioides by two different researchers using morphology, and finally as a new species, F. kyushuense, using molecular phylogenetics [3]. Marasas et al. [86] made a huge effort to correctly identify hundreds of toxigenic isolates associated with the production of various mycotoxins. These authors debunked and confirmed many reports of mycotoxin production associated with particular Fusarium species, but their efforts could only reflect the current state of the art of morphological species recognition. They recognized 30 Fusarium species, and while it served as an advance over the previous 9 species system as described by Snyder and Hansen [126, 127, 128], it too vastly under-estimated the underlying phylogenetic diversity and grouped species into sections that do not reflect evolution. Nearly 20 years later, we found overselves again needing to correlate previous reports of mycotoxin production with phylogenetically defined species, such that identifying toxigenic isolates still remains an enormous challenge. Species newly identified using phylogenetic methods are generally difficult or impossible to identify using conventional morphological traits [4]. The most important stimulus to molecular research so far has been the exploitation of DNA polymerases for in vitro amplification of DNA, i.e. the polymerase chain reaction (PCR). PCR-based technology has revolutionized DNA research by making it widely accessible, rather than the domain of the few, as was previously the case. In general terms, PCR is having a qualitative impact by allowing molecular researchers to generate new forms of information that were previously prohibitively difficult to obtain. Secondly, PCR has a quantitative effect by increasing the number of studies generating DNA-based data. Thirdly, PCR has

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increased the range of organisms subjected to DNA analysis, thereby improving the diversity of the dataset. The potential for exploiting PCRbased methods in Fusarium research is enormous, both in applied work and systematics. In systematic studies, it is important to make a clear distinction between the procedures of classification and identification. Molecular methods should allow significant improvement in the existing classification system for Fusarium and progress towards resolving some longstanding issues, including: the phylogenetic relationship between Fusarium and other genera; the generic concept in Fusarium; sectional relationships; species concepts; population structure analysis and delineation of infra-specific taxa; mechanisms of evolutionary change in Fusarium populations; mechanisms of culture instability. PCR-BASED METHODS

The polymerase chain reaction (PCR) for amplification of specific nucleic acid sequences was introduced by Saiki et al. [119], and has subsequently proved to be one of the most important scientific innovations of the past decade. PCR uses a thermostable polymerase to produce multiple copies of specific nucleic acid regions quickly and exponentially, including noncoding regions of DNA as well as particular genes. For example, starting with a single copy of a 1 KB DNA sequence, 1011 copies (or 100 ng) of the same sequence can be produced within a few hours. Once the reaction has occurred, a number of methods for identification and characterization of the amplification products according to their size following migration on agarose gels were used. The standard method requires a DNA template containing the region to be amplified and two oligonucleotide primers flanking this region. The amplification is based on the use of a thermostable DNA polymerase isolated from Thermus aquaticus, called taq polymerase [118]. All PCR reaction components are mixed and the procedure consists of a succession of three steps which are determined by temperature conditions: template denaturation, primer annealing and extension. In the first step, the incubation of the reaction mixture at a high temperature (90-95°C) allows the denaturation of the double-stranded DNA template. By cooling the mixture to an annealing temperature typically around 55°C, the target-specific oligonucleotide primers anneal to the 5’ end of the two single-stranded templates. For the extension step, the temperature is raised to 72°C. The time of incubation for each

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step is usually 1-2 minutes. This sequence of denaturation, annealing and extension generates newly synthesized DNA strands which are separated from the original strands by denaturation and each strand serves again as template in the annealing and extension steps. Theoretically, n cycles of PCR allow a 2n-fold amplification of the target DNA sequence. 1. Random Amplified Polymorphic DNA (RAPD-PCR)

The random amplified polymorphic DNA (RAPD) or arbitrarily primed polymerase chain reaction (AP-PCR) fingerprinting assay detects small inverted nucleotide sequence repeats through genomic DNA [143, 145]. In RAPD-PCR, amplification involves only single primers of arbitrary nucleotide sequence. The principle of RAPD assays is discussed in detail by Hadrys et al. [62] and Tingey and del Tufo [134]. In brief, a single oligonucleotide of an arbitrary DNA sequence is mixed with genomic DNA in the presence of a thermostable DNA polymerase and a suitable buffer, and then is subjected to temperature cycling conditions typical of the polymerase chain reaction. At an appropriate annealing temperature during the thermal cycle, the single primer binds to sites on opposite strands of the genomic DNA that are at an amplifiable distance of each other (for example, within a few thousand nucleotides), and a discrete DNA segment is produced. The presence or absence of this specific product, although amplified with an arbitrary primer, will be diagnostic for the oligonucleotide-binding sites on the genomic DNA. In practice, the DNA amplification reaction is repeated on a set of DNA samples with several different primers, under conditions that result in several amplified bands from each primer. The amplified products are visualized on an agarose gel, polymorphic bands are noted, and the similarity/ genetic distance values can be calculated, for example, among the fungal strains investigated. Amplification conditions should be well standardized, because the fragment pattern depends on the concentration of magnesium ions, Taq-DNA polymerase, primer and template DNA, and the annealing temperature. The primers designed for this application are required to be 9-15 nucleotides in length, between 50 and 80% G+C in composition. Amplification with RAPD primers is extremely sensitive to singlebase changes in the primer-target site. This feature suggests that RAPDPCRs should be highly useful for phylogenetic analysis among closely related individuals, but less useful for analysis of genetically diverse

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individuals. RAPD primers are useful for distinguishing species, for discriminating between different isolates of the same species, and to measure similarity among individuals within natural or artificial (through breeding) populations within a species. The identification of shared characters in this analysis is limited to the resolution of the agarose gel. In fact, many more bands are present among the amplification products than are detected by ethidium bromide staining. Caetano-Annoles et al. [24] were able to detect over 100 bands amplified with a single random primer, by resolving the reaction products on a polyacrylamide gel and staining with silver. RAPD offers several advantages that may be useful in studying formae specialis and races of Fusarium oxysporum, to identify RAPD markers for formae specialis and race identification [52, 57]. RAPD reduces the time needed for race identification in diseased plants, and provides genetic information on isolates studied, allowing for fingerprinting of isolates [52, 53, 142, 143]. Random PCR approaches are being increasingly used to generate molecular markers which are useful for taxonomy and for characterizing fungal populations. The main advantage of these approaches is that previous knowledge of DNA sequences is not required, hence any random primer can be tested to amplify any fungal DNA. RAPD primers are chosen empirically and tested experimentally to find RAPD banding patterns which are polymorphic between the taxa studied. These advantages also include rapidness, quickness, and small amount of template DNA needed. RAPD-PCR assays have been used extensively to define fungal populations at species, intraspecific, race and strain levels. In general, most studies have concentrated on intraspecific grouping, although others have been directed at the species level. Some examples of RAPDPCR at species level include the production of species-specific probes and primers from RAPD data for Fusarium oxysporum f. sp. dianthi: Phytophthora cinnamomi, Tuber magnatum, Glomus mosseae, Fusarium sambucinum, members of Fusarium section Fusarium, and Fusarium oxysporum [34, 52, 55, 63, 77, 78, 85, 112, 152]. Kelly et al. [69] by using the cluster analysis of RAPD data divided F. oxysporum f. sp. ciceris into two distinct clusters that correlated with the pathotypes causing the yellowing or wilt disease syndrome of chickpea. These clusters were clearly distinct from other F. oxysporum formae speciales and from other chickpea pathogens such as F. eumartii and F. solani. Gherbawy et al. [54] used RAPD techniques to differentiate between isolates of F. subglutinans,

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F. proliferatum and F. verticillioides. Their results indicated that these fusaria are distinct species and hence RAPD markers can be quick and reliable for differentiating them. Also, RAPD analysis provided markers to differentiate races A, 3 and 4 of F. oxysporum f. sp. vasinfectum [7], races 0, 2 and 1,2y of F. oxysporum f. sp. Melonis [103] and races 0, 1B/ C, 5, and 6 of F. oxysporum f. sp. ciceris [68]. Bentley et al. [15] reported that the comparison of the RAPD banding patterns, both visually and by phonetic analysis, sub-divided isolates of F. oxysporum f. sp. cubense into two major groups; there was no correlation between RAPD banding pattern and race. Assigbetse et al. [7] reported that the RAPD method revealed polymorphisms within isolates of Fusarium oxysporum f.sp. vasinfectum and established DNA fingerprints useful for race characterization. They differentiated the isolates into three main groups (RAPD I, II and III) directly related to both virulence and geographic origin. RAPD analysis conducted on Fusarium oxysporum f. sp. pisi isolates allowed differentiation of only one race (race 2) of four races [57]. Fernandez et al. [42] reported that there is no variation in the mtDNA and RAPD analysis of Algerian population of Fusarium oxysporum f. sp. albedinis, indicating that they were genetically very closely related. Möller et al. [98] created a specific primer for F. moniliforme and F. subglutinans based on sequences of RAPD fragments. Yli-Mattila and Hyvönen [148] reported that the strains of both F. oxysporum and F. redolens are nested within the large F. avenaceum clade. In their morphological studies Nelson et al. [105] and Baayen and Gams [8] regarded F. redolens conspecific with F. oxysporum, while Gerlach and Nirenberg [51] considered them as distinct species. Cramer et al. [32] reported identical RAPD banding patterns in F. oxysporum f. sp. phaseoli isolates suggesting low genetic diversity in the region studied. F. oxysporum f. sp. betae isolates showed a greater degree of diversity, but in general clustered together in a grouping distinct from F. oxysporum f. sp. phaseoli isolates. As RAPD markers revealed such a high level of genetic diversity across all isolates examined, their study concluded that RAPD markers had only limited usefulness in correlating pathogenicity among the isolates and races. Chiocchetti et al. [29] identified Fusarium oxysporum f. sp. basilica isolated from soil, basil seed, and plants by RAPD analysis. They found that 30 of the F. oxysporum f. sp. basilica isolates obtained from soil or wilted plants gave identical amplification patterns using 31 different random primers. All tested primers allowed clear differentiation of Fusarium oxysporum f. sp. basilica from representatives of other formae

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speciales and from nonpathogenic strains of Fusarium oxysporum. Two isolates of F.redolens from Japan and seven nonpathogenic isolates of F. proliferatum collected from diseased carnations in Italy, Israel and The Netherlands were clearly distinguishable according to their RAPD fingerprint [95]. RAPD method was used to differentiate and identify Group 1 and Group 2 strains of Fusarium graminearum [98, 120, 121]. Aoki and O’Donnell [3] characterized F. pseudogarminearum sp. nov., formerly recognized as the group 1 populations of F. garminearum. The amount of genetic variation in 32 isolates of Fusarium oxysporum f. sp. lentis was evaluated by PCR amplification with a set of 6 RAPD primers and all amplifications revealed scorable polymorphisms among the isolates, and a total of 8 polymorphic fragments were scored [14]. Gherbawy and Prillinger (unpublished data) during their work with molecular identification for different Fusarium strains isolated from some Austrian plants reported that (i) Fusarium colmurum strains were homogenous species according to the RAPD patterns. Therefore, there was no problem in making correct identification of this species by using RAPD analysis with the aid of type strain (Fig. 1); and (ii) Fusarium poae strains were separated into two distinct groups according to RAPD patterns (Fig. 2), one of those two groups gained new name as Fusarium langsethiae by Trop and Nirenberg [136].

Fig. 1. Random amplification patterns obtained using Fusarium culmorum, printed by M13 oligonucleotide (GAGGGTGGCGGTTCT).

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Fig. 2. Random amplification patterns obtained using Fusarium poae, printed by M13 oligonucleotide (GAGGGTGGCGGTTCT).

2. Restriction Fragment Length Polymorphism (RFLP)

Restriction fragment length polymorphism (RFLP) is a technique in which organisms may be differentiated by analysis of patterns derived from cleavage of their DNA. If two organisms differ in the distance between sites of cleavage of a particular restriction endonuclease, the length of the fragments produced will differ when the DNA is digested with a restriction enzyme. The similarity of the patterns generated can be used to differentiate species (and even strains) from one another. Restriction endonucleases are enzymes that cleave DNA molecules at specific nucleotide sequences depending on the particular enzyme used. Enzyme recognition sites are usually four to six base pairs in length. Generally, the shorter the recognition sequence, the greater the number of fragments generated. If molecules differ in nucleotide sequence, fragments of different sizes may be generated. The fragments can be separated by gel electrophoresis. Restriction enzymes are isolated from a wide variety of bacterial genera and are named by using the first letter of the genus, the first two letters of the species, and the order of discovery.

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RFLP is made on mitochondrial as well as nuclear ribosomal DNA. Since rDNA occurs as multiple copies, it lends itself to analysis based on RFLP. The patterns obtained are sufficiently different to allow recognition of individual species, as well as individual strains of a species. Consequently, the method has considerable diagnostic value. The results from RFLP are absolutely reproducible and thereby suitable for databases [90]. The estimates of evolutionary relationships from RFLP patterns would be expected to be less accurate than estimates derived from sequence comparisons because as evolutionary distances increase, the extent of pattern similarities becomes less certain [71]. For fungal mitochondrial genomes, which are usually small (19-121 kb) and exist in relatively high copy number [58], RFLPs are usually easy to detect. Total DNA of the fungus is extracted and the mitochondrial DNA separated from nuclear DNA by centrifugation in CsCl gradients containing bis-benzimide [46]. The relatively small size of mitochondrial genomes results in a simple banding pattern when cut with restriction endonucleases. Mitochondrial RFLP markers have been used in a number of fungi although the extent of variation differs considerably. Variation in mitochondrial (mt) DNA has been used to assess inter- and intra- species relationships in various fungi (reviewed by Taylor et al. [133]). RFLPs of mtDNA have been shown to be useful genetic markers for estimating genetic divergence and phylogenetic relationships between natural populations in various taxa [43, 67, 70, 125, 133]. RFLP [106, 149] and RAPD-PCR analyses [151] made it possible to distinguish nearly all F. avenaceum isolates from each other. RFLP of amplified fragments (PCR-RFLP) analysis has been used to study fungal population biology, such as the differentiation of species, species forms and isolates [25, 87]. This method has often utilized the analysis of nuclear ribosomal DNA (nrDNA) sequences that are found in all eukaryotic cells and contain both regions and showed substantial resolution at different taxonomic units in a majority of fungi including Fusarium species [5, 16, 18, 35, 37, 38, 65, 75, 96, 106, 140]. PCR-RFLP is a simple and inexpensive method compared with the traditional RFLP or sequence analyses as it avoids the need for blotting, probing and/or sequencing. Correll et al. [31] used RFLP to study genetic diversity in California and Florida populations of the pitch canker fungus Fusarium subglutinans

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f. sp. Pini. Appel & Gordon [5] used RFLP method to study the intraspecific variation within populations of Fusarium oxysporum (56 isolates F. oxysporum f. sp. melonis and nonpathogenic strains, were chosen from a larger collection to represent diversity in vegetative compatibility groups (VCGs), mitochondrial DNA (mtDNA) haplotype, geographic distribution, and virulence). Using PCR, a 2.6-kb fragment including the intergenic spacer (IGS) region of the ribosomal DNA was amplified from each isolate. The enzymes EcoRI, Sau3A, CfoI, and AvaII, cut this fragment differentially, revealing 5, 6, 6, and 7 patterns, respectively. Among the 56 isolates, a total of 13 unique IGS haplotypes were identified. Among most F. o. melonis isolates, IGS haplotype correlated with VCG and mtDNA haplotype, but did not differentiate among races. Overall, the IGS haplotype was more variable among the nonpathogenic F. oxysporum VCGs among which 12 of the 13 IGS haplotypes were found. Nonpathogenic isolates that shared a common mtDNA haplotype, but were associated with different VCGs, often had different IGS haplotypes [5]. Chakrabarti et al. [26] reported that, EcoRI restriction pattern of the nuclear ribosomal DNA of four isolates of Fusarium oxysporum f.sp. ciceris representing four races prevalent in India indicated that the races could be grouped into three distinct groups; races 1 and 4 representing one group, and race 2 and race 3, the other two. The restriction pattern indicated presence of three EcoRI sites on the nuclear rDNA of this species, one each on the 5.8S and the 25S regions, conserved to all, and the other one, i.e. the variable site, on the IGS region of the nuclear rDNA. The same was confirmed by PCRamplification of the IGS region followed by digestion with EcoRI and a set of other enzymes. It is suggested that amplification of the IGS region and digestion with restriction enzymes could be used to study polymorphism in Oxysporum f.sp. ciceris, and to rapidly identify the races existing in India. 3. Ribosomal RNA (rRNA) and Ribosomal DNA (rDNA) Sequence Comparisons

Sequence comparisons of ribosomal RNA (rRNA) and its template ribosomal DNA (rDNA) have been used extensively to assess both close and distant relationships among many kinds of organisms. The interest in rRNA/rDNA comes from two important properties. First; ribosomes are present in all cellular organisms and appear to share a common

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evolutionary origin, thus providing a molecular history shared by all organisms; second; some rRNA/rDNA sequences are sufficiently conserved that they are homologous for all organisms and serve as a reference point that enables alignment of the less conserved areas used to measure evolutionary relationships [71]. Sequences of rDNA are often used for taxonomic and phylogenetic studies because they are found universally in living cells and their evolution might reflect the evolution of the whole genome. These sequences also contain both variable and conserved regions, allowing the comparison and discrimination of organisms at different taxonomic levels. The nuclear rDNA in fungi is organized as an rDNA unit which is tandemly repeated. One unit includes three rRNA genes: the small nuclear (18S-like) rRNA, the 5.8S rRNA, and the large (28S-like) rRNA genes. In one unit, the genes are separated by two internal transcribed spacers (ITS1 and ITS2). The 18S rRNA evolves relatively slowly and is useful for comparing distantly related organisms whereas the non-coding regions (ITS) evolve faster and are useful for comparing fungal species within a genus or strains within a species. Some regions of the 28S rDNA are also variable between species. Ribosomal RNA occurs in several size classes in eukaryotes. The genes coding for large-subunit (25S to 28S), small-subunit (16S to 18S), and 5.8 rRNAs occur as tandem repeats with as many as 100 to 200 copies. The separately transcribed 5S r RNA gene may also be included in the repeats [45]. Each of the RNA size classes has been examined for extent of phylogenetic information present. The first rRNA to be sequenced was the 5S rRNA. Due to the conserved nature and small size (ca. 120 nucleotides) of 5S rRNAs, their sequences were easily determined and have been widely used for estimating broad phylogenetic relationships [64]. The 5.8S rRNA sequence analysis was not often used in the studies of yeast and fungal phylogeny. Its sequence includes only about 160 nucleotides and does not offer much more information than the 5S rRNA. Besides that, the 5.8S rRNA has modified nucleotides and is therefore less prone to sequencing. The larger size of small-subunit rRNAs (ca. 1800 nucleotides) and large-subunit rRNA (ca. 3200 nucleotides) compared with 5S and 5.8S rRNAs provide an opportunity for greater resolution of phylogenetic relationships. Despite recent advances in sequencing technology, the

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determination of complete sequences is laborious, and only a few hundred complete sequences are available for either the large subunits or the small subunits [33, 61]. Partial sequences are relatively easy to determine and form the basis for most phylogenetic comparisons. It was demonstrated that partial sequences of rRNAs reveal essentially the same phylogenetic relationships as complete sequences [88, 76]. Comparisons of sequences from a wide variety of organisms have shown that for both large- and small-subunit rRNAs, different regions have different rates of nucleotide substitution [33, 61]. Due to this, the molecules are regarded as a collection of chronometers, with each region offering a different glimpse of the evolutionary history of the organism [146]. The compilation of large-subunit RNA sequences by Gutell and Fox [61] demonstrated that the 5’ end of the molecule is quite variable and is of potential use for detection of the closely related species. With a few exceptions, the D2 region (25S-635) is sufficiently variable to recognize ascomycetous and basidiomycetous yeast species, including most sibling pairs [71]. Conspecific strains ordinarily show 0-1% nucleotide divergence, and closely related species differ by 1-5% [72, 73, 81, 89, 115, 116]. For this region, species regarded as congeneric showed 3-23% nucleotides divergence [74]. According to Peterson and Kurtzman [116], the 18S rRNA gene is the most conserved ribosomal gene, hence the small-subunit rRNA is used for estimating broad phylogenetic relationships. Given the wide divergence in taxonomic opinions reflected in the competing morphologically-based systems of classification, a molecular systematic approach based on discrete DNA sequence data offers considerable promise in the establishment of an objective, phylogenetically-based system of classification of Fusarium and its teleomorphs [22]. Previous molecular systematic studies on the genus have demonstrated the utility of partial nuclear large subunit (LSU) 28S rDNA sequence comparisons for phylogenetic analysis [60, 107, 117]. Taxon sampling in these studies, however, was limited and the level of resolution obtained using LSU 28S rDNA sequence data was insufficient, for example, to distinguish the biological species within section Liseola and to apply a phylogentic species concept [108, 110]. O’donnel [108] amplified and sequenced specific regions of the genome from a large number of taxa to test the morphologically-based taxonomic hypotheses developed for the Fusaria. He had developed a DNA sequence database

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from the following four loci for some species of Fusarium: beta tubulin gene introns/exons, mitochondrial small subunit (mtSSU) rDNA, nuclear large subunit (LSU) rDNA, and nuclear rDNA transcribed spacer (ITS) region. Another approach was the development of primers specific for mycotoxigens produced by Fusarium spp. to identify the toxin producing strains of fusaria [39]. Much of the motivation for the streamlined morphology-based Fusarium taxonomic systems of the mid-to late-20th century came from a desire to make identification simple and reliable, and in hindsight at the cost of over-simplification. There is a need to apply the same tools to provide reliable species identification in the laboratory. Phylogenetic species recognition in Fusarium has relied mostly on Genealogical Concordance Phylogenetic Species Recognition (GCPSR; Taylor et al. [132]), a method which identifies shared partitions among multiple gene genealogies as landmarks for species boundaries. While two or more gene genealogies are required for this method of recognizing species, in many instances a species could be identified accurately using a single DNA sequence marker, as long as background phylogenetic analyses have been performed using the marker along with others, thereby validating its diagnostic utility. The markers of choice for species-level phylogenetics in fungi are intron-rich portions of protein-coding genes [50]. These gene regions tend to evolve at a rate higher than that which is observed at the species level in more commonly applied markers, such as the internal transcribed spacer (ITS) regions of the nuclear ribosomal RNA gene repeat [109, 111]. Moreover, many fusaria within the Gibberella clade possess non-orthologous copies of the ITS2,which can lead to incorrect phylogenetic inferences [110, 111]. The translation elongation factor 1a (TEF) gene, which encodes an essential part of the protein translation machinery, has high phylogenetic utility because it is (i) highly informative at the species level in Fusarium; (ii) non-orthologous copies of the gene have not been detected in the genus; and (iii) universal primers have been designed that work across the phylogenetic breadth of the genus. This gene was first used as a phylogenetic marker to infer species-and generic-level relationships among Lepidoptera [30, 97]. Primers were first developed in the fungi to investigate lineages within the F. oxysporum complex [113]. The ef1 and ef2 primers were designed based on sites shared in exons between Trichoderma reesei (Hypocreales/ Sordariomycetes/Pezizomycotina/Ascomycota) and Histoplasma capsulatum (Eurotiales/Eurotiomycetes/Pezizomycotina/Ascomycota), and they

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can be applied to a wide variety of filamentous ascomycetes. These primers amplify a ~700 bp region of TEF, flanking three introns that total over half of the amplicon’s length, in all known fusaria. This gene appears to be consistently single-copy in Fusarium, and it shows a high level of sequence polymorphism among closely related species, even in comparison to the intron-rich portions of protein-coding genes, such as calmodulin, beta-tubulin and histone H3. For these reasons, TEF has become the marker of choice as a single-locus identification tool in Fusarium. Fusarium subglutinans f. sp. pini (= F. circinatum) is a pathogen of pine and is one of eight mating populations (i.e. biological species) in the Gibberella fujikuroi species complex. This species complex includes F. thapsinum, F. moniliforme (= F. verticillioides), F. nygamai, and F. proliferatum, as well as F. subglutinans associated with sugarcane, maize, mango, and pineapple. Differentiating these forms of F. subglutinans usually requires pathogenicity tests, which are often time-consuming and inconclusive. Steenkmap et al. [131] sequenced the histone H3 gene from a representative set of Fusarium isolates. The H3 gene sequence was conserved and contained two introns in all the isolates studied. From both the intron and the exon sequence data, they developed a PCRrestriction fragment length polymorphism technique that reliably distinguishes F. subglutinans f. sp. pini from the other biological species in the G. fujikuroi species complex. ITS regions

The internal transcribed spacers (ITS) are noncoding regions of DNA sequence that separate genes coding for the 28S, 5.8S, and 18S ribosomal RNAs. These ribosomal RNA (rRNA) genes are highly conserved across taxa while the spacers between them may be species-specific. The conservation of the rRNA genes allows for easy access to the ITS regions with “versatile” primers for polymerase chain reaction (PCR) amplification. The variation in the spacers has proved useful for distinguishing among a wide diversity of difficult-to-identify taxa. The ITS region is subdivided into the ITS1 region, which separates the 18S and 5.8S rRNA genes, and the ITS2 region, which is found between the 5.8S and 28S rRNA genes [66]. The highest sequence variation can be obtained from the rapidly evolving internal transcribed spacers. They are recommended for analysis

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of closely related species and may show variation within a single species [91]. The development of PCR and the design of primers for the amplification of the various rDNA regions have considerably facilitated taxonomic studies of fungi [144]. These primers were designed from conserved regions, allowing the amplification of the fragment they flank in most fungi. The ITS primers designed by White et al. [129] have enabled the determination of many ITS sequences from different fungi and these have been used to investigate taxonomic and phylogenetic relationships between species within different genera [80, 82, 129]. ITS sequences are generally constant, or show little variation within the species but vary between species in the genus, and so these sequences have been widely used to develop rapid procedures for the identification of fungal species by PCR-RFLP analysis [27, 37, 38, 137] and to design species-specific primers [13, 99, 130]. The ITS region is a particularly useful area for molecular characterization studies in fungi for four main reasons: (1) The ITS region is relatively short (500-800 pb) and can be easily amplified by PCR using universal single primer pairs that are complementary to conserved regions within the rRNA subunit genes [144]. (2) The multicopy nature of the rDNA repeat makes that ITS region easy to amplify from small, dilute or highly degraded DNA samples [48]. (3) The ITS region may be highly variable among morphologically distinct species [12, 28, 47, 48, 49, 80] and so ITS-generated RFLP restriction data can be used to estimate genetic distances and provide characters for systematic and phylogenetic analysis [22]. (4) PCRgenerated ITS species-specific probes can be produced quickly, without the need to produce a chromosomal library [129] and many researchers have selected sequences from ITS regions to develop species-specific probes because the sequences occur in multiple copies and tend to be similar within and variable between fungal species. The genus Fusarium is heterogeneous and the identification of its individual species is based on morphological or biochemical criteria which can in some cases be difficult and confusing. Edel et al. [38] differentiated several strains of F. oxysporum at the species level by RFLP analysis of a region of ITS and a variable domain of the 28S rDNA. Schilling et al. [121] evaluated sequence variation in the ITS regions of F. avenaceum, F. culmorum and F. graminearum in order to distinguish between the three species. They found that ITS sequences of F. culmorum and F. graminearum were not polymorphic enough to allow the

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construction of species-specific primers; however, sufficient sequence variation was found in the ITS1 and ITS2 regions of F. culmorum and F. graminearum to distinguish them from F. avenaceum. Waalwijk et al. [141] and O’Donnell and Cigelnik [110] have found differences between the morphological and molecular classification of several Fusarium species including F. oxysporum and F. redolens isolates based on ITS2 sequences. ITS sequence analysis of rDNA has shown that F. oxysporum and F. redolens are not conspecific [141]. They have suggested that this incongruity may be due to the hypothetical coexistence of several ITS types in the ancestral Fusarium species, which would mean that ITS2 groupings do not accurately reflect the phylogeny of Fusarium species. This might also explain the difference between rDNA sequence (based on ITS2 and 28s rDNA and on ITS1, 5.8s rDNA and ITS2). Using the latter method, Paavanen-Huhtala et al. [114] supported the morphological classification according to which F. oxysporum and F. redolens were grouped together (section Elegans), while F. avenaceum formed a monophyletic group (section Athrosporiella/Roseum). Waalwijk and Baayen [139] and Waalwijk et al. [140, 141] have demonstrated that F. oxysporum and F. redolens can be distinguished unambiguously on the basis of sequence difference in the ITS2 region of the rDNA. Using ITS criterion, six of the vegetative compatibility groups associated with Fusarium wilt of carnation could be identified as belonging to F. oxysporum f. sp. dianthi, while four belonged to F. redolens f. sp. dianthi [9]. Bao et al. [10] during their studies on genetic analysis of pathogenic and nonpathogenic Fusarium oxysporum from tomato plants using ITS regions sequences, showed that ITS region analysis grouped the strains into four clusters. The nonpathogenic F. oxysporum strains were in two groups, and the pathogenic strains were placed in two different groups. Due to the morphological similarities, F. langsethiae has long been considered to be a “powdery” variant of Fusarium poae. However, the mycotoxin production pattern of F. langsethiae is very similar to that of Fusarium sporotrichioides and produces type A trichothecenes such as T2 toxin [2, 79, 83, 135]. Distinguishing F. langsethiae from the other species in Fusarium section Sporotrichiella is currently based on morphological characters and with mycotoxin analysis [135]. However, these methods are time-consuming, labour-intensive and need specialists. Recently Yli-Mattila et al. [150] investigated the molecular phylogeny of F. langsethiae and other members of section Sporotrichiella by ITS1 and 2,

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intergenic spacer region (IGS), and >-tubulin (tub1) gene sequence analysis. This study also showed that the sequence of a fragment of tub1, while only leading to incomplete phylogenetic resolution, was best suited to distinguish these species. However, the nucleotide differences between various strains were not suitable for RFLP analysis. 4. Micro- and Minisatellites

Micro- and minisatellites are short tandem repeats found scattered through the genome. Microsatellite DNA consists of only 2-10 pb, whereas minisatellite DNA is made up of 15-30 bp [92]. The method developed to detect micro- and minisatellite DNA includes digestion of genomic DNA with a suitable restriction enzyme, separation of fragments by gel electrophoresis, blotting on a membrane, hybridization of multiple polymorphic restriction fragments with a labeled probe and visualization by autoradiography. Different combinations of DNA probes and restriction enzymes are used to obtain informative DNA fingerprints. Microsatellite-primed PCR has been demonstrated to be adapted for investigations at the intraspecific level. In ascomycetous yeasts, it allowed characterization of individuals [59, 92, 123, 143]. Meyer et al. [92] used the oligonucleotide probes (CT)8, (GTG)5, (GACA)4 and the core sequence of the phage M13 (GAGGGTGGCGGTTCT) in combination with the enzymes EcoRI, ApaI, HinfI and Sma for fingerprinting 11 strains of Penicillium, Aspergillus and Trichoderma. The probe (GACA)4 produced good fingerprints only in the genomic DNA of A. niger, which indicated that DNAs of Trichoderma and Penicillium contained only a low amount of the simple tandem sequences homologous to (GACA)4 . The other probes in combinations with the restriction enzymes produced informative fingerprints. The authors concluded that DNA fingerprinting based on the hypervariable minisatellite regions is a powerful method to differentiate species and strains of filamentous fungi. Single, simple repetitive primers have been designed to amplify the microsatellite regions of fungal chromosomal DNA [19, 94, 122]. In most applications these primers have given similar levels of specificity to those seen with RAPD, hence results have been used to group fungi at intraspecific levels [20]. However, in some instances microsatelliteprimed PCR has been used to generate species-specific patterns, and the good example of this is the work on morels by Buscot et al. [23] who

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found considerable homogeneity from both mono- and polysporic isolates of individual species. Further repetitive sequences have been examined for use in fungal systematics, including primers derived from M13 bacteriophage internal repeat [94]. These primers have so far provided differentiation at the intraspecific level, allowing differentiation of closely related isolates within single species [6, 19, 36]. Recently, microsatellite markers in combination with ISSR-PCR (internal short sequence repeats—PCR) have been developed to differentiate and identify Fusarium spp., a sensitive and economical technique that promises better intraspecies specificity [11, 21, 56]. 5. DGGE

Denaturing gradient gel electrophoresis (DGGE) is frequently used for the restriction-enzyme independent detection of DNA sequence variations like single-base substitutions [40]. DGGE exploits the principle that sequence alterations cause changes in the melting temperature of double-stranded DNA, which can be analysed by a linearly increasing gradient of DNA denaturants established in polyacrylamide gels. In practice, nearly all single-base substitutions in amplicons up to 500 bp joined to a GC-clamp could be detected by PCRDGGE-based analysis [102, 124]. The method is used routinely in the detection and enumeration of prokaryotic microorganisms in situ [101]. PCR-denaturing gradient gel electrophoresis (DGGE) method was developed to assess Fusarium species diversity in asparagus plant samples by Yergeau et al. [147]. Fusarium-specific PCR primers targeting a partial region of the translation elongation factor-1 alpha (EF-1 alpha) gene were designed, and their specificity was tested against genomic DNA extracted from a large collection of closely and distantly related organisms isolated from multiple environments. Amplicons of 450 bp were obtained from all Fusarium isolates, while no PCR product was obtained from non-Fusarium organisms. The ability of DGGE to discriminate between Fusarium taxa was tested over 19 different Fusarium species represented by 39 isolates, including most species previously reported from asparagus fields worldwide. The technique was effective to visually discriminate between the majority of Fusarium species and/or isolates tested in pure culture, while a further sequencing step permitted to distinguish between the few species showing similar migration patterns. Total genomic DNA was extracted from field-grown asparagus

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plants naturally infested with different Fusarium species, submitted to PCR amplification, DGGE analysis and sequencing. The two to four bands observed for each plant sample were all affiliated to F. oxysporum, F. proliferatum or F. solani, clearly supporting the reliability, sensitivity and specificity of this approach for the study of Fusarium diversity from asparagus plant samples. 6. Amplified Fragment Length Polymorphism (AFLP)

Another random PCR approach that has recently been developed for DNA fingerprinting and genetic mapping is the amplified fragment length polymorphism (AFLP) technique [138]. This procedure allows high stringency PCR amplification of DNA fragments randomly chosen for restriction fragments. In this technology, genomic DNA is first digested with a restriction endonuclease and oligonucleotide adaptors are ligated to the ends of the restriction fragments. Then, a PCR amplification is performed using primers which include the adaptor sequence, the part of the restriction site sequence remaining on the fragment, and between one and five additional nucleotides which are randomly chosen. This step allows selective amplification of the restriction fragments in which the nucleotides flanking the restriction site match the additional nucleotides of the primers. The amplified fragments are analyzed by denaturing polyacrylamide gel electrophoresis to generate the fingerprint. AFLP also has high resolution and the complexity of the AFLP fingerprint can be planned since that number of resulting fragments depends on the choice of restriction enzyme and the number of additional selective nucleotides in the primers. The amount of DNA used in AFLPs is manyfold less than needed for a single RFLP analysis, plus several more polymorphic markers may be observed per AFLP reaction. Compared to microsatellite markers, AFLP provides an advantage because no sequence knowledge is required prior to analysis. AFLP combines the accuracy of restriction endonuclease digest along with the precision of the PCR. This is a method of obtaining hundreds to thousands of markers for a given individual, each highly reproducible and easily scored against large populations. The AFLP technique is similar to RAPD, in that it analyses the whole genome but it differs from RAPD, i.e. it uses stringent PCR conditions and produces results that are very reproducible.

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AFLP fingerprinting has been developed to evaluate polymorphisms among various organisms, and this has already been applied to the detection of inter- and intraspecific genetic variation in fungi [84, 100]. The method has great potential for revealing variations among many fungi, especially at the intraspeceific level. Recently, AFLP analysis was used to study the genetic relationships within and between natural populations of five Fusarium spp. [1]. Also, Bao et al. [10] used the AFLP method during their studies on genetic analysis of pathogenic and nonpathogenic Fusarium oxysporum from tomato plants. Their results indicated that pathogenic and nonpathogenic strains could be separated into different clusters based on AFLP data, although some nonpathogenic strains grouped with pathogenic ones. Abdel-Satar et al. [1] used AFLP analysis to study the genetic relationships within and between natural populations of five Fusarium spp. They obtained a total of 80 AFLP polymorphic markers using four primer combinations, with an average of 20 polymorphic markers observed per primer pair. Their UPGMA analyses indicated five distinct clusters at the phenon line of 30% on the genetic similarity scale corresponding to the five taxa. The similarity percent of each group oscillated between 87 and 97%. The phenetic dendrogram generated by UPGMA as well as principal coordinate analysis (PCA) grouped all of the Fusarium spp. isolates into five major clusters. Also, no clear trend was observed between clustering in the AFLP dendrogram and geographic origin and host genotype of the tested isolates with a few exceptions. They reported that the results of the present study provide evidence of the high discriminatory power of AFLP analysis, suggesting the possible applicability of this method to the molecular characterization of Fusarium. Belabid et al. [14] used three AFLP selective nucleotide primer pairs to evaluate the amount of genetic variation among isolates of Fusarium oxysporum f. sp. lentis. Ninety-three polymorphic fragments were scored. The isolates could be grouped into two subpopulations based on RAPD and AFLP analysis. Results obtained indicate that there is little genetic variability among a subpopulation of Fusarium oxysporum f. sp. lentis as identified by RAPD and AFLP markers and that there is no apparent correlation with geographical origin or aggressiveness of isolates. Also, the data suggest that Fusarium oxysporum f. sp. lentis isolates are derived from two genetically distinct clonal lineages.

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CONCLUSION

Methods based on polymerase chain reaction (PCR) offer many new tools that are directly applicable to fungal systematics at the species level. PCR-based methods have given a greater insight into molecular variability within fungi and have highlighted the need to consider carefully sampling strategies and sample size, prior to making taxonomic decisions. This insight has also shown that molecular variability is not constant within different fungal species, and that levels of both homoand heterogeneity will vary depending on the species studied. Most filamentous fungi cannot be grouped into traditional biological species, and the species epithet has been applied to different population levels between and within different genera. As a result, the degree of variability observed within one species will not necessarily be comparable with that found in another. These differences in variability within populations, therefore, make it very difficult to select any one molecular technique as the definitive method for defining fungal species. Surprisingly, the introduction of PCR-based techniques has not led to a widespread revision of fungal species names and concepts, and in many cases existing species concepts have been reinforced. However, the wide range of existing molecular heterogeneity found in some species has led to the suggestion that there may be many more cryptic and undescribed species within existing collections. Molecular data will undoubtedly contribute to our understanding of Fusarium diversity, but it will not automatically define species boundaries. We will always have to interpret patterns of diversity in their biological context, incorporating what we know about the ecology, population genetics, incompatibility systems, life cycle, dispersal mechanisms, and host-parasite relationships, etc. References [1] Abdel-Satar MA, Khalil MS, Mohmed IN, Abdel-Salam KA, Verreet JA. Molecular phylogeny of Fusarium species by AFLP fingerprint. Afr J Biotechnol 2003; 2: 51–55. [2] Abramson D, Clear RM, Smith EM. Trichothecene production by Fusarium spp. isolated from Manitoba grain. Can J Plant Pathol 1993; 15: 147–152. [3] Aoki T, O’Donnell K. Morphological and molecular characterization of Fusarium pseudograminearum sp. nov., formerly recognized as the Group 1 population of F. graminearum. Mycologia 1999; 91: 597–609.

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[4] Aoki T, O’Donnell K, Homma Y , Lattanzi AR . Sudden-death syndrome of soybean is caused by two morphologically and phylogenetically distinct species within the Fusarium solani species complex—F. virguliforme in North America and F.tucumaniae in South America. Mycologia 2003; 95: 660–684. [5] Appel DJ, Gordon TR. Intraspecific variation within populations of Fusarium oxysporum based on RFLP analysis of the intergenic spacer region of the rDNA. Exp Mycol 1995; 19: 120–128. [6] Arora DK, Hirsch PR, Kerry BR. PCR-based molecular discrimination of Verticillium chlamydosporium isolates. Mycological Res 1996; 100: 801–809. [7] Assigbetse KB, Fernandez D, Dubois MP, Geiger J-P. Differentiation of Fusarium oxysporum f. sp. vasinfectum races on cotton by random amplified polymorphic DNA (RAPD) analysis. Phytopathology 1994; 84: 622–626. [8] Baayen RP, Gams W. The Elegans fusaria causing wilt disease of carnation. I. Taxonomy. Neth J Plant Pathol 1988; 94: 273–278. [9] Baayen RP, Van Dreven F, Krijger MC, Waalwijk C. Genetic diversity in Fusarium oxysporum f. sp. dianthi and Fusarium redolens f. sp. dianthi. Euro J Plant Pathol 1997; 103: 395–408. [10] Bao JR, Fravel DR, O’Neill NR, Lazarovits G, van Berkum P. Genetic analysis of pathogenic and nonpathogenic Fusarium oxysporum from tomato plants. Can J Bot 2002; 80, 271–279. [11] Barve MP, Haware MP, Sainani MN, Ranjekar PK, Gupta VS. Potential of microsatellites to distinguish four races of Fusarium oxysporum f. sp. circeri prevalent in India. Theoretical and Applied Genetics 2001; 201: 138–147. [12] Baura G, Szaro TM, Bruns TD. Gastrosuillus laricinus is a repeat derivative of Suillus grevellei: molecular evidence. Mycologia 1992; 84: 592–597. [13] Beck JJ, Ligon JM. Polymerase chain reaction assays for the detection of Stagonospora nodorum and Septoriatritici in wheat. Phytopathology 1995; 85: 319–324. [14] Belabid L, Baum M, Fortas Z, Bouznad Z, Eujayl I. Pathogenic and genetic characterization of Algerian isolates of Fusarium oxysporum f. sp. lentis by RAPD and AFLP analysis. African Journal of Biotechnology 2004; 3: 25–31. [15] Bentley S, Pegg KG, Dale JL. Genetic variation among a world-wide collection of isolates of Fusarium oxysporum f.sp. cubense analysed by RAPD-PCR fingerprinting. Mycolo Res 1995; 99: 1378–1384. [16] Blanz PA, Unseld M. Ribosomal RNA as a taxonomic tool in mycology. In: De Hoog GS, Smith MT, Weijman ACM, eds. The Expanding Realm of Yeast Like Fungi. Elsevier, Amsterdam: CBS, 1987; 247–248. [17] Booth C. The genus Fusarium. Kew; Commonwealth Mycological Institute, 1971; pp. 237. [18] Brayford D. Fusarium-molecules maketh the mould. Sydowia 1996; 48, 163–183. [19] Bridge PD, Pearce DA, Rivera A, Rutherford MA. VNTR derived oligonucleotides as PCR primers for population studies in filamentous fungi. Letters in Applied Microbiology 1997; 24: 426–430. [20] Bridge PD, Prior C, Sagbohan J, Lomer CJ, Carey M, Buddie A. Molecular characterization of isolates of Metarhizium from locusts and grass - hoppers. Biodiversity and Conservation 1997; 6: 177–189.

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PLANT GENOME: BIODIVERSITY AND EVOLUTION Volume 2, Part B Lower Groups

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Use of Molecular Markers to Study HostPathogen Co-evolution: A Case Study of the Fungal Pathogen Phialophora gregata WEIDONG CHEN1 and CRAIG R. GRAU2 1

USDA-ARS, Grain Legume Genetics and Physiology Research, 303 Johnson Hall, Washington State University, Pullman, WA 99164, USA 2 Department of Plant Pathology, University of Wisconsin, 1607 Linden Drive, Madison, WI 53706, USA

ABSTRACT The brown stem rot pathosystem was taken as a case study using molecular markers targeted at various taxonomical levels to investigate host-pathogen co-evolution. A series of molecular markers have been developed for differentiation at three taxonomical levels to study the interaction of Phialophora gregata with soybean. First, molecular markers based on ITS region of nuclear rDNA were developed to differentiate the pathogen P. gregata from the co-colonizing fungus Plectosporium tabacinum. Results showed that there was a negative correlation in natural occurrence between the two fungal species in soybean plants. Second, a molecular marker based on an intron near the 5’ end of nuclear small subunit rDNA was used to differentiate two formae speciales of P. gregata, f. sp. adzukicola infecting adzukibean and f. sp. sojae infecting soybean. This marker could be employed to differentiate the two formae speciales without using pathogenicity tests. Third, three sets of molecular markers were developed to separate isolates of P. gregata f. sp.

Address for correspondence: Weidong Chen, USDA-ARS, Grain Legume Genetics and Physiology Research, 303 Johnson Hall, Washington State University, Pullman, WA 99164, USA. Tel: 509-335-9178, Fax: 509-335-7692, E-mail: [email protected].

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sojae into two distinct populations (A and B) and a third population for f. sp. adzukicola. The three independent sets of molecular markers (IGS region of the nuclear rDNA, three microsatellite loci, eight anonymous DNA loci) were all correlated in identifying the two populations. Furthermore, the two genetic populations correspond to defoliating and non-defoliating pathotypes, and differential cultivar preference. Isolates of population A preferentially infect susceptible soybean cultivars and isolates of population B preferentially infect resistant soybean cultivars. This series of molecular markers greatly helped advancing our understanding of adaptation of P. gregata to host plants. Key Words: Brown stem rot, co-evolution, Phialophora gregata, soybean

Abbreviations: IGS: intergenic spacer; ISSR: inter simple sequence repeat; ITS: Internal transcribed spacer; LP-RAPD: long primer random amplified polymorphic DNA. rDNA: ribosomal DNA INTRODUCTION

Plant pathogens and their hosts form intriguing complex relationships. During the evolutionary process, the host plants may have diverged morphologically so much to become different genera, while their pathogen also has evolved parasitically along with the hosts through host specialization but with no detectable morphological changes. A case in point is that soybean [Glycine max (L.) Merr.] and adzuki bean [Phaseolus angularis (Willd.) Ohwi & H. Ohashi] have diverged to different genera, while their common pathogen Phialophora gregata (Allington & D. W. Chamberlain) W. Gams is morphologically the same. In cultivated agroecosystems where host genotypes are human directed, the pathogen also adapts to the human-selected host genotypes with no observable morphological changes. The adaptation could be in the form of cultivar specialization, symptom differentiation and preferential infection. A detailed knowledge of the pathogen population structure and pathogen adaptation will help in selecting cultivars for planting and in breeding for resistance. Sometimes saprophytes or non-virulent parasites, morphologically similar to the pathogen, may also be ecologically associated with the host plant. Molecular separation can expedite differentiation of the pathogen from other fungi that may also cocolonize soybean stems. In all such cases, molecular markers appropriate at different taxonomical levels of the pathogen are very useful and may be necessary for the investigation into the co-evolution of the pathosystem. The fungal pathogen Phialophora gregata provides an ideal example for such investigations. This chapter describes a series of molecular markers that have been developed over the years for P. gregata

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at three taxonomic levels and their applications in studying the hostpathogen co-evolution. THE BROWN STEM ROT PATHOSYSTEM

Brown stem rot is caused by the fungal pathogen Phialophora gregata, first discovered in diseased soybean plants in Illinois, USA, in 1944 [4], and subsequently reported from many other countries including Argentina, Brazil, Canada, Egypt, Japan, Mexico and Yugoslavia [23]. The pathogen also infects adzuki bean and mung bean. It is haploid and reproduces by means of phalidic conidia (asexual or mitotic spores). Its sexual state has never been found and is suspected to be non-existent. It grows slowly on culture media compared to other fungi with colonies expanding about 1 cm diameter in 1 week on potato dextrose agar. P. gregata is thought to be a poor competitor in nature because it has never been isolated directly from soil, but isolated only from diseased plants or plant debris previously colonized parasitically [1]. It resides in organic matter in soil and infects plants through roots and moves up in the plant through the vascular system. It causes symptoms on stems and leaves. Stem symptoms include brown discoloration of the vascular bundle and the pith. Foliar symptoms include wilt, interveinal necrosis and chlorosis. The stem and foliar symptoms may not always occur simultaneously, as they are subject to the pathogen isolate, host genotype and environmental conditions [21, 30, 37, 43, 58]. Tillage practice, rotation sequence, soybean cultivar and plant age all affect disease development and soybean yield [3, 22]. Yield loss in soybean can be up to 40%, but losses of 10 to 20% are more common [2, 42, 43, 51]. Phialophora gregata shows host specialization, and is separated into two formae speciales, f. sp. adzukicola for adzuki bean and f. sp. sojae for soybean [25, 32]. P. gregata f. sp. adzukicola infects adzuki bean and mung bean, but not soybean, whereas P. gregata f. sp. sojae infects soybean and mung bean but not adzuki bean. Further population subdivision has been detected within each of the formae speciales with variations in infectivity or in symptoms they cause on cultivars of their respective host species. Two races of P. gregata f. sp. adzukicola are reported in Japan and can be distinguished by inoculating the differential cultivar Kita-no-otome of adzuki bean. Race 1 is avirulent to, and race 2 is virulent to the differential cultivar Kita-no-otome [34]. There is no definitive cultivar specialization among isolates of P. gregata f. sp. sojae, but isolates do differ

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in severity and the types of symptoms they cause. Gray [21] reported two pathotypes of P. gregata f. sp. sojae. Pathotype I isolates cause stem and foliar symptoms and defoliation, whereas pathotype II isolates cause stem symptoms, and foliar symptoms on lower leaves, but no defoliation [30]. Isolates within each pathotype express a range of aggressiveness on susceptible host genotypes. All soybean genotypes are infected by P. gregata f. sp. sojae. However, soybean genotypes do differ in type and severity of stem and foliar symptoms [51]. A lack of foliar symptoms is considered as resistant. Significant research efforts have been devoted to understanding soybean resistance to brown stem rot and developing resistant soybean cultivars [7, 17, 26, 36, 46, 57]. There are several taxonomical levels that are needed for molecular markers for differentiation: between the pathotypes, between the formae speciales, and also between P. gregata and other fungi inhabiting soybean stems, particularly Plectosporium tabacinum which, although not pathogenic to soybean, is morphologically similar to and frequently causes confusions in isolation and identification of P. gregata [27]. MOLECULAR MARKERS FOR DIFFERENT TAXONOMICAL LEVELS Molecular Markers for Separating the Pathogen from other co-colonizing Fungi

Soybean stem is a habitat of many fungi including pathogens and saprophytes [29]. Often isolation and identification of specific pathogen can be difficult and time consuming using traditional mycological techniques, such as obtaining pure cultures and microscopic examinations. For instance, Plectosporium tabacinum (van Beyma) M. E. Palm, W. Gams et Nireberg, is non-pathogenic to soybean but forms conidia morphologically very similar to Phialophora gregata, frequently inhabits soybean stems, and often causes confusion in isolation and identification of the BSR pathogen P. gregata [27]. Despite similar morphology, P. gregata and Plectosporium tabacinum can be readily separated based on isozyme polymorphisms [43]. A pair of PCR primers based on the nuclear ribosomal DNA was developed specifically for P. gregata [12]. This pair of PCR primers did not amplify DNA from P. tabacinum, Acremonium spp., Cercospora sp. Pythium ultimum, Septoria sp. and Verticillium dahliae [12]. The marker was used not only to identify

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unknown pure cultures to P. gregata, but also to detect P. gregata in infected soybean plants without obtaining pure cultures, which helped speed up the process of pathogen detection and disease diagnosis [12]. Furthermore, a specific marker was developed for identifying and detecting P. tabacinum in soybean plants [14]. Combination of the marker for P. tabacinum and the marker for P. gregata f. sp. sojae enabled detection of both fungal species in soybean plants to investigate the interaction between the two fungi in nature. P. gregata f. sp. sojae infected 51% of 130 arbitrarily sampled soybean plants from four locations, whereas P. tabacinum infected 33% of the plants. However, they occurred simultaneously only in 5% of the plants [14]. They seemed to have a tendency to exclude one another. The study led to a research hypothesis that pre-establishment of P. tabacinum in soybean stem could help reduce brown stem rot incidence. Molecular Markers for Separating Formae Speciales of P. gregata

Separation of the two formae speciales of P. gregata is based on host specialization. Strains that infect adzuki bean but not soybean are grouped in P. gregata f. sp. adzukicola and strains that infect soybean but not adzuki bean are grouped in P. gregata f. sp. sojae. Therefore, pathogenicity assays with adzuki bean and soybean plants are used to identify the two form species, but pathogenicity assays require special environmental condition in plant growth facility, which are time consuming, and laborious. Molecular markers separating and identifying the two formae speciales will simplify the differentiation process, and will also further demonstrate the separate evolution of the two form species. A number of biochemical and molecular markers have been found to differentiate the two form species. Yamamoto et al. [59] found that these two form species differ in four of 15 isozymes. The two form species also differ in total genome contents and GC contents [60], in RFLP patterns of mitochondrial DNA [24], and in sensitivity to antibiotic nystatin [60]. All these differences demonstrate the genetic dissimilarities between the two formae speciales and could be possibly used to separate them. However, with the advent of the PCR technology, molecular markers based on PCR would be more convenient to separate the two formae speciales. While conducting the study on the internal transcribed spacer

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(ITS) of nuclear rDNA of P. gregata, we discovered an intron difference between the two formae speciales. When the ITS region was amplified with the conserved PCR primers ITS1 and ITS4, we obtained a PCR product of about 620 bps from all isolates of both formae specialis of P. gregata. However, when PCR primers ITS5 and ITS4 were used in PCR, which is expected to amplify a product of about 646 bps, we obtained two PCR products of considerably different sizes. One product of 646 bps was found in all isolates of P. gregata f. sp. sojae, and another of 948 bps in all isolates of P. gregata f. sp. adzukicola examined (Fig. 1). The difference was due to the presence of a 304-bp insertion sequence located between primers ITS1 and ITS5 in the isolates of P. gregata f. sp. adzukicola, but not in isolates of P. gregata f. sp. sojae [13]. This insertion sequence has all characteristics of group I introns, including four conserved sequence elements, a U at the 5’ splice site of the exon, a G at the 3’ splice site of the intron, a putative internal guiding sequence. The insertion sequence also fits a secondary structure model for group I introns. Similar to most group I introns found in nuclear small subunit rDNA, this insertion sequence is located in a highly conserved region and is devoid of long open reading frames [13]. This intron provides a convenient marker for separating P. gregata f. sp. adzukicola from P. gregata f. sp. sojae. Molecular markers for separating pathotypes of P. gregata f. sp. sojae

Pathogenic variation among isolates of P. gregata has been frequently observed [21, 30, 37, 41]. Gray reported that isolates of P. gregata could be classified into two pathotypes: defoliating and non-defoliating pathotypes. Recently three classes of molecular markers were developed to study populations of P. gregata f. sp. sojae: rDNA marker [11], three microsatellite loci [16] and eight anonymous DNA markers [40]. Nuclear rDNA marker

P. gregata f. sp. sojae exhibits very limited genetic variation. Studies on isozymes, mitochondrial DNA and nuclear small subunit rDNA and region of rDNA found no variation among isolates of P. gregata f. sp. sojae [14, 24, 41]. Ribosomal DNA is shown to evolve at different rates in coding and non-coding regions. For example, the regions coding for small subunit and large subunit rRNAs are highly conserved and can be useful for comparison among distantly related taxa [9, 50]. The ITS region is

622 646

620 948

Product size (bp) in P. g. f. sp. adzukicola sojae

ITS4

28S

28SF BSRIGS1

Hind III

P. g. f. sp. sojae

18S

CNS1R

1020 bp Populatio A 830 bp Populatio B

BSRIGS2

7 Kbs Intergenic spacer

Fig. 1. Schematic diagram showing the structure of nuclear rDNA of Phialophora gregata and its utilization to separate two formae speciales based on an intron near the 5’ end of small subunit (18S) rDNA, and to separate two populations (A and B) of P. g. f. sp. sojae based on size variation in the intergenic spacer region between transcriptional units (not drawn to scale). The locations of primers 28SF, CNS1R, BSRIGS1, BSRIGS2, ITS1, ITS4 and ITS5 are indicated. For details, see Chen et al. [11,13].

ITS5 ITS1

18S

5.8S

Transcriptional unit

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conserved at species level and is very useful for comparison among closely related species [12, 15]. The intergenic space (IGS) region located between transcriptional units is highly variable and shows considerable variation among populations of the same species [6, 31]. In order to study genetic variation among isolates of P. gregata f. sp. sojae, we initially focused on the IGS region and identified a unique marker for populations of P. gregata f. sp. sojae. Conserved PCR primers 28SF (5’CTG AAC GCC TCT AAG TCA GAA3’) and CNS1R (5’GAG ACA AGC ATA TGA CTA C3’) located on the coding regions of 28S rDNA and 18S rDNA, respectively, were used to amplify the IGS region between transcriptional units (Fig. 1). Normally the IGS region is about 2 to 4 kb long for most fungi. However, the IGS region in P. gregata is unusual, i.e. it is over 7 kb long, longer than any reported IGS region of fungi. Normal Taq DNA polymerase was not efficient enough to amplify this region. We had to employ the high fidelity Platinum Taq DNA polymerase from GIBCO BRL Life Technologies to successfully amplify this 7 kb IGS region [11]. The amplified products were digested with the restriction enzyme Hind III and the digested products revealed size differences among isolates of P. gregata f. sp. sojae (Fig.1). The digested fragments were purified and cloned into pGEM 3ZF(+) vector, and were sequenced. The variable region responsible for the size difference was identified and specific PCR primers BSRIGS1 (5’GGG GTT CCG GGA TTC ACA GG3’) and BSRIGS2 (5’GAG TGG TAA ATG GGG TAA TCA AC3’) were designed to flank the variable region [11]. The specificity of the two PCR primers (BSRIGS1 and BSRIGS2) was tested on a number of morphologically similar and ecologically associated fungi, and the primers were found to be specific for P. gregata. The PCR product amplified using these two primers has become a very useful molecular marker (referred to as rDNA marker hereafter) in studying P. gregata f. sp. sojae. Application of this rDNA marker to 118 isolates of P. gregata f. sp. sojae separated the isolates into two distinct populations: Population A identified by a 1020-bp PCR product, and population B by an 830-bp product of the rDNA marker. A number of investigations ensued that have documented that the two populations are correlated with a number of other independent molecular markers and pathogenic traits.

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Microsatellite markers

Microsatellites or simple sequence repeats are segments of DNA with tandem repeats of short nucleotide motifs. They are normally abundant and scattered throughout eukaryotic genomes [48, 49]. Microsatellites can be highly polymorphic due to their variable number of repeats, and are reproducible and easy to score with appropriate techniques. In order to study evolution of P. gregata, we developed and employed an elaborate procedure for enriching and isolating microsatellites from P. gregata [16]. A primary short-insert (300-600 bp) DNA library was constructed using genomic DNA of P. gregata. Microsatellite enrichment was further carried out using microsatellite oligonucleotides (CAC)5, (GTG)5 and (GACA)4 as PCR primers to synthesize second strand of plasmids from single stranded plasmid DNA prepared from the primary library. Positive clones from the microsatellite-enriched library were further screened with PCR using a primer based on the promoter sequence and another primer of the microsatellite oligonucleotide used in the enrichment. Selected clones were then sequenced to determine the nature of the microsatellite repeats and to design locus-specific primers flanking the microsatellite. The detailed procedure was described by Chen et al. [16]. Twenty-four microsatellite loci were selected for further study by designing locus-specific primers. Initially these were screened for their applicability to, and variability among 16 isolates of P. gregata (8 isolates each of the two populations identified by the rDNA marker). Four of the 24 microsatellite loci were polymorphic among the 16 isolates, whereas the remaining 20 loci were monomorphic. Among the four polymorphic loci, two (GTG-B11B and GTG-B11C) were located on the same clone. Due to their linkage, only one (GTG-B11B) was used for the rest of the P. gregata isolates. The three polymorphic microsatellite loci (CACA20A, GTG-B11B and GTG-H1B) were applied to a collection of more than 100 isolates of P. gregata from soybean and adzuki bean. Only two alleles were found in each of the polymorphic loci among the 118 isolates from soybean, and a third allele in the adzuki bean isolates [16]. The variation in alleles of the microsatellite loci correlated perfectly with the populations identified with the rDNA marker. Population A isolates shared the same alleles of all the microsatellite loci, and population B isolates all showed the other identical allele of the loci. Results also showed that P. gregata has very low levels of genetic variation which was manifested in two ways. First, the number of polymorphic loci was low.

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Only four of the 24 microsatellite loci were polymorphic and two of them were closely linked (located on the same piece of cloned DNA). Second, the number of alleles at the polymorphic loci was low. Only two alleles were observed for each of the three polymorphic loci among 118 isolates representing two distinct populations from a wide geographic area. A third allele was found only in the isolates of P. gregata f. sp. adzukicola [16]. Such a low level of genetic variation is in stark contrast with those reported in other fungi. In Venturia inaequalis, seven polymorphic loci were found in 23 sequenced clones and the number of alleles in each locus ranged from 2 to 48 [55]. In Aspergillus fumigatus, 10 to 23 alleles were detected in four microsatellite loci among 102 isolates [8]. In Ascochyta rabiei, 20 of 37 microsatellite loci were polymorphic with number of alleles ranging from 2 to 14 in only 22 isolates [18]. Anonymous DNA markers

The association between the rDNA marker and the alleles of the three microsatellite loci provided evidence that the rDNA marker identifies genetically distinct populations. However, there is a potential ascertainment bias that can arise either due to a small panel of isolates used in initial screening, or due to the small number of loci used on the test population [10]. In addition, the two populations are sympatric and in close proximity [40], genetic exchanges between the two populations may occur through anastomoses [35]. Since the rDNA marker is playing increasingly important roles in studying the ecology of P. gregata [37], it is desirable to have additional molecular markers to further ascertain that the rDNA marker identifies distinct populations. In this regard, Meng et al. [40] took two approaches to generate anonymous markers: inter simple sequence repeat (ISSR) amplification [20] and long primer random amplified polymorphic DNA (LP-RAPD) [19]. These two approaches were chosen because they tend to randomly survey variation within the whole genome and they are likely to generate DNA markers independent from previous rDNA and microsatellite markers [11, 16]. Generally very low levels of genetic variation were observed among isolates of P. gregata f. sp. sojae. Most of the amplicons were monomorphic among 16 isolates tested in the initial screening. Three primers that produced polymorphic amplicons were subsequently applied onto 189 isolates. PCR amplifications with the three ISSR primers generated a

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total of 14 reproducible amplicons, of which five were polymorphic. Amplification with primer ISSR817 (CA)8A produced six amplicons, one of which was polymorphic and was present in all population A isolates, but absent in all population B isolates. Primers ISSR819 and ISSR824 each generated two polymorphic amplicons with one specific for population A isolates and the other specific for population B isolates. Twenty primers based on sequences of P. gregata developed in a previous study [16] were used in LP-RAPD analyses. Initial screening with the 20 long primers showed that only two of the primers generated polymorphic amplicons among the 16 isolates used in screening. Therefore, only these two primers were used on the rest of the 189 isolates. The two primers generated a total of 17 amplicons, of which three were polymorphic. LP-RAPD with primer A16AF generated six amplicons, and only one (A16AFm1) of which was polymorphic and specific for all population B isolates. The other primer (B4AF) generated two polymorphic amplicons, with one specific for population A and the other specific for population B [40]. Correspondence of molecular markers with pathotypes

It has become clear that there are two distinct populations in P. gregata f. sp. sojae identified by the rDNA marker, alleles of three microsatellite loci, and eight anonymous markers [11, 16, 40]. It is tempting to see if the two populations correspond with the two pathotypes of P. gregata f. sp. sojae reported by Gray in 1971 [21]. Gray [21] observed pathogenic variation in symptoms caused among soybean isolates of P. gregata. One group of isolates called defoliating pathotype caused stem discoloration, and necrosis and chlorosis of leaves, and another group of isolates, nondefoliating pathotype, caused only stem discoloration. Such pathogenic variations were also observed in subsequent studies [41, 58]. Two studies were conducted to examine the relationship between the two populations defined by the molecular markers and the two pathotypes. Hughes et al. [30] employed 54 isolates arbitrarily selected to represent the two populations (27 isolates from each population) and tested their pathogenicity on five soybean cultivars with different levels of resistance. The defining difference between the two pathotypes in causing symptoms is in defoliation. Isolates of the defoliating pathotype can cause stem discoloration, foliar necrosis and chlorosis, and

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defoliation, whereas isolates of non-defoliating pathotype can also cause stem discoloration, and necrosis and chlorosis of lower leaves, but no defoliation. Second, all isolates of population A belonged to the defoliating pathotype, whereas all isolates in the population B belonged to the non-defoliating pathotype. A second study [40] using limited number of isolates on a different set of soybean cultivars also confirmed the findings by Hughes et al. [30]. Correlation of molecular markers with host cultivar preference

Host cultivar specialization is the criterion used in designating races for P. gregata f. sp. adzukicola [34]. In P. gregata f. sp. sojae, there is no definitive cultivar specialization; isolates of both pathotypes infect all soybean cultivars tested in controlled inoculation studies. However, isolation data from field samples showed that the two pathotyes may differ in cultivar preference. Host preference has been documented in a number of pathosystems. Macrophomina phaseolina is a pathogen of both maize and soybean. Chlorate-resistant isolates preferentially colonize maize, whereas chlorate-sensitive isolates preferentially colonize soybean [47]. The cereal pathogen Gaeumannomyces graminis var. avenae infects wheat, rye and other grasses, whereas G. graminis var. tritici preferentially infects wheat, rye and barley. Within G. graminis var tritici, N isolates preferentially infect wheat and R isolates infect wheat and rye. Isolates of G. graminis with the same host preference share the same or similar DNA hybridization patterns, and are differentiated from other isolates with different host preference [45]. Another cereal pathogen, Microdochium nivale, also shows host preference. Both M. nivale var. majus and var. nivale, when used individually in inoculations, are strongly pathogenic on wheat and rye, with var. nivale causing greater disease in rye. However, when the inoculum from the two varieties were mixed in different proportions, M. nivale var. majus showed preference for wheat, whereas M. nivale var. nivale showed a strong preference for rye [52]. M. nivale var. majus is significantly more sensitive than var. nivale to benzoxazolinone, a hydroxamic acid compound derived from rye leaves, indicating a possible mechanism of host preference [52]. These documented cases all show host preference towards different host species. Host preference towards different cultivars of the same species has not been reported. The host preference exhibited by the populations

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of P. gregata f. sp. sojae is towards different cultivars of the same host species. The development of molecular markers has facilitated investigations into the interactions of P. gregata f. sp. sojae with soybean cultivars, particularly into the cultivar preference. Three lines of evidence have been obtained to show that the two populations of P. gregata f. sp. sojae, based on the molecular markers, have differences in cultivar preference. First, after a collection of over 200 field isolates were separated into the two populations using the molecular markers, it was found that most isolates recovered from susceptible soybean cultivars belonged to population A, and most of the isolates from resistant cultivars were classified as belonging to population B [11]. Second, seven soybean cultivars with different levels of resistance to brown stem rot were planted in replicated plots at each of the ten locations in states of Illinois, Minnesota, and Wisconsin for two years. Samples of plant stems were arbitrarily collected near the end of growing seasons from each plot and were assayed using the rDNA marker through polymerase chain reaction to detect pathogen populations in the plants. Population A was predominantly detected in susceptible soybean cultivars Williams 82 and LN92-12054, with 70 and 78% of infected stems, respectively, positive for population A [37]. Population A was predominant in another susceptible cultivar Sturdy in most of the testing locations. Population B was predominant in partially resistant soybean cultivars Dwight, LN9212033 and Bell, with 56, 85 and 99% of the infected stems, respectively, testing positive for population B [37]. Finally, Meng et al. [40] demonstrated the differential cultivar preference using a competitive bioassay under controlled conditions. A mixture of inoculum, consisting of both populations A and B at a 1:1 ratio, was used to inoculate 24 plants each of the resistant cultivar Bell and the susceptible cultivar Sturdy. Five weeks after inoculation, the pathogen populations in the infected plants were detected using the rDNA marker though PCR. Seventeen of the 24 Bell plants were infected by population B, whereas 18 of the 24 Sturdy plants were infected by population A [40]. Chisquare analysis showed that the cultivar preference is significant (P<0.001). All data show that cultivar preference exists in P. gregata f. sp. sojae and the cultivar preference correlates with the separation of populations A and B delineated by the molecular markers.

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The pathogenic features of P. gregata show different levels of host specialization. The two formae speciales show host species specialization, f. sp. adzukicola on adzuki bean and f. sp. sojae on soybean. Within the formae speciales, parallel evolution is evident. The two races of f. sp. adzukicola show host cultivar specialization, whereas the two pathotypes of f. sp. sojae show host cultivar preference and symptom differentiation (Fig. 2). Molecular markers targeting the different taxonomic levels of P. gregata have helped advancing our understanding of the co-evolution of P. gregata with its host plant and will continue to play an important role in studying this pathosystem. Association of Independent Genetic Traits and Clonal Reproduction

Fungal populations may be recombining (outcrossing), clonal (asexual and/or selfing) or both [5, 53]. Clonal populations have distinctive Phialophora gregara

Pathogenic feature

Host species specialization

Molecular markers ITS sequence Intron in 18S rDNA mtDNA P. g. f. sp. adzukicola Adzuki bean

P. g. f. sp. sojae Soy bean

Pathogenic features

Host cultivar specialization or preference Symptom differentiation

Molecular markers

IGS sequence Microsatellite alleles ISSR & LP-RAPD markers

Race 1 Avirulent on Kita-no-otome

Race 2 Virulent on Kita-no-otome

Pathotype I Defoliating Prefers Sturdy

Pathotype II Non-defoliating Prefers Bell

Fig. 2. Schematic diagram depicting the co-evolution of P. gregata with its host plants and parallel evolution in respective formae speciales, and the accompanying pathogenic features and molecular markers. For details, see Kobayashi et al. [32], Gray et al. [21, 24, 25], Chen et al. [11, 12, 13, 16], Hughes et al. [30], and Meng et al. [40].

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features for microorganisms in general [38, 56] and for fungi in particular [5, 33, 44, 54]. The hallmark of sexual reproduction is genetic recombination. In strictly clonal species, there is a lack of genetic recombination, and genetic traits governed by DNA located on different chromosomes will remain associated. P. gregata f.sp. sojae has all the characteristics consistent with clonal genetic structure: identical genotypes are widespread and stable; there is complete absence of recombinant genotypes; alleles of independent marker loci are associated and also three sets of molecular markers consisting of 12 loci are all correlated (Table 1). Furthermore, the two genetic populations of P. gregata f. sp. sojae are correlated with pathogenic (and likely polygenic) traits such as defoliating and nondefoliating pathotypes [30,40] and cultivar preferential infection [37,40]. The two populations represent two clonal lineages. New genotypes may arise within each lineage through mutation [28]. It is conceivable that additional genotypes can be detected within the clonal lineages with molecular techniques, with increasing levels of sensitivity to DNA variation like single nucleotide polymorphism. The clonal lineages are evolutionary units adapting to new host genotypes. Molecular markers developed for the clonal lineages play an important role in studying host-pathogen co-evolution of the brown stem rot pathosystem. Table 1. Correlation among three independent sets of molecular markers, pathotypes and host cultivar preference of Phialophora gregata f. sp. sojae Population

rDNA markera

A B

1020 bp 880 bp

a

Microsatellite Anonymous markersb markersc 111 222

11001001 00110110

Pathotyped

Cultivar preferencee

Defoliating Non-defoliating

Sturdy Bell

Based on the PCR product size defined by two specific primers BSRIGS1 and BSRIGS2 located in a variable region in the non-transcribed intergenic spacer of nuclear ribosomal DNA [11]. b Microsatellite alleles of three loci CAC-A20A, GTG-H1B and GTG-B11B defined by the specific PCR primers [16]. c Based on alleles of eight anonymous DNA markers [40]. d Based on pathogenicity tests with 54 isolates (27 of each rDNA genotype) on five soybean cultivars [30], and eight isolates on two cultivars [40]. e Based on field isolation data [11], replicated field plots in ten locations in three states [37], and under controlled conditions measured by cultivar-preferential infection [40].

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Acknowledgments

The authors wish to thank Teresa Hughes, Xiangqi Meng, Dean Malvick and James Kurle for contributing to this study. This research was supported by various funding agencies including USDA, CSREES, North Central IPM Program, The Illinois Soybean Program Operating Board, The Illinois Council on Food and Agricultural Research (C-FAR), The North Central Soybean Research Program and The Wisconsin Soybean Marketing Board. References [1] Adee EA, Grau CR. Population dynamics of Phialophora gregata in soybean residue. Plant Dis 1997; 81: 199–203. [2] Adee EA, Grau CR, Oplinger ES. Inoculum density of Phialophora gregata related to severity of brown stem rot and yield of soybean in microplot studies. Plant Dis 1995; 79: 68–73. [3] Adee EA, Oplinger ES, Grau CR. Tillage, rotation sequence, and cultivar influences on brown stem rot and soybean yield. J Prod Agri 1994; 7: 341–347. [4] Allington WB, Chamberlain DW. Brown stem rot of soybean. Phytopathology 1948; 38: 793–802. [5] Anderson JB, Kohn LM. Clonality in soilborne, plant-pathogenic fungi. Annu Rev Phytopath 1995; 33: 369–391. [6] Appel DJ, Gordon TR. Intraspecific variation within populations of Fusarium oxysporum based on RFLP analysis of the intergenic spacer region of the rDNA. Exp Mycol 1995; 19: 120–128. [7] Bachman MS, Nickell CD, Stephens PA, Nickell AD, Gray LE. The effect of Rbs2 on yield of soybean. Crop Sci 1997; 37, 1148–1151. [8] Bart-Delabesse E, Humbert JF, Delabesse E, Bretagne S. Microsatellite markers for typing Aspergillus fumigatus isolates. J Clinic Microbio 1998; 36: 2413–2418. [9] Bowman BH, Taylor JW, Brownlee AG, Lee J, Lu S-D, White TJ. Molecular evolution of the fungi: Relationship of the Basidiomycetes, Ascomycetes, and Chytridiomycetes. Mole Bio Evol 1992; 9: 285–296. [10] Brumfield RT, Beerli P, Nickerson DA, Edwards SV. The utility of single nucleotide polymorphisms in inferences of population history. Trend Ecol Evol 2003; 18:249– 256. [11] Chen W, Grau CR, Adee EA, Meng XQ. A molecular marker identifying subspecific populations of the soybean brown stem rot pathogen, Phialophora gregata. Phytopathology 2000; 90: 875–883. [12] Chen W, Gray LE, Grau CR. Molecular differentiation of fungi associated with brown stem rot and detection of Phialophora gregata in resistant and susceptible soybean cultivars. Phytopathology 1996; 86: 1140–1148. [13] Chen W, Gray LE, Grau CR. Characterization of a group I intron in nuclear rDNA differentiating Phialophora gregata f. sp. adzukicola from P. gregata f. sp. sojae. Mycoscience 1998; 39, 279–283.

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[14] Chen W, Gray LE, Kurle JE, Grau, CR. Specific detection of Phialophora gregata and Plectosporium tabacinum in infected soybean plants using polymerase chain reaction. Mol. Ecol 1999; 8: 871–887. [15] Chen W, Hoy JW, Schneider RW. Species-specific polymorphisms in transcribed ribosomal DNA of five Pythium species. Exp Mycol 1992; 16: 22–34. [16] Chen W, Shi X, Chen YC. Microsatellite markers and clonal genetic structure of the fungal pathogen Phialophora gregata. Mycol Res 2002; 106, 194–202. [17] Eathington SR, Nickell CD, Gray LE. Inheritance of brown stem rot resistance in soybean cultivar BSR 101. J. Heredity 1995; 86: 55–60. [18] Geistlinger J, Weising K, Kaiser W, Winter P, Kahl G. Locus-specific microsatellite markers for the fungal chickpea pathogen Didymella rabiei (anamorph) Ascochyta rabiei. Mol Ecol 2000; 9: 1939–1941. [19] Gillings M, Holley M. Amplification of anonymous DNA fragments using pairs of long primers generates reproducible DNA fingerprints that are sensitive to genetic variation. Electrophoresis 1997; 18, 1512–1518. [20] Godwin ID, Aitken EAB, Smith LW. Application of inter simple sequence repeat (ISSR) markers to plant genetics. Electrophoresis 1997; 18, 1524–1528. [21] Gray LE. Variation in pathogenicity of Cephalosporium gregatum isolates. Phytopathology 1971; 61: 1410–1411. [22] Gray LE. Role of temperature, plant age, and fungus isolate in the development of brown stem rot in soybeans. Phytopathology 1974; 64: 94–96. [23] Gray LE, Grau CR. Brown stem rot. In: Hartman GL, Sinclair JB, Rupe JC, eds. Compendium of Soybean Diseases. St. Paul, MN, USA: APS Press, 1999; 28–29. [24] Gray LE, Hepburn AG. Mitochondrial DNA restriction patterns of Phialophora gregata isolates from soybean and adzuki bean. Phytopathology 1992; 82, 211–215. [25] Gray LE, Pataky JK. Reaction of mung bean plants to infection by isolates of Phialophora gregata. Plant Dis 1994; 78: 782–785. [26] Hanson PM, Nickell, CD, Gray, LE, Sebastian SA. Identification of two dominant genes conditioning brown stem rot resistance in soybean. Crop Sci. 1988; 28: 41–43. [27] Harrington TC, Steimel J, Workneh F, Yang XB. Molecular identification of fungi associated with vascular discoloration of soybean in the North Central United States. Plant Dis. 2000; 84: 83–89. [28] Harrington TC, Steimel J, Workneh F, Yang XB. Characterization and distribution of two races of Phialophora gregata in the North-Central United States. Phytopathology 2003; 93, 901–912. [29] Hartman GL, Sinclair JB, Rupe JC. eds. Compendium of Soybean Diseases. St Paul, MN, USA: APS Press, 1999 (4th ed). [30] Hughes TJ, Chen W, Grau CR. Pathogenic characterization of genotypes A and B of Phialophora gregata f. sp. sojae. Plant Dis 2002; 86, 729–735. [31] Jackson CJ, Barton RC, Evans EG. Species identification and strain differentiation of dermatophyte fungi by analysis of ribosomal-DNA intergenic spacer regions. J Clinic Microbiol 1999; 37: 931–937. [32] Kobayashi K, Yamamoto H, Negishi H, Ogoshi A. Formae speciales differentiation of Phialophora gregata isolates from adzuki bean and soybean in Japan. Ann Phytopathol Soc Japan 1991; 57, 225–231.

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[33] Kohn LM. The clonal dynamic in wild and agricultural plant-pathogen populations. Can J Bot 1995; 73 (Suppl. 1): 1231–1240. [34] Kondo N, Fujita S, Murata K, Ogoshi A. Detection of two races of Phialophora gregata f. sp. adzukicola, the causal agent of adzuki bean brown stem rot. Plant Dis 1998; 82, 928–930. [35] Leslie JF. Fungal vegetative compatibility. Annu Rev Phytopathol 1993; 31: 127–150. [36] Lewers KS, Crane EH, Bronson CR, Schupp JM, Keim P, Shoemaker RC. Detection of linked QTL for soybean brown stem rot resistance in ‘BSR 101’ as expressed in a growth chamber environment. Mol Breeding 1999; 5, 33–42. [37] Malvick DK, Chen W, Kurle JE, Grau CR. Cultivar preference and genotype distribution of brown stem rot pathogen Phialophora gregata in the Midwestern United States. Plant Dis 2003; 87, 1250–1255. [38] Maynard Smith J, Smith NH, O’Rourke M, Spratt BG. How clonal are bacteria? Proc Natl Acad Sci USA 1993; 90: 4384–4388. [39] Meng X, Chen W. Applications of AFLP and ISSR techniques in detecting genetic diversity in the soybean brown stem rot pathogen Phialophora gregata. Mycol Res 2001; 105, 936–940. [40] Meng X, Grau CR, Chen W. Cultivar preference exhibited by two sympatric but genetically distinct populations of the soybean brown stem rot pathogen Phialophora gregata f. sp. sojae. Plant Pathology 2005; 54, 180–188. [41] Mengistu A, Grau CR. Variation in morphological, cultural, and pathological characteristics of Phialophora gregata and Acremonium sp. recovered from soybean in Wisconsin. Plant Dis 1986; 70: 1005–1009. [42] Mengistu A, Grau CR. Seasonal progress of brown stem rot and its impact on soybean productivity. Phytopathology 1987; 77, 1521–1529. [43] Mengistu A, Grau CR, Gritton ET. Comparison of soybean genotypes for resistance to and agronomic performance in the presence of brown stem rot. Plant Dis 1986; 70, 1095–1098. [44] Milgroom MG. Recombination and the multilocus structure of fungal populations. Annu Rev Phytopath 1996; 34: 457–477. [45] O’Dell M, Flavell RB, Hollins TW. The classification of isolates of Gaeumannomyces graminis from wheat, rye and oats using restriction fragment length polymorphisms in families of repeated DNA sequences. Plant Pathol 1992; 41, 554–562. [46] Patzoldt ME, Chen W, Diers BW. Evaluation of soybean plant introductions from China for resistance to brown stem rot. Online. Plant Health Progress doi:10.1094/ PHP–2003–0702–01–RS, 2003. [47] Pearson CAS, Leslie JF, Schwenk FW. Host preference correlated with chlorate resistance in Macrophomina phaeolina. Plant Dis 1987; 71, 828–831. [48] Queller DC, Strassmann JE, Hughes CR. Microsatellites and kinship. Trend Ecol Evol 1993; 8: 285–288. [49] Rassmann K, Schlötterer C, Tautz D. Isolation of simple-sequence loci for use in polymerase chain reaction-based DNA fingerprinting. Electrophoresis 1991; 12: 113– 118. [50] Rehner SA, Samuels GJ. Taxonomy and phylogeny of Gliocladium analysed from nuclear large subunit ribosomal DNA sequences. Mycol Res 1994; 98: 625–634.

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[51] Sills GR, Gritton ET, Grau CR. Differential reaction of soybean genotypes to isolates of Phialophora gregata. Plant Dis 1991; 75, 687–690. [52] Simpson DR, Rezanoor HN, Parry DW, Nicholson P. Evidence for differential host preference in Microdochium nivale var. majus and Microdochium nivale var. nivale. Plant Pathol 2000; 49, 261–268. [53] Taylor JW, Geiser DM, Burt A, Koufopanou V. The Evolutionary biology and population genetics underlying fungal strain typing. Clin Microb Rev 1999; 12: 126– 146. [54] Taylor JW, Jacobson DJ, Fisher MC. The evolution of asexual fungi: Reproduction, speciation and classification. Annu Rev Phytopath 1999; 37: 197–246. [55] Tenzer I, degli Ivanissevich S, Morgante M, Gessler C. Identification of microsatellite markers and their application to population genetics of Venturia inaequalis. Phytopathology 1999; 89: 748–753. [56] Tibayrenc M, Kjellberg F, Arnaud J. Oury B, Brenière F, Dardé M-L, Ayala FJ. Are eukaryotic microorganisms clonal? A population genetics vantage. Proc Natl Acad Sci USA 1991; 88: 5129–5133. [57] Willmot DB, Nickell CD. Genetic analysis of brown stem rot resistance in soybean. Crop Sci 1989; 29: 672–674. [58] Willmot DB, Nickell CD, Gray LE. Physiologic specialization of Phialophora gregata on soybean. Plant Dis 1989; 73, 290–294. [59] Yamamoto H, Kobayashi K, Ogoshi A. Isozyme polymorphism in Phialophora gregata isolates from Adzuki bean and soybean in Japan. Ann Phytopath Soc Japan 1990; 56, 584–590. [60] Yamamoto H, Kobayashi K, Ogoshi A. Characterization of the nuclear DNA of Phialophora gregata f. sp. adzukicola and sojae. Mycoscience 1995; 36: 117–119.

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Utility of the Internal Transcribed Spacers of the 18S-5.8S-26S Nuclear Ribosomal DNA in Land Plant Systematics with Special Emphasis on Bryophytes ALAIN VANDERPOORTEN 1*, BERNARD GOFFINET 2 and DIETMAR QUANDT 3 1

Research associate of the Belgian Funds for the Scientific Research at University of Liège, Dept. of Life Sciences, Belgium 2 University of Connecticut, Department of Ecology and Evolutionary Biology, Storrs, CT, USA 3 Institute of Botany, University of Dresden, Zellescher Weg 22, D-01062 Dresden, Germany

ABSTRACT The internal transcribed spacers (ITS) of ribosomal DNA repeat have been one of the most exploited sources of molecular data for systematic studies in plants. The molecular features of the ITS in land plants, with special emphasis on bryophytes, are summarized in order to provide a comprehensive and practical review of the production and analysis of the molecule for phylogenetic analysis. The ITS have proved to be extremely useful at resolving taxonomic issues at or below the genus level in angiosperms, and remain a promising source of data in the pteridophytes and the bryophytes. However, mounting evidence for the existence of paralogs and pseudogenes and the lack of variability of the molecule in a large array of taxa towards the species level, suggest that other strategies, such as sequence characterized amplified

*

Address for correspondence: Alain Vanderpoorten, University of Liège, Dept. of Life Sciences, B-22 Sart Tilman, B-4000 Liège, Belgium. E-mail: [email protected]

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regions, are crucially needed for resolving taxonomic issues in the context of an increasing concern for evolutionary phenomena at the inter- and intraspecific levels. Key Words: Internal transcribed phylogenetics, review, bryophytes

spacers,

nuclear

ribosomal

DNA,

INTRODUCTION

Cytoplasmic [i.e. mitochondrial (mt) and chloroplast (cp) DNA] and nuclear (n) DNA, the usual triptych of phylogenetic tools, exhibit contrasting strengths and weaknesses. Cytoplasmic DNA has the virtue of comprising single-copy genes. Most chloroplast genes have a limited potential to accumulate mutations and therefore, provide the appropriate window of resolution to study plant phylogeny at deep levels of evolution[25]. Much of the diversity is confined to non-coding regions that accumulate mutations at a higher rate than actual genes. These regions provided phylogenetic information at the intra- and interspecific level in a range of mosses[38], pteridophytes[52] and angiosperms (for example, Palmer et al. [88]), but have also been used to reconstruct the relationships at higher taxonomic levels in mosses[38], hornworts[115] or angiosperms[17]. However, the number of available loci in the chloroplast genome is highly reduced compared to the genomes of the original endosymbiont, as many genes have been transferred to the nuclear compartment[116]. Genome economization has thus led to a severe reduction of loci and of non-coding regions. Most of the chloroplast genome is indeed composed of coding rather than non-coding sequences. For example, the 121,025 nucleotides of the complete chloroplast genome of the liverwort Marchantia polymorpha correspond approximately to 70.5% coding regions[84, 117]. No genes have introns in the plastid genome of the unicellular red alga Cyanidioschyzon merolae[83]. In the cryptophyte alga, Guillardia theta, intergenic spacers are very short, no introns have been detected, and several genes overlap[34]. By contrast, genes probably make up <5% of the total nuclear genome in maize [30]. The potential of the chloroplast genome for phylogenetic inferences may thus be limited, unless the genome is broadly surveyed based on restriction digests [56, 92] or if it is completely sequenced[57]. This has been done for a small but growing number of taxa[116]. Using complete sequences of the chloroplast genome, Goremykin et al.[39] showed for example

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that, in contrast with the interpretation of a basal position of Amborella within the angiosperms based on the phylogenetic analysis of a limited number of genes, the genus is even not the most basal among dicots. The mtDNA proved to be a suitable tool for phylogenetic analysis at low taxonomic levels in animals (Avise[3]; but see Hugall et al.[54]). In plants, the mtDNA genome, that comprises 366,924 nucleotides in the herb, Arabidopsis thaliana[118], is structurally highly complex and variable[61, 89]. The sequences are, however, not particularly divergent within or even among species[87]. Hence, like most of the cpDNA genome, regions of the mtDNA seem best suited for addressing phylogenetic issues at a deep level [14, 93]. Nuclear genes potentially contain a wealth of information about organismal phylogeny but are often present in multiple, paralogous copies[85, 86]. Multiplicity of sequences from the same taxa, combined with uneven taxonomic sampling, often obfuscates the identification of orthologous sequences and greatly complicates the relationships between gene and species trees[33]. The genes encoding the RNAs composing cytoplasmic ribosomes are organized in a cistron, which is present in many copies in the nuclear genome. The number of repeat copies varies greatly across plants, even between congeneric species. Quercus species, for example, possess between 1,300 and 4,000 copies[127]. Although evidence for the existence of paralogs and pseudogenes is mounting[1, 5], nrDNA repeats are, despite their hundreds to thousands of copies in the eukaryote genome, often remarkably homogeneous within the same species[9]. This homogeneity is attributed to concerted evolution of the entire repeat[51], a process based on gene conversion and recombination that maintains great similarity among repeat units within a species (see Elder and Turner[35] for review). In addition, nrDNA includes coding and noncoding regions with contrasting evolutionary rates, providing phylogenetic signal at different taxonomic levels. Hence, nrDNA has been one of the most exploited sources of molecular data for systematic studies in plants (see Hershkovitz et al.[49] for review). This is especially true for the internal transcribed spacers of the 18S5.8S-26S rDNA(ITS). The first phylogenetic inferences from ITS sequence variation in plants were obtained in angiosperms[7] and, more recently, in bryophytes [Spagnuolo et al.[113] and Chiang and Schaal[22] based on ITS 1, and Chiang and Schaal[23] based on ITS2],

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and pteridophytes[52]. As a comparison, of 244 papers on plant phylogenetics published during the last five years in several of the leading systematics and evolution journals, two-thirds involving comparisons at the genus level or below include ITS sequence data, and one-third of all published phylogenetic hypotheses have been based exclusively on ITS sequences [1]. In this review, we briefly present the molecular features of the ITS in land plants with special emphasis on bryophytes, examine the consequences of these features for the production and analysis of ITS sequences, and discuss the phylogenetic utility of the spacers at different taxonomic levels. LOCATION AND GENERAL FEATURES OF THE ITS

The ITS region comprises two internal transcribed spacers separated by the 5.8S gene and flanked by the 18S and 26S genes. This region is part of the rDNA cistron that encodes 18S, 5.8S, and 26S rRNAs (Fig. 1a). The cistron occurs in hundreds to thousands of copies [51]. Two consecutive cistrons are separated by an intergenic spacer(IGS), which comprises an external transcribed spacer (ETS) and a nontranscribed spacer region (NTS)[69]. The repeats are concentrated at one to few chromosomal loci as revealed by fluorescent in situ hybridization [24, 50, 122]. The number of loci may vary with the ploidy level (for example in Sanguisorba [Rosaceae] [77]) or change due to chromosomal reorganizations (for example, in Silene [Caryophyllaceae] [106]). Organization of the repeat slightly differs from this general scheme in mosses and liverworts. Based on Southern blot and fluorescence in situ hybridization analysis, Sone et al. [111] demonstrated that, unlike in vascular plants, the 5S rRNA of the hepatic, Marchantia polymorpha, are encoded within the 18S-26S rDNA repeat unit (Fig. 1b). A 5S rDNA was also found in the rDNA repeat of a moss, Funaria hygrometrica, by a homology search in a database [111]. This suggests that there has been structural re-organization of the rDNAs repeat early in the evolution of land plants. Whether the insertion of the 5S gene in the rDNA cistron is diagnostic or not for mosses and liverworts is, however, yet to be established. Existence of another specific feature of the bryophyte 18S26S rDNA gene, namely the occurrence of a putative intron separating a 26S 5’ exon of about 10bp from the remaining 26S gene, has been postulated[20, 21]. However, no experimental evidence of the exon has

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ITS1 BMBC-R ITS5

5' ETS

18S (1800 bp)

TTS1

5.8S 165 bp

IGS

ITS3

TTS2

ITS2 (a) 26S

ITS4

5S IGS1

IGS

26S 3400 bp

3' ETS

LS4-R 2 kb 18S

5.8S

IGS2

26S

ITS1 ITS2

(b)

Fig. 1. Genetic map of 18S-26S ribosomal DNA genes in angiosperms (a) and bryophytes (b) with approximate lengths of the different regions. Dashed arrows indicate the universal primers used for PCR amplification (see Table 2).

Table 1. Length range of ITS1 and 2 in selected bryophyte, gymnosperm, and angiosperm taxa Taxon (number of species studied) Angiosperms Rosa (15) Saintpaulia (8) Gentiana (20) Antennaria (30) Nothofagus (22) Gymnosperms Bryophytes Amblystegiaceae s.l. (39)

Reference

ITS1

ITS2

125 78 126 13 76 70, 73

253-255 228-249 223-238 253-260 218-228 610-3100

207-211 196-245 216-234 213-219 204-227 182-370

119

280-340

307-371

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been provided and it is now clear that both exon and intron belong to the ITS2 [94]. As a consequence, a number of ITS2 sequence data must be updated in the GenBank database (see updated GenBank Accession #X80212 of the ribosomal RNA gene repeat in Funaria hygrometrica). The GC content of ITS angiosperm is almost always higher than 50%[49]. For example, it ranges between 63 and 72% in the Salicaceae[66] and between 51.7 and 59.0% in the Nothofagaceae[76]. Similar disproportional ratios have been reported in bryophytes [95, 101]. As a result, substantial portions of ITS are believed to fold into helices or more complex structures via intramolecular base pairing[29]. For both spacers, secondary structure models have been proposed. Mai and Coleman[74], and Coleman et al.[28], inferred ITS secondary structures for the green algae Volvocales, using minimum energy optimization in combination with a comparative phylogenetic analysis. Thereupon, a common secondary structure of the ITS2 for green algae and flowering plants was postulated (Fig. 2) [74]. Reported secondary structures within land plants are still limited and often lack examples of compensating base pair changes to further support the calculated structures[9, 71]. However, it seems that ITS secondary structures are somehow evolutionarily constrained. Indeed, common secondary structure motifs might have a functional role [27, 71], bringing the small and large subunits into close proximity within a processing domain during maturation of the rRNAs[68, 91]. Although the ITS appear to play a role in the maturation of nuclear rRNA, they are cleaved or otherwise digested during the assembly of the ribosomal subunits[49]. Selective constraints are thus much lower on the spacers than on the coding regions, leading to higher inter-and even intrataxon variability. Part of this variability consists of changes in the length of the spacers. In gymnosperms for instance, length of ITS1 ranges between 610 to 3,100 [70, 73] bp vs 159-360 bp in pteridophytes[73] and angiosperms[9]. Length of ITS2 seems more conserved and ranges between 180 and 370bp[73] (Table 1). However, evolutionary patterns in the two spacers (overall rates, base composition biases) are usually parallel [9].

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11 =

111 18

1 8

111 11 >

18

1

111 11 ?

1

18

Fig. 2. Secondary structure of ITS2 of Oryza sativa (Poaceae) (a), Photinia pyrifolia (Rosaceae) (b), and Sinapis alba (Brassicaceae) (c). The four major helices are labeledIV. All diagrams illustrate the universally conserved pyrimidine-pyrimidine pairing in helix II and the conserved trinucleotide motif in helix III (white lettering on a black background). In Photinia (b), nucleotides in white lettering on a gray blackground have no changes at that position among the 23 Rosaceae examined; nucleotides in bold, uppercase lettering have one change at that position; nucleotides in uppercase lettering have two exceptions; and nucleotides in lowercase have three or more exceptions at that position (modified from Mai and Coleman [74]).

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ITS AMPLIFICATION, SEQUENCING, AND ANALYSIS Amplification and Sequencing

Universal ITS primers were designed from highly conserved regions of the flanking rRNA genes in fungi[124] and were subsequently successfully used in a vast array of organisms, sometimes under a more or less modified form (Table 2). The universality of these primers definitely helped in making the ITS one of the most widely used regions in phylogenetics. The high degree of sequence conservation of the primer Table 2. Universal primer sequences of White et al.[124] used for amplifying the ITS region in plants (ITS1-ITS5). BMBC-R[64] and LS4-R[101] are used in a nested PCR amplification. 5.8S, 5.8F[114] and 5.8S-R[101], 25R[114], and 18F correspond to slightly modified primers of ITS2, ITS3, ITS4, and ITS5, respectively, redesigned for bryophytes. A, B and C, E correspond to the pair of external and internal primers, respectively, used in the recombinant PCR described in Blattner[15] for degraded DNA. 1830F and 26S 25 were designed by Nickrent et al.[81] and used in Isoëtes[52]. See Fig. 1 for primer postion Forward BMBC-R ITS1 ITS3 ITS5 5.8S-R 5.8F A E 18F 1830F

GTA CAC ACC GCC CGT CG TCC GTA GGT GAA CCT GCG G GCA TCG ATG AAG AAC GCA GC GGA AGT AAA AGT CGT AAC AAG G TCG ATG AAG AAC GCA GCG GCA ACG ATG AAG AAC GCA GC GGA AGG AGA AGT CGT AAC AAG G CGG CAA CGG ATATCT CGGCTC GGA AAG AGA AGT CGT AAC AAG G AAC AAG GTT TCC GTA GGT GA

Reverse LS4-R ITS4 ITS2 25R B C 5.8S 26S 25R

TCA AGC ACT CTT TGA CTC TC TCC TCC GCT TAT TGA TAT GC GCT GCG TTC ATC GAT GC TCC TCC GCT TAG TGA TAT GC CTT TTC CTC CGC TTA TTG ATA TG GCA ATT CAC ACC AAG TAT CGC CGC TGC GTT CTT CAT CG TAT GCT TAA AYT CAG CGG GT

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annealing sites across eukaryotic autotrophs and fungi in particular may, however, result in the inadvertent amplification and sequencing of a foreign DNA source rather than of the intended target organism. As a consequence, extreme caution is required when sequences of a new group of plants are investigated, and comparison of the obtained sequences with those available in GenBank is recommended[109]. Amplifying and sequencing the ITS has generally been achieved using standard PCR and sequencing procedures. The high number of copies of nrDNA allows for sufficient copies to be extracted from minimal amounts of plant tissue. Hence, a single PCR amplification using external primers annealing in the flanking 18S and 26S regions is usually required. Obtaining sequences from herbarium material in which DNA often is severely degraded may, however, require more refined strategies including: (i) the amplification of ITS1 and ITS2 separately using internal primers that anneal in the conserved 5.8S gene. The separately amplified ITS1 and ITS2 fragments can subsequently be combined before sequencing without impairing the quality of the sequences[15]; (ii) the use of a nested PCR with BMBC-R and LS4-R, whose product is used as the template in a subsequent amplification using primers ITS1 and ITS4[101]; (iii) the combination of both internal and external primers in a single PCR[15]. PCR’s are usually accomplished with the same temperature profile according to a more or less modified version of the following scheme: 30 cycles at 95°C for 1 minute, 50°C for 1 minute, and 72°C for 45 seconds plus an additional 5 seconds for each successive cycle; and a final 7 minute extension at 72°C. The secondary structure of the region, including helices or more complex structures, sometimes blocks polymerization steps during amplification and sequencing. In this case, addition of about 5% v/v DMSO has been advocated as a means to enhance ITS amplification[49]. Sequence Alignment

Alignment methodology is particularly relevant to nrDNA analysis due to length variability, making it necessary to insert gaps to preserve positional homology. The existence of conserved motifs may, however,

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facilitate sequence alignment. Examples of such regions include a motif of about 25 bp in ITS1 conserved among flowering plants and gymnosperms[71, 73], but apparently not in bryophytes[20]. Spermatophytes share several motifs in the ITS2, which have been highly conserved for the last 350 millions years following the early radiation[47]. Between deeper phylogenetic branches (for example, among lineages of land plants), common sequence motifs are almost absent, except for two postulated sequence motifs, namely a pyrimidine mismatch in helices II and a GGU triplet on the 5’apex of helix III (Fig. 2), which are conserved at least between algae and flowering plants[74]. At high taxonomic level or between the conserved motifs, phylogenetic analysis of the ITS is often impeded by regions that are ambiguously aligned. Comparison of rRNA secondary structure has been advocated as a means of refining alignment by improving gap positioning[26] and has been applied in different groups[2, 37, 40]. Although structural information can be used to improve alignments, the placement of some nucleotides will, however, remain arbitrary[60]. Lutzoni et al.[72] presented a new method that allows the inclusion of ambiguously aligned regions without violating homology. This procedure consists of three steps: (1) delimitation of homologous regions on either side of the region of ambiguously aligned sequences; (2) coding each ambiguously aligned region as a new character that replaces its respective ambiguous region; and (3) elaboration and application to each of the coded characters of a specific step matrix that accounts for the differential number of changes (summing substitutions and indels) needed to transform one sequence to another. The optimal number of steps included in the step matrix is the one derived from the pairwise alignment with the greatest similarity and the least number of steps. This approach potentially enhances phylogenetic resolution and support by integrating previously non-accessible characters without violating positional homology[32]. Phylogenetic Analysis

Length variability, and the consequent need to insert gaps to preserve positional homology, is a crucial issue for phylogenetic analysis of highly length variable non-coding DNA regions such as the ITS. Usually, gaps are treated as missing data. A treatment of gap positions as a fifth character, as proposed by Gottschling et al.[40], does not seem to be

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appropriate, except when the indel consists of a single nucleotide, otherwise the indel is indirectly weighted as a function of its length. Unambiguously defined insertions and deletions can be converted into binary characters and a coding scheme can be appended to the data matrix[58, 60]. These characters would be complemented by those sites within the indel that exhibit a point substitution[10]. Following this coding scheme, each indel is thus replaced by one binary character plus a number of variable sites within the indel. The most recent application of the method can be found in Baumel et al. [12]. ITS data matrices frequently violate the assumptions, under unweighted maximum parsimony, that all bases occur at equal frequencies and evolve to other bases at comparable rates. Indeed, nrDNA tends to have >50% GC content in angiosperms, and the same seems to hold true for bryophytes. Furthermore, as a consequence of the nucleotide bias in favor of energy-rich GC bounds, substantial portions of the ITS are believed to exhibit complex and functional structures via intramolecular base pairing. Compensatory mutations or secondary mutations that maintain RNA base-pairing relations, which are observed in structurally conserved regions, are regarded as correlated characters and several authors have stressed that these should be downweighted in phylogenetic analysis[48, 123]. Therefore, differential weighting schemes have been advocated or discussed during the last few years [110]. Finally, as part of the ribosomal multigene family, the ITS region can exhibit infragenomic polymorphism. Within-individual polymorphism may either occur in transition stages of concerted evolution when mutation rates exceed the rate of concerted evolution. This may happen as a result of interspecific hybridization, when pseudogenes evolve, or when location of nrDNA loci on nonhomologous chromosomes disrupts concerted evolution (see Campbel et al. [19] for review). For practical purposes, the problem of ITS polymorphism, which actually seems to be the rule rather than the exception [5, 16, 59], is not whether it exists, but whether its pattern could mislead phylogenetic analysis. At the level of closely related species, it is unlikely that paralogues are differentiated sufficiently to strongly support an incorrect organismal tree node. At greater divergences, homogenization is sufficient to fix intertaxon differences, such that intra-individual variants with a few substitutions or indels will not affect interpretation of organismal phylogeny [18]. If functional paralogues persist through multiple and substantial

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divergences [19], standard amplification and sequencing protocols should reveal the polymorphism in at least some samples and protocols can be adjusted (for example, primer redesign) to search for complete orthologous sets. This approach will necessitate cloning the PCR products and sequencing all alternative clones. PCR and/or cloning steps can, however, strongly favor one of the forms, sometimes an apparent pseudogene, as evidenced by anomalous divergences and extraordinary deletions-substitutions at otherwise highly conserved positions [18]. Patterns of nucleotide substitutions within the highly conserved 5.8S gene can be used to distinguish presumed functional sequences from putative pseudogenes [97]. Furthermore, even when divergent pseudogenes are involved, the ITS polymorphism may not necessarily mislead phylogenetic inference [97]. PHYLOGENETIC UTILITY AND LIMITATIONS

The existence of universal primers, the relative ease of the amplification in many taxa, and the frequent sequence homogeneity among copies within species, have made the ITS one of the most widely used regions in phylogenetics. Length polymorphism may be informative for delineating taxa such as genera of the moss family, Pottiaceae [112]. Useful taxonomic markers were also found by using restriction site polymorphism (PCR-RFLP) (Polymerase chain reaction-restriction fragment-length polymorphism) in angiosperms [55, 65] and in mosses [90, 121]. However, the fairly small size of the region limits the utility of PCR-RFLP, whereas it makes the region a fairly easy target for sequencing. ITS sequences have proved to be of tremendous utility to angiosperm [49], moss [102, 104, 119], and liverwort [41-44, 45-46, 98] phylogeny. Phylogenetic analysis based on ITS sequences have never been conducted on hornworts and it is only very recently that ITS sequences have been used for inferences in the pteridophyte genus, Isoëtes. In the latter, patterns of variation in ITS sequences followed a global geographical gradient. The sequences offered, however, little signal if any to resolve the relationships among North American species, a pattern that could be explained by a rapid radiation [52]. In gymnosperms, length polymorphism hampers phylogenetic utilization of ITS. In the genus Pinus in particular, the diversity in copies of the ITS1 within individuals suggests that concerted evolution is operating slowly. Tandem

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subrepeats of the molecule form stem loops comparable to those in other genera of the Pinaceae and may be promoting recombinations between repeats, resulting in ITS1 chimeras [36]. However, it has been suggested that part of ITS1 and the whole ITS2 may be potentially employed in gymnosperm phylogenetic reconstruction [109]. ITS thus clearly holds a promise for inferring phylogenetic relationships among a still very large number of taxa. In angiosperms in general, ITS is best suited for reconstructing relationships among closely related genera and infrageneric groups [9]. Given their level of polymorphism at low taxonomic levels, ITS are commonly used to compile evidence of reticulate evolution. ITS topologies have been compared to cpDNA phylogenies to discuss the possibilities of reticulate evolution in angiosperms [62, 67, 79, 100, 108] and bryophytes [103]. Sequence additivity [52] and PCR-RFLP patterns [11, 82] of the ITS were similarly used to find evidence for the origin of hybrids and determine their progenitor species. In polyploid species possessing different parental ITS sequence, ITS sequences were used to document reticulate evolution in much the same way that allozymes have been used [99]. However, increasing reports of fast rates of concerted evolution have cast doubt on their reliability for inferring past reticulation events, providing a strong incentive to abandon this methodological approach [96, 107]. At higher level (family and higher taxa), the ITS exhibit conserved sequence patterns across angiosperms. Hershkovitz and Zimmer [48] have shown that one-third to one-half of the ITS2 sequence can be aligned above the family level in angiosperms. Although the results do not challenge the notion that these sequences are not ideal for deep-level phylogenetic analysis, they suggest that ITS2 includes a weak deep-level phylogenetic signal [47] sufficient to diagnose lineages at several hierarchical levels [48]. Furthermore, information regarding the ITS2 secondary structure in highly conserved regions with compensatory base changes may also constitute another set of characters on its own and allows comparisons at deep phylogenetic levels, making the ITS a ‘double-edged tool’ for eukaryote evolutionary comparisons [26]. Besides these general trends, ITS sequences exhibit substantial differences of divergence at a given taxonomic level from one taxon to another. For example, ITS sequences were sufficiently divergent to assess the timing of speciation within or among closely related Anthyllis and

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Gaertnera species [63, 75]. Conversely, Leskinen and Alström-Rapaport [66] reported very low interspecific variability among Salix species, in contradiction with the age of the group according to evidence from fossil data. Similar levels of variation in molecular diversity across lineages have been shown in bryophytes [38]. At the species level, the ITS appears extremely useful to resolve relationships within the leafy liverwort genus Plagiochila [41-44, 45-46, 98]. Shaw [101] even documented extensive ITS variation among populations of two widespread acrocarpous mosses, whereas similar levels of divergence are reached only among families in pleurocarpous mosses [119-120]. Such differences in genetic differentiation may be due to shifts in rates of molecular evolution. However, another possibility is simply that the taxa which are being compared are not ‘equivalent’. Indeed, Shaw [101] argued that the pattern in genetic differentiation among populations of a morphological species may indicate the presence of cryptic species. By contrast, taxonomically recognized species might not show the genetic differences that would be expected from their morphology, suggesting that their morphological diversity may to a large extent reflect habitat conditions [120]. In this context, the evolutionary conserved subportions of ITS2, apparently necessary for positioning of the multimolecular transcript processing machinery, also provide material for distinguishing evolutionarily rare events such as compensatory base changes in the relatively conserved regions, that might be useful in recognizing how arbitrary are the assignments of classical taxonomic ranks [26]. CONCLUSION

The utility of the ITS in angiosperm phylogenetics has been largely discussed and documented [49]. The use of ITS also seems promising for resolving taxonomic issues in bryophytes, pteridophytes, and maybe also gymnosperms. Although ITS primarily appears as a marker best suited to resolve issues at low levels, the existence of conserved motifs indicates that ITS may also be used at high levels. Such a potential may be explored in taxa where phylogenetic relationships are still obscure, for instance, the high levels of the taxonomic system in the Bryopsida. ITS sequences can thus diagnose organismal origins and phylogenetic relationships at many levels. However, the phylogenetic signal of ITS sequences at a given level greatly varies from a taxon to another. Below the species level, ITS divergence is in many cases too little to resolve

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relationships [8-9]. Additional polymorphic nuclear markers are needed to infer infraspecific genealogies. The use of single-copy genes has been increasingly urged to resolve the issue of the lack of polymorphism of ITS at low levels and the problems of homology assessment inherent to the use of multi-copy genes [1]. However, the summary of data gathered by Alvarez and Wendel [1] suggests that only 1 out of the 15 nuclear genes provided more variation than the ITS region, but this exception is potentially an artefact of paralogy [5]. This suggests that screening of a wide range of possible low copy number nuclear genes will be necessary in order to identify amplifiable regions with sufficient variation [31]. In addition, among the dozen of ‘universal’ primers that have been designed [80] for such introns as that of the nitrate-reductase [53], many, if not all, do not allow for the successful amplification of the region in a range of taxa [6]. In this context, the use of sequence characterized amplified regions, taking advantage of the fast and easy RAPD method but improving its robustness and reproducibility through the re-designing of internal primers specific for the selected target region, might prove extremely useful to find new variable regions [6, 105]. References [1] Alvarez I, Wendel JF. Ribosomal ITS sequences and plant phylogenetic inference. Mol Phylogenet Evol 2003; 29; 417–434. [2] An SS, Friedel T, Hegewald E. Phylogenetic relationships of Scenedesmus and Scenedesmus-like coccoid green algae as inferred from ITS-2 rDNA sequence comparison. Plant Biol 1999; 1: 418–428. [3] Avise JC. Phylogeography. Cambridge, Mass., USA: Harvard University Press, 2000. [4] Bailey CD, Price RA, Doyle JJ. Systematics of the Halimoglobine Brassicaceae: evidence from three loci and morphology. Syst Bot 2002; 27: 318–332. [5] Bailey CD, Carr TG, Harris SA, Hughes CE. Characterization of angiosperm nrDNA polymorphism, paralogy, and pseudogenes. Mol Phylogenet Evol 2003; 29: 435–455. [6] Bailey CD, Hughes CE, Harris SA. Using RAPD’s to identify DNA seaquence loci for species level phylogeny reconstruction: an example from Leucaena (Fabaceae). Syst Bot 2004; 29: 4–14. [7] Baldwin BG. Phylogenetic utility of the internal transcribed spacers of nuclear ribosomal DNA in plants: an example from the Compositae. Mol Phylogenet Evol 1992; 1: 3–16. [8] Baldwin BG, Markos S. Phylogenetic utility of the external transcribed spacer (ETS) of 18S-26S rDNA: congruence of ETS and ITS trees of Calycadenia (Compositae). Mol Phylogenet Evol 1998; 10: 449–463.

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Molecular Evolution of the =JFB-H>?L Intergenic Spacer in Bryophytes MICHAEL STECH1 and DIETMAR QUANDT2 1

Institut für Biologie - Systematische Botanik und Pflanzengeographie, Freie Universität Berlin, D-14195 Berlin, Germany 2 Institut für Botanik, Technische Universität Dresden, D-01062 Dresden, Germany

ABSTRACT The atp B-rbcL intergenic spacer of the chloroplast (cp) DNA is characterized in bryophytes (liverworts and mosses) based on more than 500 sequences of about 300 species of almost all major lineages (classes and subclasses), including comparisons of lengths, GC-contents, and sequence similarities as well as analyses of promoter elements, hairpin secondary structures and simple sequence repeats. The spacer displays a mosaic structure of (hyper-)variable segments and more conserved segments, the latter comprising one putative PEP promoter each for atp B and rbc L, a second putative -35 element, and a Shine-Dalgarno-like sequence 5´ to rbc L. Sequence variation is higher in liverworts than in mosses, and only the conserved segments are unambiguously alignable across all bryophytes. Phylogenetic analyses of mosses indicate phylogenetic utility of the at p B-rbc L spacer also at higher taxonomic levels. Molecular evolutionary characteristics are compared between the atp B-rbc L spacer and the non-coding parts of the trn T-F region. Key Words: Bryophytes, at p B-rb c L intergenic spacer, molecular evolution, phylogenetic utility, promoter elements, inversions

Address for correspondence: Michael Stech, Institut für Biologie - Systematische Botanik und Pflanzengeographie, Freie Universität Berlin, Altensteinstraße 6, D-14195 Berlin, Germany. Tel: +49 (0) 30 838 56541, Fax: +49 (0) 30 838 55434, E-mail: [email protected].

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INTRODUCTION

In the first volume of the Plant Genome series we provided an overview of molecular phylogenetic research within land plants, with an emphasis on bryophytes [45]. At that time, the trnT-trnL and trnL-trnF spacers, and especially the trnL intron of the trnT-F region were the only noncoding markers of the plastid genome that were extensively used for addressing systematic relationships among land plants at various taxonomic levels [45; Fig. 1]. However, even for these regions little was known about evolutionary patterns and constraints in the different land plant lineages and their possible influence on phylogenetic reconstructions. Therefore, detailed analyses were performed of the molecular evolution and phylogenetic utility of the trnT-F region in different land plant groups, namely hornworts [61], liverworts and mosses [46, 47], basal angiosperms [6] and land plants in general [44], based on more than 1,000 sequences altogether. These investigations included comparisons of lengths, GC-contents and putative promoter elements, secondary structure calculations of the trnL intron and detection of compensatory base pair changes, analyses of repeat motives probably resulting from slipped-strand mispairing, and detection of homoplastic inversions in loops of hairpin structures. Especially the latter mutations can decrease phylogenetic structure of molecular datasets and thus lead to erroneous tree topologies [43]. Most recently, other non-coding introns and spacers of the plastid genome have become of interest for resolving systematic relationships within bryophytes, such as the psbT-psbH spacer [for example, 17, 42, 62], the rpl16 intron [for example, 41], the trnG intron [for example, 40], and especially the atpB-rbcL spacer. The atpB-rbcL spacer separates the genes atpB, coding for the beta subunit of ATP-synthase, and rbcL,

Fig. 1. Schematic representation of the atpB-rbcL spacer in bryophytes. Approximate positions of putative atpB and rbcL promoters situated in more conserved segments (black squares) and (hyper-)variable segments (V1 – V4) are indicated according to the length of the alignment of mosses (see text for details).

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coding for the large subunit of Rubisco. As atpB and rbcL are transcribed in opposite directions, the spacer contains promoter elements for both genes (Fig. 1) and is thus expected to display a mosaic structure of conserved and variable segments evolving under different evolutionary constraints. Knowledge of molecular evolutionary characteristics of the atpB-rbcL spacer is, therefore, important for phylogenetic reconstructions. In angiosperms, not only the genes atpB and rbcL, but also the atpBrbcL spacer has been utilized as a molecular marker for more than a decade [for example, 2, 3, 10, 12, 16, 22, 31, 37, 48, 49, 50, 53]. In 1998, Chiang and colleagues [11] reported on universal primers for amplification of the atpB-rbcL spacer in a wide range of land plants, and provided the first sequences of bryophytes, Rhytidiadelphus loreus (Hedw.) Warnst. and R. triquetrus (Hedw.) Warnst. (Hylocomiaceae). Two years later, analyses of the molecular evolution and phylogenetic utility of the atpB-rbcL spacer in the Hylocomiaceae [8] and in mosses in general [9] were published, however, based on relatively small data sets. Further phylogenetic studies of bryophytes using the atpB-rbcL spacer have been published only recently, mainly dealing with mosses [for example, 33, 40, 41, 57, 58, 63, 67, 68, 69], but also with liverworts [52]. Several hundreds of sequences are now available, allowing us to characterize the atpB-rbcL spacer in more detail. In this chapter, efforts have been made to present an overview of molecular evolutionary characteristics of the atpB-rbcL spacer based on sequences from almost all bryophyte classes and subclasses, including comparisons of lengths, GC-contents, and sequence similarities, analyses of putative promoter elements, hairpin structures and simple sequence repeats in (hyper-) variable segments, as well as phylogenetic inferences. Molecular evolutionary patterns are assessed on different taxonomic levels, from the major lineages to intraspecific variation, and compared with previous analyses of the trnT-F region. MATERIAL AND METHODS Sampling of sequences

More than 500 atpB-rbcL spacer sequences of about 300 species representing almost all major bryophyte lineages (classes and subclasses) were compiled, most of them from GenBank and previous investigations

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conducted by us. New sequences were generated especially from representatives of classes and subclasses not sequenced so far, both liverworts [Treubiopsida: Treubia lacunosa (Colenso) Prosk. AY864310; Haplomitriopsida: Haplomitrium chilensis R.M. Schust. AY864305; Blasiopsida: Blasia pusilla L. AY864308; MarchantiopsidaSphaerocarpidae: Sphaerocarpos donnellii Austin DQ269023, Monocleidae: Monoclea gottschei Lindb. AY864309; JungermanniopsidaMetzgeriidae: Hymenophyton flabellatum (Labill.) Dumort. AY864306, Metzgeria furcata (L.) Corda DQ269022, Pellia epiphylla (L.) Corda AY864307] and mosses [Sphagnopsida: Sphagnum fallax H. Klinggr. AY864303, S. palustre L. AY864304, Takakiopsida: Takakia lepidozioides S. Hatt. & Inoue AY864296; Andreaeopsida: Andreaea rupestris Hedw. AY864297; Polytrichopsida: Dendroligotrichum dendroides (Brid. ex Hedw.) Broth. AY864299, D. squamosum (Hook. f. & Wilson) Cardot AY864300, Notoligotrichum angulatum (Cardot & Broth.) G.L. Sm. AY864301, Oligotrichum canaliculatum (Hook. & Arn.) Mitt. AY864302, Bryopsida-Buxbaumiidae: Buxbaumia aphylla Hedw. AY864298]. Further sequences of Jungermanniopsida-Jungermanniidae [Porella crispata (Hook.) Trevis. DQ269020, P. renifolia (Stephani) Swails DQ269021; Radula complanata (L.) Dumort. AY864311] and Bryopsida-Bryidae [Dicranoloma billardieri (Brid.) Paris AY864292, Hypopterygium discolor Mitt. AY864293, H. tamarisci (Sw.) Brid. ex Müll. Hal. AY864294, Lopidium concinnum (Hook.) Wilson AY864295] were also included. Additional sequences of Hypnodendron were kindly provided by C. Pawlak (Berlin). Protocols of the methods used for DNA extraction, amplification and sequencing are given in [57]. Two data sets were designed, the first including only one sequence of each species (300 sequences, Table 1, excluding the ambiguous sequences of Bazzania fauriana and Physcomitrella patens, see below), and the second comprising all species of which more than one voucher has been sequenced. The second data set was compiled to address intraspecific variation, while all other calculations and measurements were based on the first data set. Classification of mosses follows Goffinet and Buck [15] and those of liverworts follows Stech and Frey [59]. Alignment, Secondary Structures and Statistics

Alignments for both atpB-rbcL spacer data sets (see above) were manually performed in the Domix Alignment Editor v. 09/2004 [18] and corrected considering information obtained from secondary structure

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analyses of putative hairpin structures. Secondary structures were calculated using RNAstructure 4.1.1 [32]. Alignments are available from the authors upon request, together with a list of accession numbers and taxon abbreviations used. Sequence lengths and GC-contents were obtained using the statistics function of the alignment program. Sequence variation was calculated as p-difference (substitutions per compared nucleotides) as implemented in the statistics function of SeqState [35], with the standard error estimated via bootstrapping. Due to the limited numbers of sequences sampled from several lineages (cf. Table 1), sequence variation was compared only between Jungermanniopsida, Polytrichopsida, and Bryopsida (Table 2). Phylogenetic analyses

The suitability of the atpB-rbcL spacer for large-scale phylogenetic reconstructions at higher taxonomic levels was assessed in mosses, with species of Takakiopsida, Andreaeopsida, Polytrichopsida and Bryopsida as Table 1. Overview of the number of sequences sampled from the major bryophyte lineages (data set 1). Classification of mosses follows Goffinet and Buck [15], classification of liverworts follows Stech and Frey [59] Taxon Sphagnopsida Takakiopsida Andreaeopsida Polytrichopsida Bryopsida Buxbaumiidae Funariidae Dicranidae Bryidae Hypnidae Treubiopsida Haplomitriopsida Blasiopsida Marchantiopsida Jungermanniopsida

Abbreviation

No. of sequences

Sp Ta An Po

2 1 1 4

Bu Fu Di Br Hy Tr Ha Bl Ma Ju

1 1 47 93 114 1 1 1 3 31 5 301

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Table 2. Length ranges and sequence variation (p-difference, with the standard error [S.E.] estimated via bootstrapping) of selected bryophyte lineages

Mosses Polytrichopsida Bryopsida Jungermanniopsida

Length range [bp]

Mean

S.D.

p-distance [%]

S.E.

385-683 678-683 385-589 415-614

547,57 679,75 544,58 573,032

34,971 2,046 29,306 36,292

10,055 2,692 9,302 21,849

0,781 0,488 0,706 0,786

ingroup and both included Sphagnum species (Sphagnopsida) as outgroup representatives based on previous molecular analyses (summarized by Goffinet and Buck [15]). Inference of relationships between mosses and liverworts and across land plants is problematic, firstly because of the currently limited taxon sampling of several lineages, and secondly because only small alignable segments of the spacer could be used. Phylogenetic reconstructions according to the maximum parsimony principle were performed using PRAP [36], which allows running Nixon´s Parsimony Ratchet [38] with winPAUP 4.0b10 [66]. Heuristic searches were performed with the following options: all characters unweighted and unordered, multistate characters interpreted as uncertainties, performing TBR branch swapping, collapse zero length branches, MulTrees option in effect, random addition sequence with 1,000 replicates. In addition to calculations with gaps coded as missing data, indels were coded according to the simple indel coding method [56], as implemented in SeqState [35], in order to obtain the complete phylogenetic signal of the spacer. Heuristic bootstrap searches were performed with 10,000 replicates using the “fast bootstrap” option (bootstrapping without branch-swapping) of PAUP. Fast bootstrapping was shown to provide estimates of support similar to, although generally less than, bootstrapping with branch-swapping. RESULTS AND DISCUSSION Sequences

When analyzing the trnL intron in bryophytes based on more than 1,000 sequences [47], several difficulties were encountered with sequences taken from GenBank. About 15% of the sequences were incomplete

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towards the trnL 5’ exon (stem-loop regions P1 and P2), while <2.5% of the sequences displayed unusually high numbers of 1 nt indels in P1 – P3, which were proved artificial and most likely due to sequencing errors. In the atpB-rbcL spacer, sequencing of the region towards atpB was not problematic as the forward primer is situated further downstream in the atpB gene. However, in about 22% of the sequences from GenBank, 17 nt just in front of the rbcL gene were missing. This obviously artificial gap in the conserved region adjacent to rbcL was corrected by insertion of the appropriate number of N’s to circumvent an artificial higher degree of length variation, but could affect calculations of the GC-content. It is noteworthy that the spacer sequence of Physcomitrella patens (taken from the complete plastid genome sequence; [65]) is different from all other Bryopsida species in the part towards atpB, including the putative atpB promoter region, which has to be tested by sequencing further species of subclass Funariidae. Therefore, the sequence was only used for calculations of length and GC-values, but not included in the alignment and phylogenetic analyses. Another problem concerned misidentification of vouchers or mixing up of sequences. According to sequence comparison and phylogenetic inference, at least four atpB-rbcL spacer sequences obviously did not fit the assigned species names. The sequence of Bazzania fauriana (Stephani) S. Hatt. from GenBank (AJ249037) is clearly a moss sequence and has nothing in common with the sequence of Chiang and Schaal [9]. As its identification is totally unclear, the sequence was not considered in all calculations. Two further sequences assigned to species of the Hypnidae, Climacium dendroides (Hedw.) F. Weber & D. Mohr (AJ288395) and Antitrichia californica Sull. (AF413566), most probably belong to either Mniaceae or Hypnodendraceae (both Bryidae), respectively. The sequence of Bryum billarderi Schwägr. AF546832 seems to be more closely related to Rhodobryum and might thus be misidentified as well. However, the latter three sequences were included, as they are not identical with other sequences and probably represent different species. Length and GC-content

Length of the atpB-rbcL spacer ranges from 378 bp (Treubia lacunosa) to 683 bp (Notoligotrichum angulatum) in the investigated species. Compared with lengths of 800-1,000 bp in gymnosperms and angiosperms reported

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by Chiang and colleagues [11], the atpB-rbcL spacer is considerably shorter in bryophytes than in seed plants. However, this discrepancy seems to be less obvious than in the non-conding parts of the trnT-F region, especially the trnL-F spacer [44; Fig. 5]. In addition, sequence lengths comparable to those of bryophytes were also found in angiosperms, for example, in Anemone hupehensis (Lemoine) Lemoine (632 bp; [53]) or Euphorbia sect. Meleuphorbia (559-689 bp; [49]). Length variation between the different lineages of liverworts and mosses is shown in Fig. 2. In mosses, a trend of length reduction is observed from the more basal to the more derived lineages (“basal” and “derived” according to the position in molecular trees; [for example, 15]), similar to the trnL intron [47], although less prominent. In both markers, sequences are longer in Sphagnum, Takakia, and the Polytrichopsida than

Fig. 2. Length variation (in nt) of the atpB-rbcL spacer in the major bryophyte lineages. Sp = Sphagnopsida, Ta = Takakiopsida, An = Andreaeopsida, Po = Polytrichopsida, Bu = Buxbaumiidae, Fu = Funariidae, Di = Dicranidae, Br = Bryidae, Hy = Hypnidae, Tr = Treubiopsida, Ha = Haplomitriopsida, Bl = Blasiopsida, Ma = Marchantiopsida, Ju = Jungermanniopsida.

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in most of the Bryopsida. Comparatively short sequences are characteristic of Andreaea (Andreaeopsida). Within the Bryopsida, length variation of the trnL intron follows the same pattern of longer sequences in the more basal lineages, such as Diphysciidae, and shorter sequences in the more derived pleurocarpous Hypnidae. In the atpB-rbcL spacer, the basalmost included species of the Bryopsida, Buxbaumia aphylla (Buxbaumiidae), has the longest sequence. However, no tendency of length reduction is observed, as sequences of the Hypnidae are, on average, longer than sequences of the Bryidae and of Physcomitrella patens (Funariidae). In liverworts, the opposite situation of shorter sequences in the basalmost lineages (Treubiopsida and Haplomitriopsida; [14, 59, 60]) and longer sequences in the more derived leafy liverworts (Jungermanniopsida-Jungermanniidae) is observed. These different patterns may indicate different modes of evolution of the atpB-rbcL spacer in mosses and liverworts. DNA regions evolving under low functional constraints, such as non-coding spacers and pseudogenes, are typically AT-rich and GC-poor [28]. With GC-contents ranging from 14-24% in >90% of the investigated species, the atpB-rbcL spacer fits well into this general scheme. Higher GC-values are found in Takakia lepidozioides (28.8%) and Sphagnum spp. (36.1%) among the mosses, in the Metzgeriidae (Pellia epiphylla 30.4%, Hymenophyton flabellatum 32.7%), and especially in Haplomitrium chilensis (42.5%), which exactly matches the situation in all three non-coding parts of the trnT-F region [46, 47]. A comparison of the GC-content in the major moss lineages (Fig. 3) shows the decrease of GC-values from the Sphagnopsida and Takakiopsida to the Bryopsida, and slightly increasing GC-contents within the Bryopsida. Differences of nucleotide composition between the more basal and the more derived lineages are, therefore, similar to, but much more obvious than the patterns of length variation described above. In liverworts, the GCcontent is difficult to correlate with the relationships of the major lineages (Fig. 3). Alignments, repeats, and sequence variation

Chiang and Schaal [9; p. 424] stated that the atpB-rbcL spacer is “highly conserved both within mosses and between mosses and liverworts”. However, they compared only Marchantia polymorpha and Bazzania fauriana with a selection of 11 moss sequences, and even this data set

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Fig. 3. Variation of GC-content (in %) of the atpB-rbcL spacer in the major bryophyte lineages. Sp = Sphagnopsida, Ta = Takakiopsida, An = Andreaeopsida, Po = Polytrichopsida, Bu = Buxbaumiidae, Fu = Funariidae, Di = Dicranidae, Br = Bryidae, Hy = Hypnidae, Tr = Treubiopsida, Ha = Haplomitriopsida, Bl = Blasiopsida, Ma = Marchantiopsida, Ju = Jungermanniopsida.

showed considerable differences between liverworts and mosses. Due to high sequence divergence in the present larger data set, it is in fact impossible to align the complete atpB-rbcL spacer unambiguously across liverworts and mosses, and within liverworts. In mosses, alignment is quite straightforward in the Bryopsida, but more difficult between the Bryopsida and the other classes, especially the Sphagnopsida. Because of the difficulties in aligning mosses and liverworts, the sequences of both major bryophyte groups were aligned separately. The tentative alignments of mosses and liverworts comprise 1,770 and 1,922 positions, respectively. As indicated in Fig. 1, length variation is not equally distributed across the alignment. Most conserved are the segments around the putative promoter elements of atpB and rbcL (see below), and about 50 bp at the 5´ end just in front of rbcL. The latter part probably contains a Shine-Dalgarno (SD) sequence within an AGGGAGGG stretch, whereas the 5´-untranslated part adjacent to atpB is more variable and does not contain any SD-like sequence, similar to

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the situation reported from tobacco [20]. Translational control of atpB and rbcL might thus be different in bryophytes, which should be tested experimentally. Another conserved element (GGTCAA in >90% of all investigated species) is situated about 190 positions downstream of the— 35 element of the rbcL promoter, which might belong to another atpB promoter (see below). The middle part of the spacer (V2 in Fig. 1) is the most lengthvariable segment. Similar to the most variable stem-loop region P8 of the trnL intron and the trnT-L spacer [46, 47], it is AT-rich and consists of repetitive elements of various sizes, as shown in Fig. 4 for the Bryopsida. Short repeats of only two nucleotides, for example, AT, are common, but also longer elements up to 21 nt are found, similar to the P8 region of the trnL intron. The shortest possible indels of only one nucleotide are much more numerous and sometimes difficult to define, and were, therefore, not considered in Fig. 4. Most repeats are probably derived by duplications of adjacent sequence elements (black bars in Fig. 4), events that are most likely the result of slipped-strand mispairing (SSM) [for example, 27, 71]. SSM was found to be the predominant mechanism of indel genesis in the plastid rpl16 intron [24], and has also been suggested to play a role in the evolution of the atpB-rbcL spacer in higher plants [12, 16]. In contrast, some present nucleotide stretches (white bars in Fig. 4) may either originate from farther regions of the spacer, or their repeat origin may be masked by subsequent mutations. Similar to earlier observations on sequence variation within bryophytes [46], mosses in general, and especially the Hypnidae (3.429%, S.E. 0.321), display less variation in non-coding DNA than liverworts, which is illustrated by the almost double p-distance within the Jungermanniopsida compared to the Bryopsida (Table 2). Promoter Elements

Genes associated with photosynthesis, such as rbcL, are thought to be mainly transcribed by the plastid-encoded RNA polymerase (PEP) in non-parasitic higher plants [for example, 1, 13]. PEP promoters are similar to eubacterial promoters and consist of two elements, the -35 and -10 elements [70]. In contrast, the nuclear-encoded RNA polymerase(s) (NEP) are mainly responsible for transcription of housekeeping genes [19, 29], and for plastid genes in general in most holoparasitic plants [for example, 4]. NEP promoters are more variable than PEP promoters and

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Fig. 4. Numbers of repeats (black) and nucleotide stretches not obviously originating from adjacent regions (white) classified according to their length in nt in the atpB-rbcL spacer in the Bryopsida.

include a YRTA motif that shows homology to bacteriophage and mitochondrial promoters [70]. For transcription of rbcL, a single PEP promoter was reported from angiosperms [for example, 12, 30, 39, 50]. The respective promoter elements were also found in the present alignments of mosses and liverworts. As shown in Table 3, the -35 and -10 elements are highly conserved across the major lineages (50% majority rule consensus sequences TTGCAT and TACAAT, mostly separated by 18 nt), except for Haplomitrium chilensis (Haplomitriopsida) with a single substitution in the -10 element (TAGAAT), and Sphagnum (Sphagnopsida). A comparison of atpB-rbcL spacer sequences from about 500 genera of angiosperms compiled from GenBank also revealed little variation within the promoter elements. The -10 element is the same in most angiosperms and bryophytes, whereas the -35 element is TTGCGC in most studied angiosperm sequences. The promoter of Sphagnum (CTGCAG and TACATG, with a spacing of 22 nt) is extremely different that it might

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no longer be functional, which is supported by the presence of a second, more typical, putative promoter motif situated closer to rbcL (TTGATC and TACGAT, separated by 17 nt). In contrast to rbcL, transcriptional control of atpB (co-transcribed with atpE at least in Arabidopsis; [23]) seems to be more variable. In angiosperms, different combinations of one to three functional PEP promoters and/or one NEP promoter were reported [for example, 12, 30, 39, 49, 50, 55]. In the Rubiaceae, different numbers of promoters occur even within one family, which was supposed to correlate with ecological peculiarities, as tropical woody species have three functional PEP promoters, but temperate herbaceous species only one [30]. The present analysis indicates a simple condition in bryophytes, with only one putative functional PEP promoter present in all liverworts and mosses. According to its position (Fig. 1), this promoter may correspond to the atpB-proximal promoter reported from some angiosperms [for example, 49, 50]. Its functionality, however, remains to be tested experimentally, especially in the light of the considerable variation between the different lineages (Table 3). A second putative -35 element (GGTCAA in >90% of all investigated species, i.e., TTGACC on the complementary strand) is situated closer to rbcL, separating the variable regions V2 and V3 (Fig. 1). This element might belong to a second promoter corresponding to the atpB-distal promoter in angiosperms; however, its functionality is questionable, as a -10 element could not be detected in most of the sequences. Hairpin Secondary Structures

The first putative hairpin secondary structure formed by dyad symmetrical elements in the atpB-rbcL spacer was reported from the moss genus Campylopus [57]. It was suspicious that in the variable middle part of the spacer (V2 region, cf. Fig. 1) some Campylopus species showed an A-rich segment, while other species had T-rich sequences. Secondary structure analyses revealed that these segments (23-31 nt) comprised the loop of a hairpin. The A- and T-types originated from an inversion of the loop region. Smaller inversions (less than 10 bp) in loops of hairpin structures have so far been reported in a few studies of land plants, for example, in the atpB-rbcL spacer of Epacridaceae [12], the rpl16 intron and atpB-rbcL spacer of some Poaceae [16, 25], the psbA-trnH spacer in some Paeonia species [51], the trnL-F spacer of bryophytes [46], and in

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Table 3. Majority rule consensus sequences of putative sigma70-type promoter elements (-35 and -10 elements) of the atpB and rbcL genes present in the atpBrbcL spacer in bryophytes

rbcL rbcL rbcL rbcL atpB atpB atpB atpB atpB atpB atpB atpB

(mosses excl. Sphagnum) (Sphagnum) (liverworts excl. Haplomitrium) (Haplomitrium) (mosses excl. Sphagnum, Takakia) (Sphagnum) (Takakia) (Treubia) (Haplomitrium) (Blasia) (Marchantiopsida) (Jungermanniopsida)

-35

-10

TTGCAT CTGCAG TTGCAT TTGCAT TTGATA TTGATA TTGAAA TTAACA CTGATC TTGATA TTDATA YCGATA

TACAAT TACATG TACAAT TAGAAT TATAAT TACGAT TACAAT TATACT TACGAT TACAAT TAYAAT TACAAT

the psbT-psbN spacer of green algae, bryophytes and higher plants [43], which can cause a significant loss in phylogenetic structure if they remain undetected in phylogenetic analyses [43]. Although hairpin structures in 3´ untranslated regions of genes may be functional and involved in, for example, transcription termination, cleavage site recognition or regulation of mRNA accumulation and stability [for example, 5, 21, 64], the hairpin structure in the atpB–rbcL spacer of Campylopus probably is not functional. As discussed by Stech [57], it is situated in a 5´ untranslated region, and in four Campylopus species nearly all the dyad symmetrical element is deleted. Comparison of this region over a broad range of mosses further supports its non-functionality, because in the other genera no such dyad symmetrical element was found. By inspection of the moss alignment in the present study, two further regions were presumed to form hairpin secondary structures, both situated in the variable region V4 between the rbcL promoter and the conserved region 5´ to rbcL (Fig. 1). In contrast to the hairpin in Campylopus, these structures can be formed in all investigated moss species. One of these hairpins comprises a stem of at least 8 bp and a small loop of 4-5 nt (Fig. 5). In some species the stem can extend up to 12 bp. Similar to the examples described above, an inversion of the loop region was observed, resulting in an A-rich and a T-rich type. The A-rich

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Fig. 5. Examples of hairpin secondary structures formed by the short dyad symmetrical element in the variable region V4 of the atpB-rbcL spacer of mosses, displaying an inversion of the unpaired loop region (A-rich type in Oligotrichum canaliculatum and Campylopus flexuosus, T-rich type in Acidodontium ramicola and Hypopterygium discolor).

type occurs in >90% of the investigated moss species, whereas the T-rich type is rare, and obviously developed independently in the Dicranidae (Tortula muralis Hedw.), Bryidae (Acidodontium ramicola [Spruce ex Mitt.] A. Jaeger, Fig. 5; but not in other Acidodontium species analyzed by Pedersen and colleagues [40]), and Hypnidae (Hypopterygiaceae, for example, Hypopterygium discolor, Fig. 5; and Hookeria lucens [Hedw.] Sm.). The second hairpin structure is larger, with a stem of 13-15 bp and a loop of 6-12 nt (Fig. 6). In this structure, only two Bryum species (B. bicolor Dicks., B. caespiticium Hedw.) show a putative inversion of the loop region from A-type to T-type. Formation of the hairpin is supported by compensatory base-pair changes (CBC), as can be seen, for example, between Dicranum and Bryum (Fig. 6). Two substitutions leading to the opening of the helix are characteristic for most Bryidae (except Bryaceae and Mniaceae) and Hypnidae. Interestingly, in the Hypopterygiaceae further compensatory mutations occur that close the helix again (cf. Palustriella falcata [Brid.] Hedenäs vs. Hypopterygium discolor in Fig. 6). Phylogenetic Inference

Maximum parsimony analyses were based on 432 parsimony-informative positions without indel coding or 746 positions with indel coding, respectively. Heuristic searches recovered 1,562 trees without indel coding (length 2,236 steps, CI = 0.437, RI = 0.846) or 502 trees with indel coding (length 3,302 steps, CI = 0.473, RI = 0.839). The strict consensus trees of both analyses are of almost congruent topology.

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Fig. 6. Examples of hairpin secondary structures formed by the long dyad symmetrical element in the variable region V4 of the atpB-rbcL spacer of mosses. See text for further explanation.

Relationships of the major moss lineages, as inferred from the strict consensus tree of the analysis with indel coding (shown in a simplified form in Fig. 7), are largely consistent with the “total evidence” molecular tree of Goffinet and Buck [15]. This includes (i) the basal positions of Takakiopsida and Andreaeopsida (the position of Buxbaumia needs further consideration), (ii) the sistergroup relationship of Polytrichopsida (100% bootstrap support) and Bryopsida excl. Buxbaumia (99% BS), (iii) monophyly of haplolepideous mosses (Dicranidae, 92% BS), (iv) paraphyly of Bryidae, with Rhizogonianae plus Aulacomniaceae separated from the (remaining) Bryanae and more closely related to the Hypnidae, and (v) monophyly of the Hypnidae and sistergroup relationship of Hookeriales and Hypnales. Hence, the atpB-rbcL spacer seems to reflect, at least to some extent, the evolutionary history of mosses and provides reliable information for phylogenetic reconstruction, although only few internal clades at the ordinal and family levels are well-supported by bootstrap analysis, for example, Leucobryaceae sensu La Farge and colleagues [26] (100%), Bryaceae (100%), and Aulacomniaceae (90%). Similar to other studies employing only one or two regions [for example, 7], practically no resolution within the Hypnales is achieved using the atpB-rbcL spacer alone.

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Fig. 7. Simplified representation of a strict consensus tree of 502 most parsimonious trees (with simple indel coding according to Simmons and Ochoterena [56]; length 3302 steps, CI = 0.473, RI = 0.839) inferred from a heuristic search of atpB-rbcL spacer sequences of 300 moss species belonging to the classes Bryopsida, Polytrichopsida, Andreaeopsida and Takakiopsida, with two species of Sphagnum (Sphagnopsida) as outgroup representatives. Bootstrap values >50% are above the branches. Numbers in brackets indicate the number of sequences included in the respective clades.

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Intraspecific Sequence Variation

For about 70 species (almost 25% of the sampled species), two or more atpB-rbcL spacer sequences from different populations are available. However, only few species have so far been analyzed in more detail at the population level, mainly Amblystegium spp. [69], Campylopus spp. [58, 63], and Pyrrhobryum mnioides (Hook.) Manuel [33]. According to these studies, intraspecific sequence variation seems to be a common phenomenon in the atpB-rbcL spacer in bryophytes. Sequence divergence corresponding to large-scale biogeographical patterns was observed, for example, in Campylopus flexuosus (Hedw.) Brid. (African and European/ South American clades; [63]), C. pilifer Brid. (Old World, South American, and Central/North American clades; [58]), and Pyrrhobryum mnioides (Australian/New Zealand and South American clades; [33]). As the respective Campylopus species are not monophyletic at the molecular level, the above mentioned clades may represent cryptic species. CONCLUSION

Many features of the atpB-rbcL spacer resemble those of the non-coding parts of the trnT-F region, for example, a mosaic structure of conserved and variable segments, short sequence lengths in comparison with seed plants, low GC-contents except for some lineages of mosses (Sphagnopsida, Takakiopsida) and liverworts (Haplomitriopsida, Jungermanniopsida-Metzgeriidae), and a higher sequence variation in liverworts than in mosses. Similar to the situation found in the trnT-F region, the pleurocarpous Hypnidae show a considerably low sequence variation, perhaps due to the rather rapid radiation in the early history of this group [54]. This low variation accounts for the current problems in resolving relationships among the pleurocarpous mosses even using non-coding DNA that is known to be relatively highly variable. A successful approach to resolve the Hypnalean lineages will thus need the combination of a substantial amount of regions to obtain a useful phylogenetic signal and structure. Patterns of variation in non-coding cpDNA regions, such as the atpB-rbcL spacer or the spacers and intron of the trnT-F region, seem to be comparable between bryophytes and seed plants. Within certain lineages these non-coding regions are largely alignable, whereas homology assessment between rather distant lineages (for example, between liverworts and mosses, or between Gnetales, other gymnosperms and angiosperms; [44]) is almost impossible at least for the variable segments.

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Nevertheless, both the atpB-rbcL spacer and the trnT-F region seem to allow congruent circumscriptions and delimitations of the bryophyte classes. As they are not functional, large parts of the atpB-rbcL spacer may evolve under low evolutionary constraints, thus more or less independently in the higher taxa, whereas in the functional segments (for example, promoter regions) possibilities of mutations are lower, probably resulting in homoplastic character states in non-related lineages. Phylogenetic utility of the atpB-rbcL spacer is, therefore, expected to be limited at higher taxonomic levels. However, the atpB-rbcL spacer can be a suitable marker for phylogenetic reconstructions at least for mosses, and combined analyses of several introns and spacers of the plastid genome, in combination with coding indels as additional characters, may result in phylogenies that are equally well-resolved as phylogenies based on coding genes. So far, only the trnT-F region has been investigated in more detail, and in particular the more conserved parts of the trnL intron have proved to be suitable to infer relationships between the major bryophyte lineages [44]. Acknowledgments

Sincere thanks to Dr. W. Frey, Dr. J.-P. Frahm and Dr. C. Neinhuis for their advice and encouraging support, to C. Pawlak for providing sequences, to the curator of B, Dr. H. Nowak-Krawietz, for loan of plant material, and to B. Giesicke for technical assistance. Furthermore, we thank the German Research Foundation (DFG) as well as the German Academic Exchange Service (DAAD), for financially supporting our research during the past few years. References [1] Allison LA, Simon LD, Maliga P. Deletion of rpoB reveals a second distinct transcription system in plastids of higher plants. EMBO J 1996; 15: 2802–2809. [2] Azuma H, Thien LB, Kawano S. Molecular phylogeny of Magnolia (Magnoliaceae) inferred from cpDNA sequences and evolutionary divergence of the floral scents. J Plant Res 1999; 112: 291–306. [3] Bell CD. Preliminary phylogeny of Valerianaceae (Dipsacales) inferred from nuclear and chloroplast DNA sequence data. Mol Phylogenet Evol 2004; 31: 340–350. [4] Berg S, Krause K, Krupinska K. The rbcL genes of two Cuscuta species, C. gronovii and C. subinclusa, are transcribed by the nuclear-encoded plastid RNA polymerase (NEP). Planta 2004; 219: 541–546.

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[5] Blowers AD, Klein U, Ellmore GS, Bogorad L. Functional in vivo analysis of the 3´ flanking sequences of the Chlamydomonas chloroplast rbcL and psaB genes. Mol Gen Genet 1993; 238: 339–349. [6] Borsch T, Hilu KW, Quandt D, Wilde V, Neinhuis C, Barthlott W. Non-coding plastid trnT-trnF sequences reveal a well resolved phylogeny of basal angiosperms. J Evol Biol 2003; 16: 558–576. [7] Buck WR, Goffinet B, Shaw AJ. Testing morphological concepts of orders of pleurocarpous mosses (Bryophyta) using phylogenetic reconstructions based on trnLtrnF and rps4 sequences. Mol Phylogenet Evol 2000; 16:180–198. [8] Chiang T-Y, Schaal BA. Molecular evolution of the atpB-rbcL noncoding spacer of chloroplast DNA in the moss family Hylocomiaceae. Bot Bull Acad Sin 2000; 41: 85–92. [9] Chiang T-Y, Schaal BA. Molecular evolution and phylogeny of the atpB-rbcL spacer of chloroplast DNA in the true mosses. Genome 2000; 43: 417–426. [10] Chiang T-Y, Chiang YC, Chen YJ, Chou CH, Havanond S, Hong TN, Huang S. Phylogeography of Kandelia candel in East Asiatic mangroves based on nucleotide variation of chloroplast and mitochondrial DNAs. Mol Ecol 2001; 10: 2697–2710. [11] Chiang T-Y, Schaal BA, Peng CI. Universal primers for amplification and sequencing a noncoding spacer between the atpB and rbcL genes of chloroplast DNA. Bot Bull Acad Sin 1998; 39: 245–250. [12] Crayn DM, Quinn CJ. The evolution of the atpb-rbcL intergenic spacer in the Epacrids (Ericales) and its systematic and evolutionary implications. Mol Phylogenet Evol 2000; 16: 238–252. [13] deSantis-Maciossek G, Kofer W, Bock A, Schoch S, Maier RM, Wanner G, Rüdiger W, Koop HU, Herrmann RG. Targeted disruption of the plastid RNA polymerase genes rpoA, B and C1: molecular biology, biochemistry and ultrastructure. Plant J 1999; 18: 477–489. [14] Forrest LL, Crandall-Stotler BJ. A phylogeny of the simple thalloid liverworts (Jungermanniopsida, Metzgeriidae) as inferred from five chloroplast genes. Monogr Syst Bot Missouri Bot Gard 2004; 98: 119–140. [15] Goffinet B, Buck WR. Systematics of the Bryophyta (mosses): from molecules to a revised classification. Monogr Syst Bot Missouri Bot Gard 2004; 98: 205–239. [16] Golenberg EM, Clegg MT, Durbin ML, Doebley J, Ma DP. Evolution of a noncoding region of the chloroplast genome. Mol Phylogenet Evol. 1993; 2: 52–64. [17] He-Nygrén X, Piippo, S. Phylogenetic relationships of the generic complex Chiloscyphus-Lophocolea-Heteroscyphus (Geocalycaceae, Hepaticae): Insights from three chloroplast genes and morphology. Ann Bot Fennici 2003; 40: 317–329. [18] Hepperle D. Domix Alignment Editor v. 09/2004. Program distributed by the author, Neu-Globsow, 2002. [19] Hess W, Börner T. Organellar RNA polymerases of higher plants. Int Rev Cytol 1999; 190: 1–59. [20] Hirose T, Sugiura M. Multiple elements required for translation of plastid atpB mRNA lacking the Shine-Dalgarno sequence. Nucleic Acids Res 2004; 32: 3503–3510. [21] Hong L, Stevenson JK, Roth WB, Hallick RB. Euglena gracilis chloroplast psbB, psbT, psbH and psbN gene cluster: regulation of psbB-psbT pre-mRNA processing. Mol Gen Genet 1995; 247: 180–188.

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[22] Hoot SB, Douglas AW. Phylogeny of the Proteaceae based on atpB and atpB-rbcL intergenic spacer region sequences. Australian Syst Bot 1998; 11: 301–320. [23] Isono K, Niwa Y, Satoh K, Kobayashi H. Evidence for transcriptional regulation of plastid photosynthetic genes in Arabidopsis thaliana roots. Plant Physiol 1997; 114: 623–630. [24] Kelchner SA, Clark LG. Molecular evolution and phylogenetic utility of the chloroplast rpl16 intron in Chusquea and the Bambusoideae (Poaceae). Mol Phylogenet Evol 1997; 8: 385–397. [25] Kelchner SA, Wendel JF. Hairpins create minute inversions in non-coding regions of chloroplast DNA. Curr Genet 1996; 30: 259–262. [26] La Farge C, Shaw AJ, Vitt DH. The circumscription of the Dicranaceae (Bryopsida) based on the chloroplast regions trnL-trnF and rps4. Syst Bot 2002; 27: 435–452. [27] Levinson G, Gutman, GA. Slipped-strand mispairing: a major mechanism for DNA sequence evolution. Mol Biol Evol 1987; 4: 203–221. [28] Li WH. Molecular evolution. Sinauer, Sunderland, Massachusetts, Publisher 1997. [29] Maliga P. Two plastid RNA polymerases of higher plants: an evolving story. Trends Plant Sci 1998; 3: 4–6 [30] Manen J-F. Relaxation and evolutionary constraints in promoters of the plastid gene atpB in a particular Rubiaceae lineage. Plant Syst Evol 2000; 224: 235–241. [31] Manen J-F, Natali A, Ehrendorfer F. Phylogeny of the Rubiaceae-Rubieae inferred from the sequences of a cpDNA intergene region. Plant Syst Evol 1994; 190: 195– 211. [32] Mathews DH, Disney MD, Childs JL, Schroeder SJ, Zuker M, Turner DH. Incorporating chemical modification constraints into a dynamic programming algorithm for prediction of RNA secondary structure. Proc Natl Acad Sci USA 2004; 101: 7287–7292. [33] McDaniel SF, Shaw AJ. Phylogeographic structure and cryptic speciation in the transAntarctic moss Pyrrhobryum mnioides. Evolution 2003; 57: 205–215. [34] Mort ME, Soltis PS, Soltis DE, Mabry ML. Comparison of three methods for estimating internal support on phylogenetic trees. Syst Biol 2000; 49: 160–171. [35] Müller K. SeqState: primer design and sequence statistics for phylogenetic DNA datasets. Appl Bioinformatics 2005; 4: 65–69. [36] Müller K. PRAP - calculation of Bremer support for large data sets. Mol Phylogenet Evol 2004; 31: 780–782. [37] Natali A, Manen J-F, Ehrendorfer F. Phylogeny of the Rubiaceae-Rubioideae, in particular the tribe Rubieae: evidence from a non-coding chloroplast DNA sequence. Ann Missouri Bot Gard 1995; 82: 428–439. [38] Nixon KC. The parsimony ratchet, a new method for rapid parsimony analyses. Cladistics 1999; 15: 407–414. [39] Orozco EM, Chen L-J, Eilers RJ. The divergently transcribed rbcL and atpB genes of tobacco plastid DNA are separated by nineteen base pairs. Curr Genet 1990; 17: 65–71. [40] Pedersen N, Cox CJ, Hedenäs L. Phylogeny of the moss family Bryaceae inferred from chloroplast DNA sequences and morphology. Syst Bot 2003; 28: 471–482. [41] Pedersen N, Hedenäs L. Phylogenetic investigations of a well supported clade within the acrocarpous moss family Bryaceae: evidence from seven chloroplast DNA sequences and morphology. Plant Syst Evol 2003; 240: 115–132.

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[42] Quandt D, Huttunen S. Evolution of pendent life-forms in bryophytes. J Hattori Bot Lab 2004; 95: 207–217. [43] Quandt D, Müller K, Huttunen S. Characterisation of the chloroplast DNA psbT-H region and the influence of dyad symmetrical elements on phylogenetic reconstructions. Plant Biol 2003; 5: 400–410. [44] Quandt D, Müller K, Stech M, Frey W, Hilu KW, Borsch T. Molecular evolution of the chloroplast trnL-F-region in land plants. Monogr Syst Bot Missouri Bot Gard 2004; 98: 13–37. [45] Quandt D, Stech M. Molecular systematics of bryophytes in context of land plant phylogeny. In: Sharma AK, Sharma A. eds. Plant Genome: Biodiversity and Evolution, Vol. 1, Part A. Science Publishers, Enfield, NH, USA, 2003: 267–295. [46] Quandt D, Stech M. Molecular evolution of the trnTUGU-trnF GAA region in bryophytes. Plant Biol 2004; 6: 545–554. [47] Quandt D, Stech M. Molecular evolution and secondary structure of the chloroplast trnL intron in bryophytes. Mol Phylogenet Evol 2005; 36: 429–443. [48] Renner SS, Foreman DB, Murray D. Timing Transantarctic disjunctions in the Atherospermataceae (Laurales): Evidence from coding and noncoding chloroplast sequences. Syst Biol 2000; 49: 579–591. [49] Ritz CM, Zimmermann NFA, Hellwig FH. Phylogeny of subsect. Meleuphorbia (A. Berger) Pax & Hoffm. (Euphorbia L.) reflects the climatic regime in South Africa. Plant Syst Evol 2003; 241: 245–259. [50] Samuel R, Pinsker W, Kiehn M. Phylogeny of some species of Cyrtandra (Gesneriaceae) inferred from the atpB/rbcL cpDNA intergene region. Bot Acta 1997; 110: 503–510. [51] Sang T, Crawford DJ, Stuessy T. Chloroplast DNA phylogeny, reticulate evolution, and biogeography of Paeonia (Paeoniaceae). Amer J Bot 1997; 84: 1120–1136. [52] Schill DB, Long DG, Moeller M, Squirrell J. Phylogenetic relationships between Lophoziaceae and Scapaniaceae based on chloroplast sequences. Monogr Syst Bot Missouri Bot Gard 2004; 98: 141–149. [53] Schuettpelz E, Hoot SB, Samuel R, Ehrendorfer F. Multiple origins of Southern Hemisphere Anemone (Ranunculaceae) based on plastid and nuclear sequence data. Plant Syst Evol 2002; 231: 143–151. [54] Shaw AJ, Cox CJ, Goffinet B, Buck WR, Boles SB. Phylogenetic evidence of a rapid radiation of pleurocarpous mosses (Bryophyta). Evolution 2003; 57: 2226–2241. [55] Silhavy D, Maliga P. Mapping of promoters for the nucleus-encoded plastid RNA polymerase (NEP) in the jojap maize mutant. Curr Genet 1998; 33: 340–344. [56] Simmons MP, Ochoterena H. 2000. Gaps as characters in sequence-based phylogenetic analyses. Syst Biol 2000; 49: 369–381. [57] Stech M. Supraspecific circumscription and classification of Campylopus Brid. (Dicranaceae, Bryopsida) based on molecular data. Syst Bot 2004; 29: 817–824. [58] Stech M, Dohrmann J. Molecular relationships and biogeography of two Gondwanan Campylopus species, C. pilifer and C. introflexus (Dicranaceae). Monogr Syst Bot Missouri Bot Gard 2004; 98: 415–431. [59] Stech M, Frey W. CpDNA relationship and classification of the Jungermanniopsida. Nova Hedwigia 2001; 72: 45–58.

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[60] Stech M, Frey W. Molecular circumscription and relationships of selected Gondwanan species of Haplomitrium (Calobryales, Haplomitriopsida, Hepaticophytina). Studies in austral temperate rain forest bryophytes 24. Nova Hedwigia 2004; 78: 51–70. [61] Stech M, Quandt D, Frey W. Molecular circumscription of the hornworts (Anthocerotophyta) based on the chloroplast DNA trnL-trnF region. J Plant Res 2003; 116: 389–398. [62] Stech M, Quandt D, Lindlar A, Frahm J-P. The systematic position of Pulchrinodus inflatus (Pterobryaceae, Bryopsida) based on molecular data. Studies in austral temperate rain forest bryophytes 21. Australian Syst Bot 2003; 16: 561–568. [63] Stech M, Wagner D. Molecular relationships, biogeography, and evolution of Gondwanan Campylopus species (Dicranaceae, Bryopsida). Taxon 2005; 54: 377–382. [64] Stern DB, Radwanski ER, Kindle KL. A 3´ stem/loop structure of Chlamydomonas chloroplast atpB gene regulates mRNA accumulation in vivo. Plant Cell 1991; 3: 285–297. [65] Sugiura C, Kobayashi Y, Aoki S, Sugita C, Sugita M. Complete chloroplast DNA sequence of the moss Physcomitrella patens: evidence for the loss and relocation of rpoA from the chloroplast to the nucleus. Nucleic Acids Res 2003; 31: 5324–5331. [66] Swofford DL. PAUP*: Phylogenetic analysis using parsimony (*and other methods). Version 4.0b10. Sinauer, Sunderland, Massachusetts; 2002. [67] Vanderpoorten A, Goffinet B, Hedenäs L, Cox CJ, Shaw AJ. A taxonomic reassessment of the Vittiaceae (Hypnales, Bryopsida): evidence from phylogenetic analyses of combined chloroplast and nuclear sequence data. Plant Syst Evol 2003; 241: 1–12. [68] Vanderpoorten A, Hedenäs L, Cox CJ, Shaw AJ. Circumscription, classification, and taxonomy of Amblystegiaceae (Bryopsida) inferred from nuclear and chloroplast DNA sequence data and morphology. Taxon 2002; 51: 115–122. [69] Vanderpoorten A, Shaw AJ, Cox CJ. Evolution of multiple paralogous adenosine kinase genes in the moss genus Hygroamblystegium : phylogenetic implications. Mol Phylogenet Evol 2004; 31: 505–516. [70] Weihe A, Börner T. Transcription and the architecture of promoters in chloroplasts. Trends Plant Sci 1999; 4: 169–170. [71] Wolfson R, Higgins KG, Sears BB. Evidence for replication slippage in the evolution of Oenothera chloroplast DNA. Mol Biol Evol 1991; 8: 709–720.

PLANT GENOME: BIODIVERSITY AND EVOLUTION Volume 2, Part B Lower Groups

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Molecular Phylogeny and Biogeography of Plagiochila (Jungermanniidae: Plagiochilaceae) JOCHEN HEINRICHS Department of Systematic Botany, Georg August University, Göttingen, Germany

ABSTRACT The cosmopolitan genus Plagiochila (Jungermanniidae: Plagiochilaceae) with about 450 species is the largest genus of hepatics. The genus is morphologically well-defined, whereas taxonomy at species and section level is hampered by the scarcity of stable morphological characters. Traditional classification systems based on morphology often do not reflect phylogenetic relationships within Plagiochila. Recent molecular studies based on phylogenetic analyses of internal transcribed spacer sequences of nuclear ribosomal DNA and chloroplast gene r p s 4 sequences led to new insights into the phylogeny and biogeography of Plagiochila. Contrary to earlier belief, intercontinental ranges at section level are common. Molecular data also support intercontinental ranges of species. Close links exist between the Plagiochila floras of the Neotropics and Africa as well as Atlantic Europe, and between those of Australasia and Southern South America. The Asian Plagiochila flora is closely related to that of the Holarctics. Plagiochila sects. Vagae and Cucullatae have pantropical ranges. The monospecific neotropical genera Rhodoplagiochila, Steereochila, and Szweykowskia have proved to be elements of Plagiochila. Due to the frequency of morphological homoplasy, it is not possible to identify monophyletic species groups of Plagiochila by morphology alone.

Address for correspondence: Department of Systematic Botany, Albrecht von Haller Institute of Plant Sciences, Georg August University, Untere Karspüle 2, D-37073 Göttingen, Germany. E-mail: [email protected].

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Key Words: Plagiochilaceae, Plagiochila, molecular phylogeny, biogeography, ITS, r p s 4, rb c L

INTRODUCTION

Plagiochila (Dumort.) Dumort., with over 1,600 published binomials [52, 54] and an estimated number of 400-450 species [83], is the largest genus of hepatics. Species of Plagiochila occur worldwide with the exception of the Antarctic mainland. Centers of diversity, however, are the humid tropics of Asia and America [29, 52, 79]. Large numbers of species also occur in temperate regions with oceanic climate (for example, 16, 25, 49, 50, 55]. Representatives of Plagiochila are able to form large mats, and are among the most diverse and abundant cryptogams in humid tropical forests. The genus is well defined by its laterally compressed perianth with a dorsal keel usually longer than the ventral one, dioicism, succubous leaves, alternating foliation, and almost exclusively lateral branching. In contrast, taxonomy at species and section level is hampered by the scarcity of stable gametophytic characters [for example, 29, 84] and has been regarded as “daunting, forbidding, confusing and notoriously difficult” [79]. SPECIES RANGES: TRADITIONAL CONCEPTS

Many authors [10, 85] tried to cope with the huge variability of Plagiochila gametophytes by describing numerous species that were usually assigned to small ranges only. However, the application of a narrow species concept is problematic because of the presence of numerous intermediates. Modern revisions [29, 52, 56, 62, 79, 89] have thus adopted a wider species concept, resulting in binomials with larger geographical ranges and numerous synonyms. Nevertheless, intercontinental ranges are considered to be rare in Plagiochila and have been proposed for a few species only, for example, Plagiochila bifaria (Sw.) Lindenb. [36], Plagiochila exigua (Taylor) Taylor [51], Plagiochila longispina Lindenb. & Gottsche [30], Plagiochila parvifolia Lindenb. [74, 79] or Plagiochila semidecurrens (Lehm. & Lindenb.) Lindenb. [74, 79].

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SUPRASPECIFIC CLASSIFICATION OF PLAGIOCHILA BASED ON MORPHOLOGY

Several morphology-based classification systems were proposed from the mid-19th to the late 20th century [4, 8, 11, 52, 55, 56, 58, 70, 71, 79, 85, 88]. The first attempt to classify Plagiochila was published by Lindenberg [58] who recognized six sections, based on characters of the dried gametophytes including perianth and leaf shape as well as branching pattern. Subsequent attempts to define units solely based on leaf shape by Dugas [8] and Stephani [88] were not successful and soon rejected [4]. An important contribution to the taxonomy of Plagiochila was made by Spruce [85] who divided the genus into “Cauliflorae” and “Ramiflorae”, based upon branching pattern and perianth position. This subdivision was modified by later authors [55, 56]. The most comprehensive classification was proposed by Carl [4] who gave attention to Plagiochila worldwide, with the exception of Africa, and studied more than 400 binomials. Carl accepted three Plagiochila subgenera; subgenus Oppositae Carl (= Plagiochilion S.Hatt. [26]), the monospecific subgenus Cuculifolliae Carl (= Szweykowskia Gradstein & Reiner-Drehwald [12]), and subgenus Eu-Plagiochila, with the latter subgenus agreeing more or less with the current concept of Plagiochila. In a first step, Carl [4] divided the species of subgenus Eu-Plagiochila based on their distribution in three floristic kingdoms (Neotropics, Paleotropics, Austral-Antarctic; Africa not treated). In a second step, he defined 50 sections which were based on several characters of the dried gametophytes, including leaf cell pattern, density of foliation, branching, vegetative reproduction, position of the androecia and gametangia, as well as dentation of the male bracts. These sections were usually restricted to a single floristic kingdom or region. Some authors improved Carl’s concepts by the introduction of new morphological characters. Their works [32, 33, 52, 71] usually affected the separation or summarization of supraspecific taxa recognized by Carl [4] or the exclusion of particular groups from Plagiochila, for example, Acrochila R.M.Schust. or Xenochila R.M.Schust. Inoue and Schuster [55], Inoue [52] and Heinrichs et al. [30] demonstrated the value of oil body characters for the taxonomy of Plagiochila and used both gametophyte and sporophyte characters. Substantial changes of Carl’s concepts primarily arose from the work of Inoue [52] who raised two sections to subgenera (Plagiochila sect. Peculiares Schiffn. ® P. subgen.

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Paraplagiochila Inoue, P. sect. Cucullatae Schiffn. ® P. subgen. Metaplagiochila Inoue), lowered Chiastocaulon Carl to a subgenus of Plagiochila and established the subgenus Pleurochila Inoue for a single New Zealand species, P. pleurata (Hook. f. & Taylor) Gottsche, Lindenberg & Nees. Five subgenera, including the type subgenus, arose from this study, of which one (the subgenus Plagiochila) includes the vast majority of species. Inoue’s hierarchical subdivision of Plagiochila into subgenera and sections was accepted by some recent authors [for example, 18, 62, 79, 84]. IMPLICATIONS FROM MOLECULAR PHYLOGENIES Molecular Markers

Plagiochilaceae are the subject of ongoing molecular studies. Up to now, sequences of about 100 species of Plagiochila, i.e. of a quarter of the genus have been included in phylogenetic analyses. Molecular investigations within Plagiochila have so far been based on phylogenetic analyses of the internal transcribed spacer region of nuclear ribosomal DNA (nrITS15.8S-ITS2) [19, 22, 24, 29, 35, 38, 40, 42, 43, 44, 45, 59, 64, 65, 66, 76], analyses of nrITS and chloroplast gene rps4 sequences [20, 23, 41], as well as of chloroplast gene rbcL sequences [21]. Several authors included single Plagiochila species in phylogenetic analyses of larger groups of liverworts [2, 3, 6, 7, 27, 28, 63, 68, 69, 86, 87, 90]. Accordingly, numerous nrITS and cp rps4 sequences, a few rbcL sequences and single sequences of the chloroplast psbA gene, the trnL-F and the ITS2-4.5SITS3 regions, the mitochondrial nad4-2 spacer region, the nad1 and nad5 gene, as well as nuclear encoded 18S and 26S ribosomal DNA genes are available from GenBank. Separate analyses of chloroplast gene rps4 and nrITS sequences of Plagiochila led to largely similar topologies [20, 23, 41], indicating that chloroplast and nucleus share the same history. The nrITS region often shows infraspecific variation [39, 64]. nrITS topologies are usually well supported at crown groups but lack support at many deeper nodes [24, 29, 44]. The most variable regions of Plagiochila ITS sequences are usually difficult to align, leading to exclusion of extensive parts of the sequences in datasets which cover larger parts of the morphological variation of the genus [23, 41]. Within closely related species groups, however, practically the entire nrITS region can be aligned unambiguously [41]. The chloroplast gene rps4 is more conservative than

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the nrITS region and usually not informative at species level, whereas deeper nodes are often well resolved [20, 23]. Combination of rps4 and ITS leads to robust topologies both at deeper nodes and crown groups [20, 23, 41]. The rbcL gene allows the discrimination of the main clades of Plagiochila [21]. CURRENT HIERARCHICAL SUBDIVISIONS OF PLAGIOCHILA ARE NOT REFLECTED IN MOLECULAR TOPOLOGIES

To date, representatives of four of the five subgenera of Plagiochila accepted by Inoue [52] have been included in molecular investigations. In phylogenetic analyses of nrITS and cp rps4 sequence alignments, Plagiochila subgen. Chiastocaulon (Carl) Inoue clustered with Pedinophyllum interruptum (Nees) Kaal. and Plagiochilion mayebarae S.Hatt. (Plagiochilaceae) rather than with Plagiochila, leading to the reinstatement of the genus Chiastocaulon [20]. Morphologically, Chiastocaulon differs from Plagiochila by the frequent occurrence of ventral intercalary branches [5, 20]. A robust sister relationship of Chiastocaulon and Plagiochilion was also confirmed in phylogenetic analyses of rbcL sequences; the Chiastocaulon-Plagiochilion clade was placed sister to Plagiochila in a robust sister relationship [21]. The type species of the Plagiochila subgenera Metaplagiochila and Paraplagiochila, P. sandei Sande Lac. and P. peculiaris Schiffn. were resolved as members of the robust main clade B of Plagiochila sensu Groth & Heinrichs [20] which also includes the generitype, P. asplenioides (L.) Dumort. Consequently, both subgenera were lowered to synynoms of subgenus Plagiochila [23]. The robust main clades identified so far in Plagiochila (Fig. 1) lack morphological synapomorphisms. Hence, Groth et al. [23] abstained from a hierarchical subdivision of Plagiochila and applied only to sections, representing the most widely used rank for species groups below Plagiochila. NATURAL SPECIES GROUPS SPAN SEVERAL FLORISTIC KINGDOMS

Schuster [72] criticized Carl’s division of Plagiochila [4] based on geographical criteria and the often very small sections, which can be monospecific after elimination of the synonyms [61]. Molecular phylogenetic studies corroborate Schuster’s objections: natural species

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outgroup

Pedinophyllum interruptum Chiastocaulon dendroides [20] Plagiochilion mayebarae E, N [20, 23, 24, 29, 30, 48]

D Glaucescentes

Durae Au, S [4, 23, 55]

including Rhodoplagiochila Arrectae Af, E, N, W [17, 22-24, 29, 34, 36, 39, 41, 43, 44, 59, 65, 67, 75, 76] P. rubesce ns S

C

Rutilantes

Vagae

A, Af, E, N, P, S, W [19, 22, 23, 29, 37, 38, 41, 51, 66]

A, Au, Af, E, N, W, P [23, 29, 33, 40-42, 45, 47, 52, 56, 62, 77, 80]

Africanae Af [41, 56] A, W [20, 23, 42]

Trabeculatae

Plagiochila sects.

Fruticosae Plagiochila

B

Poltiae

Adiantoideae

A

A, Au [20, 23] A, E, W [20, 23, 29, 61, 64, 79]

A tropical and temperate Asia Au Australasia (extended to New Zealand) Af Africa E Eastern Holarctic N Neotropics P Pacific S Southern South America W Western Holarctic

A, Au, E [13, 23, 62, 64, 79] A, Au, P, W {P. subgen. Paraplagiochila} Peculiares [23, 24, 29, 41, 42, 52, 62, 78] {P. subgen. Metaplagiochila} Cucullatae A, Au, Af, N, P, W [23, 24, 29, 41, 42, 52, 62, 78] N [23, 29] Fusco- N, E [23, 24, 29, 32, 46, 57] luteae including Steereochila Hylacoetes Szweykowskia N, A [12, 23, 24, 29, 35, 41, 53] Alternantes Au, S, N [23, 29, 55, 61]

Fig. 1. Maximum likelihood phylogeny of Plagiochila based on a combined nrITS / cp rps4 sequence alignment. Sectional clades all with good bootstrap support, robust basal clades double bold (Simplified from [41]; distribution of sections indicated). For details on distribution, morphology, species assemblages, and synonymy of the sectional clades, see the literature indicated by bracketed reference numbers.

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groups (sections) often span several floristic kingdoms [22, 23, 24, 29, 41, see also Fig. 1]. Examples of intercontinental species groups include the originally neotropical sect. Alternantes Carl [4] which is nested in a clade with species from Southern South America and New Zealand [23] [Fig. 2], the originally Asian sect. Cucullatae Schiffn. [70, 71] which is now recognized as pantropical [23, 24], and the originally neotropical section Arrectae Carl [4] with representatives in Atlantic Europe, tropical Africa, and North America [29, 39, 40, 43, 59, 65, 67, 76]. Current knowledge on the distribution of Plagiochila sections so far included in molecular studies is summarized in Fig. 1. Molecular surveys also indicate that the majority of proposed supraspecific taxa belong in the synonymy. Possibly not more than 25% of the more than 100 supraspecific taxa will be reflected in molecular topologies [23]. A good example concerning this matter is Plagiochila sect. Vagae Lindenb. As an outcome of molecular phylogenetic studies [23, 41] Plagiochila sect. Vagae represents a robust pantropical clade (Fig. 3) with numerous representatives in all parts of the tropics. Morphologically the clade can be identified by the frequent terminal branching of its representatives, their colorless oil bodies, a capsule wall with cell wall thickenings in all layers, and vegetative reproduction by propagules on the surface of the leaves, or occasionally by fragmenting leaves [23, 47]. Numerous sections were resolved as representatives of the Vagae clade including P. sects. Contiguae Carl, Crispatae Carl, Hypnoides Carl, Parallelae Carl, and Subtropicae Carl [23, 29, 41]. Although defined at most by weak morphological tendencies in density of foliation or leaf shape, these sections were widely accepted in recent literature [52, 74, 79]. The similarity of these groups caused problems with the assignment of several species that have been moved between some of the above sections [81]. However, the new morphological circumscription derived from molecular topologies allows the certain identification of representatives of the lineage. Molecular data also support intercontinental ranges at species level. Disjunct ranges have been verified in several studies of nrITS sequence variation (Plagiochila bifaria, Neotropics-Europe [39]; P. boryana Steph., Neotropics-Africa, [41]; P. carringtonii (Balf.) Grolle, Himalaya-Europe [64]; P. corrugata (Nees) Mont. & Nees, Neotropics-Africa, [40]; P. punctata (Taylor) Taylor, Neotropics, Europe, Africa [43]; Fig. 4, P. sciophila Lindenb., Asia-North America [42]).

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MOLECULAR TOPOLOGIES AND MORPHOLOGY: THE CASE STUDY PLAGIOCHILA SECT. SUPERBAE

Carl [4] set up an illegitimate Plagiochila sect. Superbae Carl (Art. 52.1 ICBN) to include tropical American species with toothed, ampliate leaves consisting of large cells, imbricate foliation, terminal, fan-shaped, rarely simple, intercalary androecia and lack of well-developed underleaves. To find a name for the illegitimate Superbae, Heinrichs [29] carried out a combined molecular-morphological study. Neotropical species matching Carl’s description were placed in a robust main clade of Plagiochila (Fig. 1, clade A) which was further subdivided into three clades assignable to Plagiochila sects. Adiantoideae Lindenb., Fuscoluteae Carl and Hylacoetes Carl. Not only “Superbae” sensu Carl were identified as members of this main clade, but also species without ampliate leaves or with copious underleaves. Renewed morphological analyses of members of “main clade A” led to a new circumscription of the related sections Adiantoideae, Fuscoluteae and Hylacoetes (Fig. 5) and to relegation of numerous supraspecific taxa to synonymy including the monospecific genera Steereochila Inoue and Szweykowskia Gradstein & Reiner-Drehwald which were separated from Plagiochila because of a peculiar type of vegetative reproduction (Steereochila [53]) or saccate leaves (Szweykowskia [12]). Within “main clade A”, Hylacoetes (Fig. 6) stand out by their sporophyte with a capsule wall without cell wall thickenings in the epidermal layer [29]. Most Hylacoetes species have terminal, fan shaped androecia with opposite bracts not overlapping dorsally. A few of its representatives build simple androecia and male bracts which overlap dorsally (P. guevarii H.Rob., P. cucullifolia Jack & Steph. [29, 35]). The latter type of androecia is typical both for Adiantoideae and Fuscoluteae (Fig. 7) which have a capsule with thickenings in all cell layers. However, Fuscoluteae can be separated from Adiantoideae by the presence of surface wax [29, 31, 46]. The above study demonstrates that morphological characters used in traditional classification systems of Plagiochila (for example, leaf shape and dentation) are unemployable for a subdivision of Plagiochila into monophyletic units. Characters of the sporophyte are more promising with regard to a natural classification of Plagiochila [29, 30]; however, they are usually also homoplastic at section level [for example, 23, 29, see p. 447].

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MORPHOLOGICAL DATASETS OF PLAGIOCHILA HAVE A HIGH CONTENT OF HOMOPLASY

Although many sectional clades of Plagiochila can be characterized by a couple of morphological characters, it is evident from molecular topologies that morphological datasets of Plagiochila have a high content of homoplasy [20, 23, 29]. All members of the neotropical-African sect. Hylacoetes have a capsule without cell wall thickenings in the epidermal layer (referred above). This type of sporophyte recurs in the pantropical sect. Cucullifoliae. Both groups are placed in different main clades of Plagiochila [20, 23, 29, 41, Figs. 1, 5]. Müller et al. [61] demonstrated that the originally Neotropical Plagiochila sect. Alternantes shares most of its sporophytic features with the holarctic sect. Plagiochila (Fig. 8) and proposed a synonymy of both names. Representatives of both groups form a robust monophyletic lineage in phylogenetic analyses of a morphological matrix of numerous species of Plagiochila [29]. In contrast, molecular topologies (Fig. 1) clearly indicate that both groups are not closely related and should be kept separate. Dominating terminal branches, serving as a basis for the subdivision of Plagiochila into Cauliflorae and Ramiflorae [85] occur in different lineages of Plagiochila, for example, Glaucescentes and Vagae [30, 45]. Many other characters are diffusely distributed and occur in single representatives of different sections of Plagiochila. Paraphyllia are, for instance, known from some representatives of the sects. Fruticosae Inoue, Fuscoluteae, Peculiares Schiffn., and Vagae [23]. Unispiral elaters, once considered to occur only rarely in Plagiochila [56], have been detected in some representatives of Arrectae [17, 36, 65], Durae Carl [23], Rutilantes Carl [22, 23], and Vagae [40, 42, 56], i.e. in some representatives of main clades B, C, and D (Fig. 1). Other representatives of the above sections, respectively main clades, have bispiral elaters [22, 23, 29, 67]. Due to the frequency of homoplasy it is not possible to identify monophyletic species groups of Plagiochila by morphology alone [23]. It seems that the “morphotype Plagiochila” is consistently modified and that certain character combinations recur regularly. However, morphologically similar sectional clades are often geographically separated [23, 29].

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Fig. 8. Morphology of Plagiochila (sect. Plagiochila) porelloides (Nees) Lindenb. (Germany, specimen Heinrichs & Groth 4043): A, top of shoot with sporophyte; B, leaf cells with oil bodies; C, cross section of capsule wall; D, innermost layer of capsule wall, surface view; E, epidermal layer of capsule wall, surface view; F, G, inner layers of capsule wall, surface view.

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TAXONOMIC DECISIONS BASED ON MOLECULAR TOPOLOGIES: PLAGIOCHILA DETECTA, PLAGIOCHILA MADERENSIS, PLAGIOCHILA RUTILANS AND RHODOPLAGIOCHILA

Molecular data may help to evaluate the taxonomic value of certain morphological characters. So & Grolle [82] examined large sized East Asian Plagiochila morphotypes with unispiral elaters which otherwise matched the features of P. trabeculata Steph. Using RAPD analysis, they were able to clearly discriminate the large sized morphotype against typical P. trabeculata. As a consequence of their RAPD investigation and the morphological differences, the authors established a new species, Plagiochila detecta So & Grolle, hitherto treated as fo. stipulata Inoue of P. trabeculata [50]. Schuster [73] established Rhodoplagiochila R.M.Schust. as a new, monospecific genus of Plagiochilaceae because of the rose-red pigmentation of its shoot apices. A few years later, Inoue [52] transferred the genus to Lophoziaceae. However, Schuster [75] depicted the plagioichilid perianths of Rhodoplagiochila and provided convincing evidence that the genus should remain within Plagiochilaceae. Heinrichs [29] doubted the generic status of Rhodoplagiochila and assumed the taxon to be a member of Plagiochila sect. Arrectae. An isotype, as well as specimens from nearby the type locality in the Andes of Venezuela, were remarkably similar to small sized forms of Plagiochila (sect. Arrectae) bifaria. However, the plants differed from P. bifaria by their papillose leaf surface and the leaf margins toothed all-around [44]. Judging from the morphology, Rhodoplagiochila rosea could represent both a species of P. sect. Arrectae or an extreme morphotype of the polymorphic P. bifaria. Phylogenetic analyses of nrITS sequences revealed Rhodoplagiochila in a robust clade together with several accessions of P. bifaria. As an outcome of the molecular investigation, Rhodoplagiochila was treated as a variety of P. bifaria [44] rather than a distinct species (also see Fig. 5). Gametophyte morphology supported the reinstatement of the Madeiran endemic Plagiochila maderensis Steph. [66], which had been treated as a synonym of P. (sect. Arrectae) spinulosa (Dicks.) Dumort. by Grolle [14] and subsequent authors. Sectional placement of P. maderensis was impossible by morphology alone. Maximum likelihood analyses of nrITS sequences resolved P. maderensis as a member of P. sect. Rutilantes, as was also indicated by lypophylic secondary metabolites [66].

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Lipophilic secondary metabolites alongside nrITS sequence topologies allowed the clarification of the systematic position of several Plagiochila binomina which are morphologically characterized by a “free” perianth (Plagiochila deflexa Mont. & Gottsche, P. moritziana Hampe, P. permista Spruce [38]) but were placed in several different sections of Plagiochila by Carl [4]. Anton [1] defined a “Chemogruppe IX” of Plagiochila being characterized by the presence of various phenolic compounds and lack of flavonoids and plagiochilines. “Chemogruppe IX” included the above species in addition to taxa having a perianth covered by bracts. In molecular analyses of chloroplast rps4 and of nrITS sequences, however, the species with free perianth were resolved as members of P. sect. Rutilantes, the species with a perianth covered by bracts as members of P. sect. Arrectae (Fig. 1, main clade C). Plagiochila moritziana, as well as several other Rutilantes, stand out by a distinct odor of peppermint caused by highly volatile monoterpenoids [22, 23, 37, 38]. Plagiochila moritziana differs from typical P. rutilans Lindenb. mostly in leaf shape. Forming a monophyletic lineage with P. rutilans, it was reduced to a variety of the latter [38]. PHYLOGEOGRAPHY

The oldest known fossil of Plagiochila, P. groehnii Grolle & Heinrichs, dates back to the Eocene [15]. Currently it is impossible to infer the diversification time of Plagiochila based on the poor fossil record within the genus and the lack of a stable liverwort phylogeny [6, 9, 27]. Despite this constraint, several tentative conclusions have been drawn with respect to the biogeography of Plagiochila [23, 24, 39-44]. The apparent lack of Australasian-southern South American sections of Plagiochila in Africa and the relationships of the African and Asian Plagiochila floras (Fig. 1, main clade B) was taken as an argument for a diversification of at least major parts of the genus a long time after the breakup of Gondwana at about 180 million years ago [60]. The results of several studies point to a center of phylogenetic diversity of Plagiochilaceae in Southeast Asia and Australasia [20, 23, 29, 41, 44]. In these regions Plagiochilaceae are represented by several genera, whereas in Africa and tropical America the family is limited to Plagiochila [56]. In the Holarctic, Plagiochilaceae are represented by Plagiochila and Pedinophyllum (Lindb.) Lindb. [20], in Southern South America by Plagiochila and Pedinophyllopsis R.M.Schust. & Inoue [28, 29]. The Alternantes clade of

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Plagiochila is of Gondwanan range (Fig. 1). In combination with the phylogenetic diversity described above, this was taken as an argument for a possible origin of major parts of extant Plagiochila from ancestors in Australasia and adjacent regions [23]. Molecular data clearly demonstrate that the Plagiochila binomials from Atlantic Europe belong to neotropical or pantropical sections, or to sections from Asia [23, 24, 41, 64]. Contrary to earlier belief [48, 57], most of these taxa do not represent endemics of Europe but belong to far more widespread species. Close links exist between the Plagiochila floras of atlantic Europe and tropical America [overviews in 24, 66]. The Plagiochila flora of the western Holarctic has links to the Neotropics as well as Asia and Europe [23, 42]. Heinrichs et al. [35, 43, 44] presumed that late Miocene uplift of the Andes led to a rapid speciation within some neotropical species groups of Plagiochila. They also proposed that P. sect. Arrectae originated from southern South American ancestors, diversified in tropical America after the uplift of the Andes and reached the Holarctic and Africa by long distance dispersal [43]. Long distance dispersal of spores was favored as an explanation for intercontinental ranges of several Plagiochila species with similar nrITS sequences throughout their range [24, 39, 41]. Molecular topologies indicate that Plagiochila boryana reached tropical Africa by long distance dispersal originating from the Neotropics [41]. Heinrichs et al. [41] assumed that the low diversity of Plagiochila in Africa may reflect extensive drought periods over the Pleistocene and that the extant tropical African Plagiochila flora is a mixture of old elements and rather recent immigrants. Intercontinental ranges at species level exist between tropical America and Africa, whereas similarities between tropical Africa and Asia were only recovered at section level. Topologies within the pantropical Vagae clade support the hypothesis of several switches from Africa to Asia and vice versa, as well as an African origin of the group [41]. Molecular data seem to indicate that Plagiochila is able to react to climatic changes relatively fast. Many species seem to be able to spread out by long distance dispersal of diaspores indicating that ancient vicariance is not a standard model for bryophyte disjunctions.

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Acknowledgments

I thank Hermann Anton, Kathrin Feldberg, S. Rob Gradstein, Henk Groth, Florian Hartmann, Melanie Lindner, Jochen Müller, Rüdiger Mues, Thomas Pröschold, Carsten Renker, David Rycroft, Alfons Schäfer-Verwimp, Harald Schneider, and Rosemary Wilson for fruitful collaboration. Work on taxonomy and phylogeny of Plagiochilaceae in Göttingen has been supported by the Deutsche Forschungsgemeinschaft (grants DFG Gr 1588/1, He 3584/1). References [1] Anton H. Neue Ergebnisse zur Chemie und Chemotaxonomie der Plagiochilaceae (Lebermoose). Thesis, University of Saarbrücken, Saarbrucken, Germany, 2001. [2] Beckert S, Steinhauser S, Muhle H, Knoop V. A molecular phylogeny of bryophytes based on nucleotide sequences of the mitochondrial nad5 gene. Pl Syst Evol 1999; 218: 179–192. [3] Bopp M, Capesius C. New aspects of bryophyte taxonomy provided by a molecular approach. Bot Acta 1996; 109: 368–372. [4] Carl H. Die Arttypen und die systematische Gliederung der Gattung Plagiochila Dum. Ann Bryol Suppl 1931; 2: 1–170. [5] Carl H. Morphologische Studien an Chiastocaulon Carl, einer neuen Lebermoosgattung. Flora 1931; 126: 45–60. [6] Davis EC. A molecular phylogeny of leafy liverworts (Jungermanniidae: Marchantiophyta). Monogr Syst Bot Missouri Bot Gard 2004; 98: 61–86. [7] Dombrovska O, Qiu YL. Distribution of introns in the mitochondrial gene nad1 in land plants: phylogenetic and molecular evolutionary implications. Mol Phyl Evol 2004; 32: 246–263. [8] Dugas M. Contribution a l’étude du genre “Plagiochila” Dum. Ann Sci Nat Bot 1929; Sér 10 11: 1–199. [9] Forrest LL, Crandall-Stotler BJ. A phylogeny of simple thalloid liverworts (Jungermanniopsida, Metzgeriidae) as inferred from five chloroplast genes. Monogr Syst Bot Missouri Bot Gard 2004; 98: 119–140. [10] Gottsche ACM. De mexikanske Levermosser. Beskrevne efter Prof. Fr. Liebmanns Samling. Copenhagen: Bianco Luno, 1863–1867. [11] Gottsche ACM, Lindenberg JBG, Nees ab Esenbeck CG. Synopsis hepaticarum. Hamburg: Meissner, 1844–1847. [12] Gradstein SR, Reiner-Drehwald ME. Szweykowskia, a new genus of Plagiochilaceae (Hepaticae) from tropical America. Fragm Florist Geobot 1995; 40: 31–38. [13] Grolle R. Jamesoniella carringtonii - eine Plagiochila in Nepal mit Perianth. Trans Brit Bryol Soc 1964; 4: 653–663. [14] Grolle R. Miscellanea Hepaticologica (71-80). Trans Brit Bryol Soc 1967; 5: 271–282.

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[15] Grolle R, Heinrichs J. Eocene Plagiochila groehnii sp. nov. - the first representative of Plagiochilaceae in Baltic Amber. Cryptogamie Bryol 2003; 24: 289–293. [16] Grolle R, Long DG. An annotated check-list of the Hepaticae and Anthocerotae of Europe and Macaronesia. J Bryol 2000; 22: 103–140. [17] Grolle R, Schumacker R. Zur Synonymik und Verbreitung von Plagiochila spinulosa (Dicks.) Dum. und P. killarniensis Pears. J Bryol 1982; 12: 215–225. [18] Grolle R, So ML. Notes on Plagiochila subgenus Paraplagiochila (Hepaticae). J Bryol 1999; 21: 197–199. [19] Groth H, Hartmann FA, Wilson R, Heinrichs J. nrITS sequences and morphology indicate a synonymy of the Patagonian Plagiochila rufescens Steph. and the Central American Plagiochila bicuspidata Gottsche. Cryptogamie Bryol 2004; 25: 19–28. [20] Groth H, Heinrichs J. Reinstatement of Chiastocaulon Carl (Plagiochilaceae), based on evidence from nuclear ribosomal ITS and chloroplast gene rps4 sequences. Pl Biol 2003; 5: 616–622. [21] Groth H, Heinrichs J. Maximum likelihood analyses of chloroplast gene rbcL sequences indicate relationships of Syzygiella (Jungermanniopsida) with Lophoziaceae rather than Plagiochilaceae. Cryptogamie, Bryol. 2005; 26: 49–57. [22] Groth H, Helms G, Heinrichs J. The systematic status of Plagiochila sect. Bidentes Carl and Caducilobae Inoue (Hepaticae) inferred from nrDNA ITS sequences. Taxon 2002; 51: 675–684. [23] Groth H, Lindner M, Heinrichs J. Phylogeny and biogeography of Plagiochila (Plagiochilaceae) based on nuclear and chloroplast DNA sequences. Monogr Syst Bot Missouri Bot Gard 2004; 98: 365–387. [24] Groth H, Lindner M, Wilson R, Hartmann FA, Schmull M, Gradstein SR, Heinrichs J. Biogeography of Plagiochila (Hepaticae): natural species groups span several floristic kingdoms. J Biogeogr. 2003; 30: 965–978. [25] Hässel de Menendez GG. Informaciones nomenclaturales sobre las especies del genero Plagiochila (Hepaticae) de Argentina y Chile. Bol Soc Argent Bot 1983; 22: 87–129. [26] Hattori S. Five new genera of Hepaticae. Biosphaera 1947; 1: 3–7. [27] He-Nygrén X, Ahonen I, Juslén A, Glenny D, Piippo S. Phylogeny of liverworts beyond a leaf and a thallus. Monogr Syst Bot Missouri Bot Gard 2004; 98: 87–118. [28] He-Nygrén X, Piippo S. Phylogenetic relationships of the generic complex Chiloscyphus-Lophocolea-Heteroscyphus (Geocalycaceae, Hepaticae): Insights from three chloroplast genes and morphology. Ann Bot Fenn 2003; 40: 317–329. [29] Heinrichs J. A taxonomic revision of Plagiochila sect. Hylacoetes, sect. Adiantoideae and sect. Fuscoluteae in the Neotropics with a preliminary subdivision of Neotropical Plagiochilaceae into nine lineages. Bryophyt Biblioth 2002; 58: 1–184, Append. 1–5. [30] Heinrichs J, Anton H, Gradstein SR, Mues R. Systematics of Plagiochila sect. Glaucescentes Carl (Hepaticae): a morphological and chemotaxonomical approach. Pl Syst Evol 2000; 220: 115–138. [31] Heinrichs J, Anton H, Gradstein SR, Mues R, Holz I. Surface wax, a new taxonomic feature in Plagiochilaceae. Pl Syst Evol 2000; 225: 225–233. [32] Heinrichs J, Gradstein SR. On Plagiochila longiramea Steph. (Hepaticae), a poorly known species of Bolivia. Candollea 1999; 54: 73–81.

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[33] Heinrichs J, Gradstein SR. A revision of Plagiochila sect. Crispatae and sect. Hypnoides (Hepaticae) in the Neotropics. I. Plagiochila disticha, P. montagnei and P. raddiana. Nova Hedwigia 2000; 70: 161–184. [34] Heinrichs J, Gradstein SR, Grolle R. A revision of the Neotropical species of Plagiochila (Dumort.) Dumort. (Hepaticae) described by Olof Swartz. J Hattori Bot Lab 1998; 85: 1–32. [35] Heinrichs J, Gradstein SR, Groth H, Lindner M. Plagiochila cucullifolia var. anomala var. nov. from Ecuador, with notes on discordant molecular and morphological variation in Plagiochila. Pl Syst Evol 2003; 242: 205–216. [36] Heinrichs J, Grolle R, Drehwald U. The conspecifity of Plagiochila killarniensis Pearson and P. bifaria (Sw.) Lindenb. (Hepaticae). J Bryol 1998; 20: 495–497. [37] Heinrichs J, Groth H, Gradstein SR, Rycroft DS, Cole WJ, Anton H. Plagiochila rutilans (Hepaticae): a poorly known species from tropical America. Bryologist 2001; 104: 350–361. [38] Heinrichs J, Groth H, Holz I, Rycroft DS, Renker C, Pröschold T. The systematic position of Plagiochila moritziana, P. trichostoma, and P. deflexa based on ITS sequence variation of nuclear ribosomal DNA, morphology, and lipophilic secondary metabolites. Bryologist 2002; 105: 189–203. [39] Heinrichs J, Groth H, Lindner M, Feldberg K, Rycroft DS. Molecular, morphological and phytochemical evidence for a broad species concept of Plagiochila bifaria (Sw.) Lindenb. (Hepaticae). Bryologist 2004; 107: 28–40. [40] Heinrichs J, Groth H, Lindner M, Renker C, Pócs T, Pröschold T. Intercontinental distribution of Plagiochila corrugata (Plagiochilaceae, Hepaticae) inferred from nrDNA ITS sequences and morphology. Bot J Linnean Soc 2004; 146: 469–481. [41] Heinrichs J, Lindner M, Gradstein SR, Groth H, Buchbender V, Solga A, Fischer E. Origin and subdivision of Plagiochila (Jungermanniidae: Plagiochilaceae) in tropical Africa based on evidence from nuclear and chloroplast DNA sequences and morphology. Taxon 2005; 54: 317–333. [42] Heinrichs J, Lindner M, Groth H. Sectional classification of the North American Plagiochila (Hepaticae, Plagiochilaceae). Bryologist 2004; 107 (4). [43] Heinrichs J, Lindner M, Groth H, Renker C. Distribution and synonymy of Plagiochila punctata (Taylor) Taylor, with hypotheses on the evolutionary history of Plagiochila sect. Arrectae (Plagiochilaceae, Hepaticae). Pl Syst Evol 2005; 250: 105–117. [44] Heinrichs J, Lindner M, Pócs T. nrDNA internal transcribed spacer data reveal that Rhodoplagiochila R.M.Schust. (Jungermanniales, Marchantiophyta) is a member of Plagiochila sect. Arrectae Carl. Org Divers Evol 2004; 4: 109–118. [45] Heinrichs J, Pröschold T, Renker C, Groth H, Rycroft DS. Plagiochila virginica A.Evans rather than P. dubia Lindenb. & Gottsche occurs in Macaronesia; placement in sect. Contiguae is supported by ITS sequences of nuclear ribosomal DNA. Pl Syst Evol 2002; 230: 221–230. [46] Heinrichs J, Rycroft DS. Leaf surface waxes and lipophilic secondary metabolites place the endemic European liverwort Plagiochila atlantica F.Rose in the Neotropical Plagiochila sect. Bursatae Carl. Cryptogamie, Bryol 2001; 22: 95–103. [47] Heinrichs J, Sauer M, Grolle R. Lectotypification and synonymy of Plagiochila sect. Vagae Lindenb. (Hepaticae). Cryptogamie, Bryol 2002; 23: 5–9.

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[48] Herzog T. Eine neue Plagiochila-Art auf den Azoren. Rev Bryol Lichénol 1945; 14: 161–162. [49] Inoue H. The family Plagiochilaceae of Japan and Formosa. I. J Hattori Bot Lab 1958; 19: 25–59. [50] Inoue H. The family Plagiochilaceae of Japan and Formosa. II. J Hattori Bot Lab 1958; 20: 54–106. [51] Inoue H. Notes on the Plagiochilaceae, X. Plagiochila corniculata (Dum.) Dum. and its allies. Bull Natn. Sci Mus, Ser B (Bot) 1980; 6: 115–124. [52] Inoue H. The genus Plagiochila in Southeast Asia. Tokyo: Academic Scientific Books, 1984. [53] Inoue H. Steereochila, a new genus of the Plagiochilaceae from the Neotropics. Mem New York Bot Gard 1987; 45: 279–282. [54] Inoue H. Plagiochila. In: Geissler P, Bischler H, eds. Naiadea to Pycnoscenus. Index Hepaticarum. Vol. 11. Berlin, Stuttgart: Cramer, 1989: 106–273. [55] Inoue H, Schuster RM. A monograph of New Zealand and Tasmanian Plagiochilaceae. J Hattori Bot Lab 1971; 34: 1–225. [56] Jones EW. African Hepatics. XV. Plagiochila in tropical Africa. Trans Brit Bryol Soc 1962; 4: 254–325. [57] Jones EW, Rose F. Plagiochila atlantica F. Rose spec. nov. - P. ambagiosa auct. J Bryol 1975; 8: 417–422. [58] Lindenberg JBG. Monographia Hepaticarum Generis Plagiochilae. Bonn, Germany: Henry & Cohen, 1839–1844. [59] Lindner M, Pócs T, Heinrichs J. On the occurrence of Plagiochila stricta on Madagascar, new to Africa. J Hattori Bot Lab 2004; 96: 261–271. [60] McLoughlin S. The breakup history of Gondwana and its impact on pre-Cenozoic floristic provincialism. Aust J Bot 2001; 49: 271–300. [61] Müller J, Heinrichs J, Gradstein SR. A revision of Plagiochila sect. Plagiochila in the Neotropics. Bryologist 1999; 102: 729–746. [62] Piippo S. Bryophyte flora of the Huon Peninsula, Papua New Guinea. XXX. Plagiochilaceae (Hepaticae). Ann Bot Fennici 1989; 26: 183–236. [63] Quandt D, Müller K, Stech M, Frahm JP, Frey W, Hilu KW, Borsch T. Molecular evulution of the chloroplast trnL-F region in land plants. Monogr Syst Bot Missouri Bot Gard 2004; 98: 13–36. [64] Renker C, Heinrichs J, Pröschold T, Groth H, Holz I. ITS sequences of nuclear ribosomal DNA support the generic placement and the disjunct range of Plagiochila (Adelanthus) carringtonii. Cryptogamie Bryol 2002; 23: 23–29. [65] Rycroft DS, Cole WJ, Heinrichs J, Groth H, Renker C, Pröschold T. Phytochemical, morphological and molecular evidence for the occurrence of the neotropical liverwort Plagiochila stricta in the Canary Islands, new to Macaronesia. Bryologist 2002; 105: 363–372. [66] Rycroft DS, Groth H, Heinrichs J. Reinstatement of Plagiochila maderensis (Jungermanniopsida: Plagiochilaceae) based on chemical evidence and nrDNA ITS sequences. J. Bryol. 2004; 26: 37–45. [67] Ryroft DS, Heinrichs J, Cole WJ, Anton H. A phytochemical and morphological study of the liverwort Plagiochila retrorsa Gottsche, new to Europe. J Bryol 2001; 23: 23–34.

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[68] Samigullin T, Yacentyuk SP, Degtyaryeva GV, Valiehoroman KM, Bobrova VK, Capesius I, Martin WF, Troitsky AV, Filin VR, Antonov AS. Paraphyly of bryophytes and close relationship of hornworts and vascular plants inferred from analysis of chloroplast rDNA ITS (cpITS) sequences. Arctoa 2002; 11: 31–43. [69] Samigullin TH, Valiejo-Roman K, Troitsky AV, Bobrova VK, Filin VR, Martin W, Antonov AS. Sequences of rDNA internal transcribed spacers from the chloroplast DNA of 26 bryophytes: properties and phylogenetic utility. FEBS Letters 1998; 422: 47–51. [70] Schiffner V. Die Hepaticae der Flora von Buitenzorg. Leiden: Brill, 1900. [71] Schiffner V. Expositio plantarum in itinere Suo Indico annis 1893/94 suscepto collectarum, II. Denkschr. Math.-Nat. Cl. K. Akad. Wiss. Wien 1900; 70: 155–218. [72] Schuster RM. A monograph of the nearctic Plagiochilaceae. Part I. Introduction and sectio I. Asplenioides. Amer. Midl. Naturalist 1959; 62: 1–166. [73] Schuster RM. Studies on Venezuelan Hepaticae. Phytologia 1978; 39: 239–251. [74] Schuster RM. The Hepaticae and Anthocerotae of North America. Vol. 4. New York: Columbia University Press, 1980. [75] Schuster RM. On the genus Rhodoplagiochila Schust. (Plagiochilaceae). Nova Hedwigia 2000; 71: 395–403. [76] Sim-Sim M, Esquível MG, Fontinha S, Cavalho S. Plagiochila stricta Lindenb. new to Madeira. Morphological and molecular evidence. Nova Hedwigia 2004; 79: 497–505. [77] So ML. Plagiochila sect. Contiguae (Hepaticae) in Australasia and the Pacific, with description of Plagiochila subjavanica sp nov. Austral Syst Bot 2000; 13: 803–815. [78] So ML. Studies on Plagiochila subgenus Metaplagiochila (Plagiochilaceae, Hepaticae). Austral Syst Bot 2001, 14: 677–688. [79] So ML. Plagiochila (Hepaticae, Plagiochilaceae) in China. Syst Bot Monogr 2001a; 60: 1–214. [80] So ML, Grolle R. Studies of Plagiochila sect. Subtropicae in Asia. Bryologist 1999; 102: 67–75. [81] So ML, Grolle R. On the Plagiochila species with paraphyllia or mamillose stems (Hepaticae). Syst Bot 1999; 24: 297–310. [82] So ML, Grolle R. Description of Plagiochila detecta sp. nov. (Hepaticae) from East Asia based on morphological and RAPD evidence. Nova Hedwigia 2000; 71: 387–393. [83] So ML, Grolle R. A checklist of Plagiochila (Hepaticae) in Asia. J Hattori Bot Lab 2000; 88: 199–243. [84] So ML, Grolle R. On Plagiochila subgenus Plagiochila section Abietinae (Hepaticae). Syst Bot 2001; 26: 459–469. [85] Spruce R. Hepaticae of the Amazon and the Andes of Peru and Ecuador. Trans. & Proc. Bot. Soc. Edinburgh 1884–1885; 15: i-xi, 1–589, plates i-xxii. [86] Stech M, Frey F. CpDNA-relationship and classification of the liverworts (Hepaticophytina, Bryophyta). Nova Hedwigia 2001; 72: 45–58. [87] Stech M, Quandt D, Frey W. Molecular circumscription of the hornworts (Anthocerotophyta) based on the chloroplast DNA trnL-trnF region. J Pant Res 2003; 116: 389–398. [88] Stephani F. Species Hepaticarum. Vol. II. Acrogynae. Geneva: Georg & Cie, 19011906.

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[89] Vanden Berghen C. Le genre Plagiochila (Dum.) Dum. (Hepaticae) à Madagascar et aux Mascareignes, principalment d’après les récoltes de M. Onraedt. Bull Jard Bot Belg 1981; 51: 41–103. [90] Wilson R, Gradstein SR, Heinrichs J, Groth H, Ilkiu-Borges AL, Hartmann FA. Phylogeny of Lejeuneaceae: a cladistic analysis of chloroplast gene rbcL sequences and morphology with preliminary comments on the mitochondrial nad4-2 spacer region. Monogr Syst Bot Missouri Bot Gard 2004; 98: 188–202.

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Genome Studies of Cactus Yeast MIGUEL DE BARROS LOPES1 and PHILIP F. GANTER2 1

School of Pharmacy and Medical Sciences, University of South Australia, Adelaide, Australia 2 Biology Department, Tennessee State University, 3500 John Merritt Blvd., Nashville, TN 37209, USA

ABSTRACT The cactophilic yeast, that live in pockets of decaying cactus tissue, are the best-studied natural yeast community from an ecological and evolutionary standpoint. Most of the yeast species commonly isolated in the necrotic tissue of cacti are specific to this habitat and the insects that live in it, with very little overlap between neighbouring yeast communities. The first part of this review summarizes the factors that shape the cactus yeast community, which include the cactus species, geography, the distribution of insects and yeast-yeast interactions. The subsequent part of the review focuses on recent molecular studies that analyze the genetic variation that exists between strains of individual species or related species, and which provide insight into the population structure of cactophilic yeast.

THE CACTUS-YEAST—DROSOPHILA COMMUNITY The Host Cacti

The cacti consist of approximately 100 genera and 1,500 species that are native to the Western Hemisphere, ranging from northern Canada to the Straits of Magellan. Several species have been introduced into many Address for correspondence: Miguel de Barros Lopes, School of Pharmacy and Medical Sciences, University of South Australia, City East Campus, North Terrace, Adelaide, SA 5000, Australia. E-mail: [email protected].

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other parts of the world, including the Mediterranean region, the Middle East, South Africa, India, Hawaii and Australia. In some areas, with their ability to withstand long periods of drought, they have become serious pest weeds [4]. Their ability to withstand drought is partially due to their capacity to store water, which makes them veritable oases of abundant moisture in the desert for animals and microbes that can colonize and utilize their tissue. When cacti of some species are injured, they can be infected by a community of organisms, dominated early on by Drosophila and yeast, that produce a long-lasting necrotic wound, commonly called a soft rot, due to the activity of pectinolytic bacteria. Not all large cacti seem to be susceptible to soft rotting, but little is known about the factors that might make some cacti immune. Host species susceptible to infection are grouped into three types that correspond to three major groups in the Cactaceae: (1) Opuntieae species (prickly pears), (2) Pachycereinae species (columnar cacti such as saguaro [Carnegiea gigantea] and cardón [Pachycereus pringle]), and (3) Stenocereinae species (columnar cacti such as the organ pipe cactus [Stenocereus thurberi] [18]). This situation may actually be more complicated as the South American cacti are both more diverse and less studied. The three cacti clades studied are chemically distinct [28], and this has been shown to have a significant effect on the Drosophila and yeast community that utilize the host plant [8, 10, 24, 28, 50]. For example, species in the subtribe Stenocereinae contain large quantities of water-soluble triterpene glycosides and unusual lipids, whereas species in the subtribe Pachycereinae possess complex alkaloids. Each of these compounds can be inhibitory to the growth of both yeast and the insects responsible for transporting the yeast to the next rot [8, 50]. Moreover, nutrients in these plants are often complexed with other molecules and not readily available. The Opuntia plants appear to be closest in composition to that of a ‘typical plant’ but the tissue contains large amounts of mucoid polysaccharides. Finally, morphological differences among cacti may alter edaphic habitat parameters such as temperature. Cactophilic Insects

The insects associated with cactus necroses are diverse, exploiting the cactus tissue and the microbes that colonize the necrosis, or preying on one another. Moran [34] references 122 different phytophagous insect

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species from the Coleoptera, Lepidoptera, Hemiptera, Diptera, and Hymenoptera associated with Opuntia species. The insect communities from other cacti are not so well-known, as Opuntia species have been intensively sampled to limit their growth with biocontrol agents [4]. Some moths in the tribe Phycitini (Pyralidae: Cactoblastis, Melitara, Sigelgaita) lay eggs on Opuntia and their larvae penetrate the tough cuticle and feed on the tissue underneath. One cactophilic yeast species (Clavispora opuntiae) has been associated with these moths, but it is not clear whether the moth is a vector for the yeast [30] (referred below). More probably, the moths initiate soft rots that are attractive to the actual vectors [43, 44, 45]. Although numerous cactus associated insects exist, the first insects to arrive are often species of Drosophila, which appear to be the most important insects in vectoring yeast and, therefore, the most well studied [61]. The Drosophila species associated with cactus necrosis generally belong to the mulleri subgroup of the repleta group, native to the Americas [70]. These species have a diffused symbiotic mutualistic relationship with cactus yeast. The yeast influence many aspects of Drosophila development and behaviour and the Drosophila are necessary to transport the yeast to new sites [19, 57]. So long as insects carrying the yeast are allowed to access the necrotic tissue in cacti, the yeast are able to exploit nutrients in this tissue and are, therefore, among the first microorganisms to colonize the rot. If insects are excluded, the rot is only colonized by bacteria and this gives rise to soft rot, particularly in the presence of pectinolytic Pectobacterium [20]. When yeast are present, their growth is rapid and they provide an important source of nutrients for the larval and adult flies [47]. Furthermore, the yeast produce volatiles that serve as important cues to Drosophila adults that are in transit from one location to another, as well as to the foraging larvae [6]. In turn, Drosophila are major vectors for the dispersal of yeast to new rots [7, 11, 22], and as the Drosophila exhibit an extensive degree of specialization in terms of host plant selection, it is expected that they strongly influence the yeast community composition. The four unrelated Drosophila species endemic to the Sonoran Desert are: D. pachea (nannoptera group), D. nigrospiracula (anceps complex), D. mojavensis (mulleri group), and D. mettleri (eremophila complex). An examination of their distributions may serve as a model of the ecology of cactophilic yeast vectors. The flies’ distributions are largely limited by the

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distribution of their specific host plants [23, 24, 27]. D. pachea is limited to senita because of a requirement for ,7–sterol–its cytochrome P450 system permits this species to tolerate the toxic alkaloids present in the host [9, 21]. Drosophila nigrospiracula is found in the decaying tissue of two cacti related to senita—saguaro and cardón—because senita alkaloids are toxic to this species. D. nigrospiracula is excluded from organ pipe cactus by the medium chain fatty acids and sterol diols found there. It is unclear why it does not inhabit Opuntia, although it has been suggested that the larvae are inhibited by the slime present in rots of this species [24]. D. mettleri, although resistant to senita alkaloids, is dependent on soil soaked by decaying saguaro and cardón for breeding [24]. D. mojavensis has a broader distribution, breeding in organ pipe cactus when given the choice, but is able to breed in agria and Opuntia as well as other species. The species is sensitive to senita alkaloids and is prevented from breeding on saguaro by D. nigrospiracula [24]. Similar studies on the Drosophila species that inhabit cacti in South America have shown that D. koepferae breeds primarily on the rotting stems of columnar cacti, whereas D. buzzatii breed primarily on the necrotic cladodes of Opuntia cacti. Both hosts appear to be suitable for viability for either of the flies, but differences in spatial and temporal availability of the rots appear to affect oviposition site preference and behaviour of the two Drosophila species. The Cactophilic Yeast

Several studies have analyzed the distribution of yeast in cacti rots from many regions [1, 10, 14, 52, 54, 63]. A major paper [63] reviewed the distribution of yeasts from 1,885 cactus rots, which included 50 different cactus species. The investigation centred on cacti from their native habitat, with 121 different localities in five defined geographic regions of the Americas (Caribbean, Sonoran Desert, southern Mexico, southwestern United States and Venezuela) being studied. Analysis of the cactophilic yeast community in countries where the cactus has been introduced (mostly studied in Australia and Hawaii), provides an interesting comparison [1, 65]. The results demonstrate the specificity of the cactophilic yeast community. Most of the yeast species commonly isolated in the necrotic tissue of cacti are specific to this habitat and the insects that live in it. There is very little overlap between cactophilic communities and neighbouring yeast communities, even with those

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growing in cactus fruits or flowers of the same plant [10, 44, 45, 46, 63]. While the fruit yeast show a greater discrepancy between localities and appear to be influenced by neighbouring yeast communities, the yeast isolated from cactus rots are distinct. The validity of this statement has increased since some isolates initially classified as members of species associated with non-cactophilic habitats have been shown to be distinct cactus rot-specific species upon closer inspection [35, 40, 42]. The findings also show that the yeast species composition is influenced more strongly by the host plant than by geography, although there is some suggestion of yeast shifting hosts where cacti have been introduced into new regions [1]. The reasons for the discrete cactus yeast community and its persistence [1, 32] include the specificity of insect vectors, yeastyeast interaction and plant chemistry. Cactus-Yeast Interactions

The chemical composition of the cactus has a significant effect on Drosophila host plant selection, which in turn is expected to affect the yeast that inhabit particular cactus species. Although cactophilic yeast generally grow on tissue from any cactus species (Ganter, unpublished results), the metabolites produced by the cacti, both stimulatory and inhibitory, are likely to have a direct impact on shaping the yeast community [51]. One such notable example is seen with the two varieties of Starmera amethionina. Strains of Starmera amethionina var. amethionina, a species usually associated with cacti of the subtribe Stenocereinae, possess a single dominant gene that minimises the inhibitory effect of the triterpene glycosides present in the host species. Starmera amethionina var. pachycereinae do not possess this gene and are generally limited to Pachycereinae hosts. Minimal or no gene flow occurs between the two varieties and it has been suggested that they are at the early stages of speciation [50]. Interestingly, Australian isolates of S. amethionina growing on Opuntia are sensitive to triterpene glycosides, although, based on DNA studies, they are much more similar to the variety amethionina. This yeast, although originating in triterpene glycoside possessing columnar cacti, is presumed to have lost resistance since its transfer to Australia. The division in habitat between the two varieties is not complete, however, and there are other factors that must also affect the distributions of these yeast - there is no obvious barrier preventing S. a. var. pachycereinae from colonizing Opuntia rots and in

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some regions (Tucson, AZ, in the northern Sonoran Desert) isolates of S. a. var. amethionina are found in Opuntia and S. a. var. pachycereinae in saguaro. Moreover, an alteration for growth between groups of Cactaceae is not a requirement, and other yeast species are commonly isolated from Stenocereinae, Pachycereinae and Opuntiae, and do not display the host shift adaptations demonstrated by Starmera amethionina. A similar host shift is currently unable to explain the triterpene glycosides tolerance of Candida orba [65]. C. orba forms part of the Phaffomyces opuntiae complex, which is composed of four taxa: Phaffomyces opuntiae, P. thermotolerans, P. antillensis as well as Candida orba [65, 71]. Their geographic distribution poses an interesting and, as of yet, unanswered question. P. thermotolerans and P. antillensis are found in Pachycereinae rots in the northern Sonoran Desert or from a limited set of islands in the Caribbean, respectively. Their closest relative, P. opuntiae, is found commonly in Australia on introduced Opuntia hosts. It has been found in only a single rot in over 2,500 rots sampled in Venezuela, the Caribbean, and North America [14, 63 and Ganter, unpublished data]. It was thought that further sampling in South America, especially Argentine Opuntia species, would reveal the native host and distribution of this yeast. However, over 500 South American rots (including 87 from Argentina) have since been sampled without a single isolate of P. opuntiae being found (Ganter, unpublished data). There are regions of South America rich in cacti yet to be sampled (Bolivia, much of the Peruvian altiplano, Ecuador, Columbia, Guyana) but the secret of the origin of P. opuntiae is proving more difficult to solve than predicted. Candida orba’s distribution poses a similar question. Only a handful of strains have ever been isolated and all were collected from Opuntia plants within 50 km of Brisbane, Queensland [65]. It has never been collected in the New World but is closely related to the Sonoran species, P. thermotolerans. However C. orba strains are resistant to the triterpene glycosides produced by Stenocereinae cacti, even though these compounds are not produced by the Opuntia that C. orba inhabit, or the hosts of P. thermotolerans, Pachycereinae cacti. Other cactophilic yeast also possess potentially beneficial physiological properties for residing on cactus soft rots. Candida caseinolytica demonstrates strong extracellular proteolytic activity, a rare activity in ascomycetes [41], whereas Dipodascus starmeri possesses lipase activity that permits its effective growth in lipid rich columnar cacti [42].

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Drosophila-Yeast Interactions

The close association of yeast and Drosophila suggests that these insects may have an important influence on yeast community structure. Several studies demonstrate that specific Drosophila/yeast interactions exist in the cactus habitat, providing a means for co-adaptation of the two organisms. Drosophila are able to discriminate between yeast species for food site selection [5, 6, 33]. In some cases, the yeast preferences of the Drosophila revealed in laboratory tests are consistent with abundance in the natural habitat. For example, Pichia cactophila, the most commonly isolated cactus yeast, is the preferred food of Drosophila mojavensis larvae [5, 6]. However, Candida sonorensis, the second most commonly isolated cactus yeast, was avoided by larvae. Drosophila buzzatii, a fly that breeds on Opuntia stricta rots, also showed preferences in oviposition site selection (which were similar to the feeding preferences of the adult flies) and the preferred yeast for oviposition were generally the species most commonly found in Opuntia stricta [2, 69]. The yeast available for feeding have been shown to affect the growth and fecundity of the Drosophila, with feeding on mixed yeast rather than a monoculture generally more beneficial to the flies [55, 62]. This is most apparent on yeast growing on their natural substrate—especially when nutrition is limited. Yeast are transmitted between courting flies [60] and in some species the quality of the yeast offered by the male can influence mating success and fecundity [37, 67, 68]. Specific yeast/Drosophila interactions have been shown to be beneficial for flies growing on particular cacti. Growth of the lipophylic yeast Dipodascus starmeri and P. mexicana may improve the suitability of the organ pipe necrosis tissue for supporting development of D. mojavensis larvae by providing a food source or eliminating the cactus lipids that are normally toxic to flies [52, 53]. If this is true, however, it remains to be explained why other Drosophila species such as D. nigrospiracula are excluded from organ pipe rots [24]. The presence of 2propanol metabolizing yeast, Candida sonorensis and Sporopachydermia cereana, increases Drosophila growth when present at concentrations similar to that in cacti rots. The presence of 2-propanol without the yeast leads to toxic effects on the Drosophila, indicating that the yeast benefit the flies by decreasing its concentration [57]. Similarly, the D. mojavensis life-span was also enhanced by C. sonorensis when grown on methanol since C. sonorensis assimilates this alcohol [12].

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Yeast-Yeast Interactions

Zymocidal or killer yeast produce an extracellular protein that is able to kill other yeast. Killer toxins are a diverse set of proteins with different temperature and pH optima, employing various mechanisms of lethality. Some are encoded by nuclear genes and others by genes on plasmids, which may be RNA or DNA. Each protein acts against a specific and limited set of sensitive strains. Several cactophilic species are killers but the activity may vary greatly even among strains from the same species [13, 58 and Ganter, unpublished data]. Pichia kluyveri killer strains (that produce killer protein with a low pH optimum, characteristic of recently colonized cactus rots) were shown to be able to exclude competitors from cohabiting cactus rots [13, 58]. However, killer strains of Phaffomyces thermotolerans were found together with strains sensitive to their toxin in saguaro, which is a very large cactus with persistent rots that are often neutral or basic. In this case the killer factor might provide a means of invading already colonized rots that contained strains sensitive to the killer’s toxins. Yeast can be beneficial for the growth of a second yeast [55]. For example, the presence of Pichia cactophila growing on agria-stem rots can double the growth rate of other species. This mutualism might be due to cross-feeding (for example, the ability of some species to free bound sugars—like those in triterpene glycosides – thus making them available to all yeast in the rot) or the catabolism of a growth inhibitory substance by specific yeast. Ganter and Starmer [13] have also demonstrated that the presence of a mixed yeast population can reduce the impact of killer toxin producing species on the survivability of sensitive strains on native host tissue. GENOME STUDIES OF CACTOPHILIC YEAST

Close to 100 different yeast species have been isolated from necrotic cactus tissue, but despite this complexity, the cactus yeast community is well defined with specific species and species complexes dominating the habitat. Pichia cactophila, Candida sonorensis and Sporopachydermia cereanus (and related species) are cosmopolitan - occurring in all cactus types and in all regions [63]. These are the most commonly isolated species, accounting for more than half of all yeasts isolated from the cactus habitat. Other yeasts generally show increased specificity in terms

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of region and host plant distribution (the two are confounded). These yeast include the Pichia kluyveri complex and a number of other distantly related Pichia species, the Phaffomyces opuntiae complex, the Starmera amethionina complex, Dipodascus starmeri, Clavispora opuntiae, Myxozyma mucilagina, Pichia mexicana and an undescribed species related to Ogataea polymorpha (=Pichia angusta or Hansenula polymorpha). Several factors of the cactophilic yeast community: including the prevalence of the core species, its relationship with Drosophila species and its independence from neighbouring yeast, have led Starmer and colleagues to conclude that the yeast-Drosophila-cactus community is relatively old and coadapted [55, 63]. Partial rRNA sequence comparisons between cactophilic species and related yeast support these conclusions and demonstrate that the community is polyphyletic - the core species having independent origins [66]. Recently, genetic variation between multiple strains of a single or closely related species has been assessed for several of the most important cactophilic yeast. The following sections will describe these comparative genetic studies and provide insight into the population structure of this community. The Pichia cactophila Species Complex

Pichia cactophila is the most common yeast isolated from cactus [25, 63], being part of the 36% GC (guanine-cytosine) species complex that includes Pichia norvegensis and P. pseudocactophila [57]. These latter two species are less common in cacti, and are geographically as well as host tissue specific. P. pseudocactophila is restricted to Sonoran Desert Pachycereinae rots, whereas P. norvegensis is mainly restricted to Opuntia from the Sonoran Desert and the Caribbean. It has also been listed as a human pathogen, but no DNA studies have confirmed that the cactus and human isolates are members of a single taxon. Ribosomal DNA analysis groups these yeast with a number of species (mostly Pichia) that have their origin in decaying fruit [29, 48, 66]. This clade also contains several other region or host specific cactus yeasts, like P. kluyveri, P. deserticola and P. heedii. Thirty-two Pichia cactophila strains isolated from several localities (the United States, the Caribbean and Argentina) and several host types have been studied using Random Amplified Polymorphic DNA (RAPD) in order to compare the relatedness of strains with geography and cactus host [15]. The results demonstrate that the geographic range is far more

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influential than host type in determining the similarity of yeast at the strain level. Strains isolated from southern Florida and the Caribbean grouped together, whether isolated from Opuntia or Stenocereinae rots. Interestingly, isolates from these two regions (2,200 km apart) were more similar to each other than southern Florida isolates were to those in northern Florida (560 km apart and all collected from Opuntia). The lack of distance effect is also demonstrated by the higher similarity of strains collected in Arizona with Argentinean isolates than with other Sonoran Desert strains, or strains collected from Texas. Consistent with the molecular data, the killer/sensitive phenotypes were more similar between Southern Florida and Antigua strains than between strains from southern and northern Florida. This significant influence of geography on intraspecific variation is in contrast to the influence of host on species community structure. Pichia norvegensis strains were also compared to each other and to the P. cactophila yeast (too few P. pseudocactophila strains are available for analysis) [15]. P. cactophila and P. norvegensis are phenotypically related, and based on DNA studies belong to the same species complex. The RAPD data confirm that the P. norvegensis strains formed a separate clade to P. cactophila. The isolates were again separated based on geography. However, the strains collected from the Caribbean were more closely related to strains from Argentina. Isolates from the Sonoran Desert produced a second lineage. The Pichia kluyveri Complex

As a comparison to P. cactophila, similar studies using amplified fragment length polymorphism (AFLP) and RAPD data were performed on Pichia kluyveri [16]. This species is in the same clade as P. cactophila but only distantly related [29, 48, 66]. Pichia kluyveri is atypical, i.e. it clearly occurs in both the cactophilic habitat and other habitats, in particular that of acidic fruits, including cactus fruit, tomatoes and oranges [10, 14, 64]. Therefore, by analyzing P. kluyveri, it is possible to establish whether spanning the cactophilic and non-cactophilic habitats creates any genetic barriers between strains of a single species. Pichia kluyveri was originally divided into three varieties based on reduced intervarietal DNA-DNA reassociation and spore viability [40]. These divisions are consistent with minor differences in physiological

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characters, specifically the ability to ferment glucose and the presence of killer activity [16, 40]. Pichia kluyveri var. eremophila and P. k. var. cephalocereana are specialists in both geography and host: P. k. var. cephalocereana has only been isolated from columnar cacti of the Pachycereinae tribe on a single island in the Caribbean; while P. k. var. eremophila has only been found in Opuntia and columnar cacti, but only in Mexico and the south-western United States. Interestingly, strains consistent with the P. k. var. eremophila phenotype have been isolated from Stenocereinae cacti on Caribbean islands, but no genetic testing has been done to establish that they actually belong to this variety (Ganter, unpublished data). Pichia kluyveri var. kluyveri is a generalist – it has been isolated from all regions, including Opuntia introduced in Australia, and from both cactus rots and fruits. As for Pichia cactophila, there is a strong influence of geography on the population structure of P. kluyveri, but there is a more obvious link with host type as well [16]. For example, a strain isolated from an Opuntia rot in Mexico had closer genetic similarity to Opuntia isolates from Arizona than to columnar cacti isolates in the same area. Also, in Florida collections, tomato isolates were distinguishable from Opuntia isolates. However, there is no barrier between strains of the Opuntia and tomato habitats as the strains between the two hosts are often more similar than those collected from either tomato or Opuntia. Interestingly, the single characterized Australian isolate grouped with many of the tomato isolates. The data also provided molecular support for P. k. var. cephalocereana as a monophyletic group [16]. All the yeast, except one, that showed physiological characters consistent with the varietal description formed a monophyletic clade. The one exception is actually more similar to a different species, Pichia barkeri, and is most easily explained by convergent evolution. Although Pichia barkeri has not been intensively studied genetically, this yeast highlights interesting aspects of genome evolution. Pichia kluyveri yeast possesses a 30% G+C content. Pichia barkeri, in contrast, possesses 36% G+C content, the same as the P. cactophila group of yeasts [57]. Surprisingly, despite the 6% discrepancy in G+C %, P. barkeri is much more closely related to P. kluyveri, a conclusion supported by partial rDNA sequencing [29, 66]. Pichia heedii, also closely related to P. kluyveri and P. barkeri, has an intermediate G+C content (between 32 and 33%), whereas Pichia

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deserticola, another cactophilic species in the same clade (although different subclade to both P. kluyveri and P. cactophila), has a very low G+C content (27%). The mechanisms accounting for these striking differences in G+C % are unknown [57]. Added to highly variable %GC values in closely related species, genetic divergence in cactophilic yeast is also apparent between strains of the same species. Although cactophilic strains generally exhibit minimal phenotypic variation, the level of intraspecific genetic variation is considerable. For example, in P. kluyveri, the average variation between the RAPD fingerprints was 19%, with very few bands amplified in all strains of the species [16]. This disparity was even higher in P. cactophila (28%) [15] and in results obtained for Candida sonorensis (24%) [17] (detailed below). Likewise, DNA-DNA reassociation value observed between strains of the same species is often below the 70% value generally accepted as the minimal for conspecifics, and rDNA sequence variation can be considerable [29, 30]. This contrasts with the results for Saccharomyces cerevisiae which appear to be very homogeneous [3, 36]. Additional evidence for an unusual level of genetic variation within and among related cactophilic strains emerges from comparisons between rDNA sequence divergence and DNA-DNA reassociation [29, 36]. Amongst 54 comparisons, 7 involving cactophilic species, there is no difference between the average percentage of DNA-DNA reassociation among cactophilic strains and the average for the other 47 comparisons. However, there is a difference in genetic distance among comparisons based on partial rDNA sequences. The average pairwise difference for cactophilic strains (17.4 bp) is almost five times that for the noncactophilic comparisons (3.7 bp). Although this is not a randomly chosen sample of cactophilic or non-cactophilic yeast, some of the variances described in these different studies may be partially explained by limitations or discrepancy in methodology. The results might also be explained by the age of the cactophilic species, increased rates of sequence divergence for at least portions of the genome, or even differences in the mechanics of speciation. For example, functional meiosis in Saccharomyces sensu stricto interspecies hybrids is much greater when the parental strains are defective in DNA mismatch repair [26].

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Candida sonorensis

Candida sonorensis is one of the three ubiquitous cactus yeast, found globally in all cactus species studied and unique to the cactus habitat [17, 63]. It is distinctive in its ability to assimilate methanol, a breakdown product of plant cell walls, consistent with results based on rDNA sequencing that indicate it was derived from tree fluxes [66]. A number of different methods have been used to look at 36 strains collected from a large geographic range—two distant sites in the USA, Texas and Florida, and from Australia—but limited to Opuntia rots [17]. A significant correlation between several of the methods exists, most clearly between DNA-DNA reassociation studies and RAPDs. Based on this data, the strains could be divided into two main groups: nine strains collected from West Texas formed group 1, and all the others, which include the isolates from Australia, Florida, East Texas and three additional strains from West Texas but collected in an earlier year (1991 versus 1994) formed the second group. Only the nine West Texas strains were able to assimilate glycerol, the physiological character that is most consistent with the division. Based on the RAPD data, ten of the twelve Australian isolates formed a separate clade. Interestingly, there is minimal correlation between rDNA sequences and any other type of variation measured, highlighting the caution required when using a single locus for phylogenetic studies. The genomes of C. sonorensis strains appear to be highly diverged. The electrokaryotype and DNA-DNA reassociation studies indicate considerable variation among the 36 strains studied, even though they are all from Opuntia and only from part of the known geographic distribution. As no sexual reproduction is known for this species, the variation may indicate that this is an ancient asexual lineage. The Sporopachydermia cereana Species Complex

The Sporopachydermia cereana species complex is the third group of yeast that are commonly and globally isolated from cactus necrosis [63]. These yeast are distinguished by their ability to assimilate erythritol as a sole carbon source. Similar to what is seen for P. barkeri and P. kluyveri, the G+C % content varies widely for the species complex, 38 – 50%, but genetic relatedness between the strains is evidenced by DNA-DNA reassociation studies and rDNA sequencing [31]. The complex, with origins in temperate tree-fluxes, has been separated into ten species or

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sub-species; although based on recent collections from South America (Ganter, unpublished data), the number of species in the complex is probably considerably larger than the ten currently known. Based on this division, none of the species on their own have been found in all cactus types or in all regions. However, as seen for the other species analyzed, the distribution appears to be shaped mainly by the geographic range and not host species [31]. Of the main groups, Sporopachydermia australis yeast are localised in South America, S. opuntiana are mainly found in the Caribbean (and Australia), whereas S. cereana are found in the Sonoran region, southern Mexico and Haiti. S. cereana has been further divided into three sub-groups and these inhabit the three distinct regions. Clavispora opuntiae

Clavispora opuntiae is a common cactus yeast with an unusual association with moths. In Brazil, the yeast host is a columnar cactus [43] and the moth is a species of Sigelgaita, but a better-studied association is between the yeast and an Opuntia-boring moth, Cactoblastis cactorum, another Phyticid moth [39, 59, 63]. Over 500 strains of this species collected worldwide have been analyzed using rDNA restriction mapping [30]. The conclusion is that the distribution of this species is closely associated with the movements of the moth Cactoblastis cactorum, an insect that has been used widely and successfully in the biological control of Opuntia weeds [4]. Cactoblastis cactorum was particularly successful in decimating the Australian population of O. stricta and has since been introduced into many cactus growing regions, including Hawaii and South Africa, for weed control. In the early 1960’s, the moth was released on some of the smaller Leeward Islands of the Caribbean to control native Opuntia populations that had expanded onto the land cleared for animal grazing [49]. By the end of the decade, the moth had reached the larger islands and by 1987 was infesting Opuntia populations in the Florida keys [59]. The moth has continued to spread in Florida and is now recognized as a threat to Opuntia populations throughout North America, although it is not known if it will have the same effect in areas where there are already native Opuntia-boring moths [38]. Lachance and colleagues have shown that the occurrence of the moth leads to the dominance of a few related C. opuntiae strains [30]. For example, in Maui, Hawaii, where cacti were introduced from Mexico in 1809 and Cactoblastis cactorum introduced from Australia in 1949, a single genotype of C. opuntiae

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appears to be ubiquitous, all isolates possessing identical rDNA sequences and almost all belonging to a single mating type. In areas where the moth does not exist, the genetic variation of the C. opuntiae isolates is much greater. These results indicate that the dispersal of this single moth species has caused the global spread of a small number of related strains. Other Cactophilic Yeast Species

Although several other cactophilic species exist, the population structures of these yeast have not been well documented. The genera Phaffomyces and Starmera, and the species Dipodascus starmeri, are most likely derived from species living in tree-fluxes [66, 71]. Myxozyma mucilagina is commonly isolated from Opuntia rots in North America and the Caribbean, and shares a peculiarity with its host: both produce copious amounts of mucopolysaccharides, an unusual characteristic for a yeast. Partial rRNA sequencing positions this species in a clade containing species isolated from trees, with deeper origins in fruit habitats [66]. Pichia mexicana is not closely related to any other cactophilic yeast. It was originally reported from columnar cacti in the Sonoran Desert, but has since been isolated more widely. It is found in Opuntia cacti in Venezuela [63], and less frequently in Australia, Brazil, and Argentina, and more rarely in columnar cacti from Peru. Three Candida species (C. entomoaea, C. veronae, and C. terebra) all have partial LSU rDNA sequences identical to that of P. mexicana but come from a diverse set of locales and substrates (insect tunnels in South African pines, grape juice in Italy, and Japanese soils, respectively). At this time, we can say little about the origin of this species or its role in the cactophilic community. Finally, there is evidence of another undescribed cactophilic species Ogataea polymorpha (= Pichia angusta = Hansenula polymorpha ) that has long been isolated from cactus samples, primarily from Opuntia rots. The species has been identified from many different habitats but the cactus strains have a G+C proportion a full percentage point higher than noncactus strains and show phenotypic divergence—slow growth on methanol and tolerance to higher maximum temperatures for growth. CONCLUSION

The cactophilic yeast community is distinct from surrounding yeast communities, even to those associated with fruits or flowers on the same

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plant. The basis for this separation includes cactus chemistry, the distribution of insects and yeast-yeast interactions. The diverse yeast species have adapted to the cactus habitat independently, and most likely coevolved with the Drosophila that feed and oviposit on the same hosts. This puts the origin of the cactus-yeast community to approximately 50 million years ago, together with the derivation of the Cactaceae themselves. One of the themes that runs through the discovery of the cactophilic yeast is that there are often either multiple related species (species complexes) with very similar physiological profiles or substantial genetic variation between strains of a single species. This is likely to be, at least partially, due to the forces promoting genomic changes in species living in this habitat. Whereas the host species appears to be the main factor dictating the yeast community structure at the species level, a number of recent genome comparative investigations show that the intraspecific variation appears to be influenced by geographic range more than host. Preliminary genomic explorations demonstrate that the cactophilic yeast community provides an excellent model to study the processes and structures found in living systems. The yeast are sufficiently diverse to be interesting and yet limited enough, in terms of habitat, to be tractable. What makes this collection of yeast uniquely valuable (as compared to the current collection of yeast genomes being sequenced) is the ecological knowledge with which genomic information can be combined. It is only with the integration of such information that the forces influencing genome-phenome relationships can be understood. Acknowledgments

The authors would like to thank Sonia Dayan for critical reading of the manuscript. References [1] Barker JSF, East PD, Phaff HJ, Miranda M. The ecology of the yeast flora in necrotic Opuntia cacti and of associated Drosophila in Australia. Microbial Ecology 1984; 10: 379–399. [2] Barker JSF, Starmer WT. Environmental effects and the genetics of oviposition site preference for natural yeast substrates in Drosophila buzzatii. Hereditas 1999; 130: 145–175.

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[3] de Barros Lopes M, Rainieri S, Henschke PA, Langridge P. AFLP fingerprinting for analysis of yeast genetic variation. International Journal of Systematic Bacteriology 1999; 49: 915–924. [4] Dodd AP. The biological campaign against prickly pear. Commonwealth Prickly-Pear Board Bulletin. Brisbane, Australia, 1940. [5] Fogleman JC, Starmer WT, Heed WB. Larval selectivity for yeast species by Drosophila mojavensis in natural substrates. Proceedings of the National Academy of Sciences of the United States of America city. 1981; 78: 4435–4439. [6] Fogleman JC. The role of volatiles in the ecology of cactophilic Drosophila. In: Barker JSF, Starmer WT, eds. Ecological genetics and evolution: the cactus-yeast-Drosophila model system. Sydney, Australia: Academic Press, 1982; 191–208. [7] Fogleman JC, Starmer WT, Heed WB. Comparisons of yeast flora from natural substrates and larval guts of southwestern Drosophila. Oecologia 1982; 52: 187–191. [8] Fogleman JC, Abril JR. Ecological and evolutionary importance of host plant chemistry. In: Barker JSF et al. eds. Ecological and Evolutionary Genetics of Drosophila, New York, USA: Plenum Publication Company, 1990; 121–143. [9] Frank MR, Fogleman JC. Involvement of cytochrome P450 in host-plant utilization by Sonoran Desert Drosophila. Proceedings of the National Academy of Sciences of the United States of America 1992; 89: 11998–12002. [10] Ganter PF, Starmer WT, Lachance MA, Phaff HJ. Yeast communities from host plants and associated Drosophila in southern Arizona: new isolations and analysis of the relative importance of hosts and vectors on community composition. Oecologia 1986; 70: 386–392. [11] Ganter PF. The vectoring of cactophilic yeasts by Drosophila. Oecologia 1988; 75: 400–404. [12] Ganter PF , Peris F , Starmer WT. Adult life span of cactophilic Drosophila: interactions among volatiles and yeasts. American Midland Naturalist 1989; 121: 331–340. [13] Ganter PF, Starmer WT. Killer factor as a mechanism of interference competition in yeasts associated with cacti. Ecology 1992; 73: 54–67. [14] Ganter PF, Bustillo E, Pendola J. Yeast interactions inferred from natural distribution patterns. Florida Scientist 1993; 57: 50–61. [15] Ganter PF, Quarles B. Analysis of population structure of cactophilic yeast from the genus Pichia: P. cactophila and P. norvegensis. Canadian Journal of Microbiology 1997; 43: 35–44. [16] Ganter PF, and de Barros Lopes M. The use of anonymous DNA markers in assessing worldwide relatedness in the yeast species Pichia kluyveri Bedford and Kudrjavzev. Canadian Journal of Microbiology 2000; 46: 967–980. [17] Ganter PF, Cardinali G, Giammaria M, Quarles B. Correlations among measures of phenotypic and genetic variation within an oligotrophic asexual yeast, Candida sonorensis, collected from Opuntia. FEMS Yeast Research 2004; 4: 527–540. [18] Gibson A, Horak K. Systematic anatomy and phylogeny of Mexican columnar cacti. Annals of the Missouri Botanical Garden 1978; 65: 999–1057. [19] Gilbert DG. Dispersal of yeasts and bacteria by Drosophila in a temperate forest. Oecologia 1980; 46: 135–137. [20] Hauben L , Moore ER , Vauterin L , Steenackers M, Mergaert J, Verdonck L, Swings J. Phylogenetic position of phytopathogens within the Enterobacteriaceae. Systematic and Applied Microbiology 1998; 21: 384–397.

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[21] Heed WB, Kircher HW. A unique sterol in the ecology and nutrition of Drosophila pachea. Science 1965; 149: 758–761. [22] Heed WB, Starmer WT, Miranda M, Miller MWHJ. Phaff HJ. An analysis of the yeast flora associated with cactophilic Drosophila and their host plants in the Sonoran Desert and its relation to temperate and tropical associations. Ecology 1976; 57: 151– 160. [23] Heed WB. Ecology and genetics of Sonoran desert Drosophila. In: P. Brussard, ed. Ecological Genetics: the Interface, New York, USA, Springer, 1978; 109–126. [24] Heed WB. The origin of Drosophila in the Sonoran Desert. In: Barker JSF, Starmer WT, eds. Ecological genetics and evolution: the cactus-yeast-Drosophila model system. New York, USA, Academic Press, 1982; 65–80. [25] Holzschu DL, Phaff HJ, Tredick J, Hedgecock D. Resolution of the varietal relationship within the species Pichia opuntiae and establishment of a new species, Pichia thermotolerans comb. nov. International Journal of Systematic Bacteriology 1985; 35: 457–461. [26] Hunter N, Chambers SR, Louis EJ, Borts RH. The mismatch repair system contributes to meiotic sterility in an interspecific yeast hybrid. EMBO Journal 1996; 15: 1726– 1733. [27] Hurtado LA, Erez T, Castrezana S, Markow, TA. Contrasting population genetic patterns and evolutionary histories among sympatric Sonoran Desert cactophilic Drosophila. Molecular Ecology 2004; 13: 1365–1375. [28] Kircher H. Chemical composition of cacti and its relationship to Sonoran Desert Drosophila. In: Barker JSF, Starmer WT, eds. Ecological genetics and evolution: the cactus-yeast-Drosophila model system. Sydney, Australia, Academic Press, 1982: 143– 158. [29] Kurtzman CP, Robnett CJ. Identification and phylogeny of ascomycetous yeasts from analysis of nuclear large subunit (26S) ribosomal DNA partial sequences. Antonie van Leeuwenhoek 1998; 73: 331–371. [30] Lachance MA, Starmer WT, Bowles JM, Phaff HJ, Rosa CA. Ribosomal DNA, species structure, and biogeography of the cactophilic yeast Clavispora opuntiae. Canadian Journal of Microbiology 2000; 46: 195–210. [31] Lachance MA, Kaden JE, Phaff HJ, Starmer WT. Phylogenetic structure of the Sporopachydermia cereana species complex. International Journal of Systematic and Evolutionary Microbiology 2001; 51: 237–247. [32] Latham BP. Yeast community persistence in a spatially structured environment. Microbial Ecology 1998; 36: 60–65. [33] Morais PB, Rosa CA, Hagler AN, Mendonca-Hagler LC. Yeast communities of the cactus Pilosocereus arrabidae as resources for larval and adult stages of Drosophila serido. Antonie Van Leeuwenhoek 1994; 66: 313–317. [34] Moran VC. Interactions between phytophagous insects and their Opuntia hosts. Ecological Entomology 1980; 5: 153–164. [35] Naumov GI, Naumova ES, Kondrativea VI, Bulat SA, Mironenko NV, MendonçaHagler LC, Hagler AN. Genetic and molecular delineation of three sibling species in the Hansenula polymorpha complex. Systematic and Applied Microbiology 1997; 20: 50–56. [36] Naumov GI, James SA, Naumova ES, Louis EJ, Roberts IN. Three new species in the Saccharomyces sensu stricto complex: Saccharomyces cariocanus, Saccharomyces

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[53] Starmer WT. Associations and interactions among yeasts, Drosophila and their habitat. In: Barker JSF , Starmer WT, eds. Ecological genetics and evolution: the cactus-yeast-Drosophila model system. Sydney, Australia, Academic Press, 1982; 159– 174. [54] Starmer W, Phaff HJ. Analysis of the community structure of yeasts associated with the decaying stems of cactus. II. Opuntia species. Microbial Ecology 1983; 9: 247–259. [55] Starmer WT, Fogleman JC. Coadaptation of Drosophila and yeasts in their natural habitat. Journal of Chemical Ecology 1986; 12: 1035–1053. [56] Starmer WT, Barker JS, Phaff HJ, Fogleman JC. Adaptations of Drosophila and yeasts: their interactions with the volatile 2-propanol in the cactus-microorganism-Drosophila model system. Australian Journal of Biological Sciences 1986; 39: 69–77. [57] Starmer WT, Ganter PF , Phaff HJ. Quantum and continuous evolution of DNA base composition in the yeast genus Pichia. Evolution 1986; 40: 1263–1274. [58] Starmer WT, Ganter PF, Aberdeen V, Lachance MA, Phaff HJ. The ecological role of killer yeasts in natural communities of yeasts. Canadian Journal of Microbiology 1987; 33: 783–796. [59] Starmer WT, Aberdeen V and Lachance MA. The yeast community associated with decaying Opuntia stricta (Haworth) in Florida with regard to the moth Cactoblastis cactorum (Berg). Florida Scientist 1988; 51: 7–11. [60] Starmer WT, Peris F , Fontdevila A. The transmission of yeasts by Drosophila buzzatii during courtship and mating. Animal Behavior, 1988; 36: 1691–1695. [61] Starmer WT, Phaff HJ, Bowles JM, Lachance MA. Yeasts vectored by insects feeding on decaying saguaro cactus. Southwestern Naturalist 1988; 33: 362–363. [62] Starmer WT, Aberdeen V. The nutritional importance of pure and mixed cultures of yeasts in the development of Drosophila mulleri larvae in Opuntia tissues and its relationship to host plant shifts. In: Barker JSF, Starmer, WT, MacIntyre RJ, eds. Ecological and Evolutionary Genetics of Drosophila. New York, USA, Plenum Press, 1990: 145–160. [63] Starmer WT, Lachance MA, Phaff HJ, Heed WB. The biogeography of yeasts associated with decaying cactus tissue in North America, the Caribbean, and Northern Venezuela. In: Hecht MK, Wallace B, MacIntyre RJ, eds. Evolutionary Biology, New York, USA, Plenum Press, 1990: 253–296. [64] Starmer WT, Ganter PF , Aberdeen V. Geographic distribution and genetics of killer phenotypes for the yeast Pichia kluyveri across the United States. Applied and Environmental Microbiology 1992; 58: 990–997. [65] Starmer WT, Phaff HJ, Ganter PF, Lachance MA. Candida orba sp nov., a new cactusspecific yeast species from Queensland, Australia. International Journal of Systematic and Evolutionary Microbiology 2001; 51: 699–705. [66] Starmer WT, Schmedicke RA, Lachance MA. The origin of the cactus-yeast community. FEMS Yeast Research 2003; 3: 441–448. [67] Steele RH. Courtship feeding in Drosophila subobscura. I. The nutritional significance of courtship feeding. Animal Behaviour 1986a; 34: 1087–1098. [68] Steele RH. Courtship feeding in Drosophila subobscura. II. Courtship feeding by males influences female mate choice. Animal Behaviour 1986b; 34: 1099–1108. [69] Vacek DC, East PD, Barker JSF, Soliman MH. Feeding and oviposition preferences of Drosophila buzzatii for microbial species isolated from its natural environment. Biological Journal of the Linnean Society 1985; 24: 175–187.

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[70] Wasserman SA. Cytological evolution in the Drosophila repleta species group. In: Barker JFS, Starmer WT eds. Ecological Genetics and Evolution: The cactus-yeastDrosophila model system, J Barker, WT Starmer, eds. (Sydney, Academic Press), 1982: 49–64. [71] Yamada Y, Kawasaki H, Nagatsuka Y, Mikata K, Seki T . The phylogeny of the cactophilic yeasts based on the 18S ribosomal RNA gene sequences: the proposals of Phaffomyces antillensis and Starmera caribaea, new combinations. Bioscience, Biotechnology, and Biochemistry 1999; 63: 827–832.

PLANT GENOME: BIODIVERSITY AND EVOLUTION Volume 2, Part B Lower Groups

Author Index

Aarts HJM 236 Abarca ML 322, 323 Abbas B 177 Abbott SP 237 Abdel-Salam KA 354 Abdel-Sallam IS 329 Abdel-Satar MA 354 Abdelzaher HM 357 Abendstein D 235 Aberdeen V 478 Abliz P 326, 330 Abouhaidar MG 31 Abramson D 354 Abril JR 475 Accensi F 322, 323 Adamson J 111 Adee EA 380 Adler A 235, 357, 361 Aebersold R 77 Aert R 230 Afzal M 328 Agabian N 279 Agostini-Carbone ML 235 Aguinagalde I 404 Ahearn DG 230, 277, 279 Ahmad A 113, 115 Ahn JH 37 Ahonen I 453 Aida K 283

Aigle M 233, 237 Ainouche AK 400 Ainouche ML 400 Aitken EAB 381 Ajello L 324 Alabouvette C 356, 357 Albermann K 234 Albers A 329 Albers-Schonberg G 322 Alberts AW 322 Alcala-Jimenez AR 358 Aldridge C 236 Aldridge DC 330 Alekhina IA 363 Alexopoulos CJ 227, 322 Alfatafta AA 332 Ali M 328 Alice LA 400 Allain FHT 31 Allard MR 359 Allen B 405 Allen JR 110 Allington WB 380 Allison LA 427 Al-Musallam A 322 Al-Shehbaz IA 403 Alström-Rapaport C 403 Altmann, R 74 Altman S 34, 36

482

Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

Alvarez Buylla ER 403 Alvarez I 399 Ambros V 34 Ammirati JF 233 An SS 399 Anaissie EJ 361 Ananvoranich S 30 Andersen AA 30, 33 Anderson DM 78, 114, 115 Anderson JB 362, 380 Andersen RA 74, 114, 175 Andersson B 282 Andersson JO 74 Ando K 234 Ando Y 234 Angus A 239 Anonymous 169, 227 Antaramian A 83 Anton H 452, 453, 454, 455 Antonov AS 456 Aoki S 431 Aoki T 354, 355 Aota S 404 Appel DJ 227, 355, 380 Apt KE 74, 81 Archer S 328 Archibald JM 74, 81 Arisan-Atac I 239 Armbrust EV 169 Arnason U 401 Arnaud J 383 Arnheim N 361 Arnold JRP 37 Arora DK 323, 355 Artigvenave F 237 Asao T 326 Ashby SF 227 Assigbetse KB 355 Atkins JF 33 Auclair K 326 Avelange I 356, 357 Avila-Campillo I 236 Avise JC 74, 399 Avner P 30

Axelrood PE 356 Ayala FJ 383 Ayliffe MA 406 Azuma H 427 Baayen RP 355, 362, 363 Babinchak JA 110 Babjeva IP 284 Bachewich J 239 Bachman MS 380 Bachmann M 322 Bachvaroff TR 74, 110 Bacigálová K 236 Bacon CW 230 Bacon JS 277 Badr A 79 Baer MF 36 Bähler J 233 Bähr M 234 Bailey CD 399 Baillie BK 110 Bainbridge BW 358 Bakan B 358 Baker AC 110 Baker JL 323 Bakos JT 36 Balague V 113 Balajee SA 322 Baldauf JG 169 Baldauf SL 74, 86, 227 Baldwin BG 399, 400 Balick M 358 Ball LA 356 Ball RM 110 Ballou C 277 Bamba T 279 Banaszak AT 110 Bandoni RJ 277 Banerji S 235 Bang A 240 Banno I 277, 281, 283, 284 Bao JR 355 Bapteste E 74 Baral HO 230

Author Index

Barber RT 171 Barbrook AC 74, 110, 111, 112 Bárcena MA 171 Barker JS 478 Barker JSF 474, 477, 478 Barnett JA 277 Barnett P 77 Barns SM 229 Baron EJ 234 Barr ME 227 Barrales H 176 Barrell BG 279 Barrie FR 230 Barriel V 400 Barron JA 169 Bart-Delabesse E 380 Bartel DP 37, 38 Barthlott W 172, 400, 428 Bartnicki-Garcia S 227, 277 Bartnik E 358 Barton RC 381 Barve MP 355 Bassam BJ 356 Bassiouny A 228 Basson M 36 Bateman RM 400 Batenburg-van der Vegte WH 282 Bates SS 172 Batista LR 322 Batra LR 228 Battilani P 322 Baucom A 38 Bauer R 228, 235, 236 Baulcombe DC 33 Baum DA 402 Baum M 355 Baum MP 229 Baumel A 400 Baura G 355 Bayer MM 174 Bayer RJ 400 Bayman P 323 Beadle G 228 Beam CA 113

483 Beattie TL 30, 37 Beck JJ 355 Beckert S 400, 452 Béclin C 38 Been MD 31, 36, 37 Beerli P 380 Begerow D 228 Behnke A 169 Beklemishev CW 177 Belabid L 355 Bell CD 427 Bell PJ 277 Bellemere A 228 Beltz SB 326 Ben Ali A 74 Ben-Yephet Y 359 Benkovic SJ 33 Bennett CA 326 Bennett DE 238 Bennett FD 477 Bennett JR 177 Bennett JW 323 Benny GL 228 Bentley S 355 Bento Soares M 79 Beraldi-Campesi H 169 Berbee ML 228, 233, 239, 277 Beretta B 323 Berg G 233 Berg S 427 Berges JA 169 Bergquist PL 111 Bergquist PR 111 Berland B 111 Berney C 82 Bernhoft A 359 Bertozzini E 111 Bertuzzi AT 322 Besebdorfer V 407 Bettinge JC 36 Beyermann B 360 Bhagwat AS 356 Bhattacharya D 74, 82, 84, 87, 111, 116, 169

484

Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

Bhaud Y 113 Bianco L 111, 113 Biehl ML 332 Bievre C 326 Bigelis R 323 Bills GF 357 Binder M 235 Bingham S 111 Birren B 280 Birren BW 280 Bissett J 232 Bittkau C 404 Blackburn SL 110 Blackwell M 227, 228, 237-239, 282, 322 Blake Boles S 404 Blakesley B 112 Blanchard J 75 Blandin G 228 Blanz PA 230, 280, 355 Blaser P 322 Blattner FR 400 Blaze KL 176 Bleeker M 362 Blobel G 38 Blowers AD 428 Blue J 477 Bobrova VK 456 Bock A 428 Boekhout T 277, 278 Bogorad L 428 Bohnert HJ 81 Bolch CJS 110, 112, 114 Bolen PL 323 Boles S 406 Boles SB 405, 430 Bolter B 75 Bon E 228 Bonaldo MF 111 Bonfante P 359 Bonnard G 75 Booth C 355 Booy G 400 Bopp M 452 Borda EJ 31

Bordner J 330 Borkovich KA 230 Borner T 360, 362 Börner T 232, 239, 360, 428, 431 Borsch T 400, 428, 430, 455 Borts RH 476 Botes L 114 Botos B 331 Bottalico A 359 Botton B 356 Boundy-Mills K 280 Bouznad Z 355 Bowers B 281 Bowers HA 115 Bowler C 169 Bowles JM 476-478 Bowman BH 228, 238, 380 Boyd PW 169 Boyen C 75 Boyle JA 169 Brachat S 229, 278 Bradley PJ 75 Bragulat MR 322, 323 Branch AD 31 Brandt P 406 Brandt U 81 Braun EL 228 Braun U 228 Brayford D 355 Breitenbach M 235 Bremer B 405 Brenière F 383 Brennicke A 406 Bretagne S 380 Breuil C 233 Brewer S 404 Briatore E 360 Brick MA 356 Bridge PD 355 Brinkmann H 74, 81 Britton DM 401 Britz H 356 Briza P 236 Bronson CR 382

485

Author Index

Brookman JL 323 Brown AE 362 Brown F 31 Brown SC 400 Browning S 403 Brownlee AG 380 Bruening G 31, 36, 37 Bruice TC 37 Brumfield RT 380 Brund TD 357 Brune DC 77 Bruns T 363, 406 Bruns TD 229, 239, 355-357 Brutto R 77 Bryant DA 81 Brygoo Y 230, 358 Bucci TJ 228 Buchbender V 454 Buchi G 323 Buck WR 401, 428, 430 Buckler ES 400 Buddie A 355 Büdel B 173 Bui ETN 75 Bulat SA 361, 363, 476 Bulgheroni A 327 Bullerwell CE 229 Burdet HM 230 Burger G 75 Burger HJ 332 Burger-Wiersma T 75 Burgers K 324 Burke JM 31, 35 Burkhead KD 323 Burkholder JJ 278 Burkholder JM 81 Burns G 233 Burt A 232, 330, 383 Buscot F 356 Busico E 111 Busse I 75 Busse JS 78 Bussey H 279 Bussiere F 31

Bussink HJD 323 Busta FF 325 Bustillo E 475 Butcher SE 31 Butrym ED 281 Buxton FP 323 Buzayan JM 31, 36, 37 Byers B 229 Byme PF 356 Cabañes FJ 322, 323 Cabib E 281 Cachon J 110 Cachon M 110, 112 Caetano-Anolles G 356 Cai J 229, 278, 279 Cai Z 31 Callender M 327 Calton GJ 323 Calvo SE 230 Cambareri EB 232 Campbell CS 400 Campbell L 83 Campbell-Platt G 323 Cannon PF 232 Cannone JJ 401 Cano J 322 Capesius C 452 Capesius I 400, 456 Cappadocia M 82, 113 Caprara MG 31 Caputo P 406 Cardinali G 475 Cardwell KF 324 Carey M 355 Carl H 452 Carlos AA 110 Caron DA 111 Carothers JM 31 Carr TG 399 Carrum G 327 Carter JP 356 Cary JW 326 Casadevall A 230

486

Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

Casaregola S 228, 233 Casavant TL 111 Castaldo R 406 Castellá G 322, 323 Castrezana S 476 Cate JH 38 Cavalho S 456 Cavalier-Smith T 74, 75, 76, 79, 80, 84, 87, 114, 116, 229 Cavalier-Smith TC 237 Cavalieri D 278 Cech TR 31, 32, 33, 34 Cedergren R 32 Cembella AD 112 Cerbah M 407 Cevallos-Ferriz SRS 169 Chacón-Baca E 169 Chadwick BP 31 Chai JY 170 Chakrabarti A 356 Chalfoun SM 322 Chamberlain DW 380 Chambers SR 476 Chamuris GP 357 Chandler FW 328 Chang Z 404 Chao EE 75 Chapelle S 283 Chargaff E 278 Chartrand C 330 Chase MW 404 Cheddadi R 404 Chelkowski J 323, 359 Chen J 322 Chen JW 323 Chen KC 323 Chen L-J 429 Chen W 356, 380-382 Chen YC 381 Chen YJ 428 Cheng DJ 278 Cheng G 281 Chepurnov VA 169, 170, 174 Chesnick JM 110, 170

Chesnick JM 76 Chiachiera SM 324 Chiang TY 428, 400 Chiang YC 428 Chiappino ML 170 Chibana H 279 Chiche J 400 Childs JL 429 Chinain M 110, 114 Chiocchetti A 356 Chisholm SW 76, 86 Chiu SW 282 Cho S 360 Cho SW 356 Choi HS 325 Choi JN 37 Choi S 229, 278 Choi YD 37 Choo QL 38 Chou CH 428 Chow VTK 235 Chrétiennot-Dinet M-J 171 Christen R 233 Christensen M 323, 325 Christensen P 328 Christensen T 76 Christianson LM 363 Christophersen C 329 Churey JJ 330 Ciegler A 323 Ciesioka J 31 Cifelli R 169 Cigelnik E 234, 235, 361 Cimino MT 80 Ciniglia C 87, 116 Clark CA 358 Clark LG 429 Clarke L 229 Clear RM 354 Clegg MT 400, 428 Cliften P 278 Clifton SW 233 Cluck DR 237 Cobb FW 230

487

Author Index

Coelho KIR 326 Coenen A 331 Coffman AD 330 Coffroth MA 114 Cohen BA 278 Cohen-Bazire G 83 Cole GT 235 Cole WJ 454, 455 Coleman A 401, 403 Coleman AW 400, 401 Coleman DC 238 Collins AG 115 Collins MD 229, 278, 279, 358 Collins RA 30, 31, 33, 36, 37 Collins RF 31 Colombo AL 325 Colorni A 112 Colson I 278 Comas CI 403 Comerio R 331 Comes HP 403 Compos-Takaki GM 326 Concepcion GT 74, 110 Conesa A 329 Confalonieri VA 403 Conforti V 81, 82 Conn GL 401 Conraux C 326 Cook ME 78 Cook PE 323 Cooper R 332 Coppejans EGG 173 Cordo HA 477 Coriglione G 324 Corliss JO 76 Cornillot E 229 Correll JC 356 Corte AM 332 Cosson J 112 Cotty PJ 324 Couch JA 75 Coulson AR 84 Coutinho TA 356, 362 Couture LA 36

Cowperthwaite M 401 Cox CJ 233, 405, 406, 429-431 Cox DR 362 Cox EJ 176 Cox ER 76, 85 Cozzolino S 406 Cramer RA 356 Crandall-Stotler BJ 428, 452 Crane EH 382 Crane F 278 Crawford DJ 404, 405, 430 Crawford RM 170, 175, 176 Crayn DM 428 Crick FHC 32, 283 Croft JH 325, 326, 331 Cronk QCB 404 Cronn RC 401, 405 Crous PW 229 Culham A 360 Cunningham FX 86 Currah RS 230 Currie S 322 Cushion MT 229, 237 Czarnecki DB 169 Dacks JB 76, 79, 112 Dahlberg OJ 85, 115 Dahm SC 32 Dalcero A 324 Dale JL 355 Dallner G 282 Daly CB 326 Dams E 356 Dangel P 235 Daniel HM 278, 280 Dardé M-L 383 Darius HT 110 Daros JA 32 Dauga C 110 Daugbjerg N 76, 79, 110, 113, 170, 173 David JC 232 Davidson E 77 Davies J 327 Davila-Aponte J 83

488

Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

Davis EC 452 Davis JH 31 Davis RW 279 Dawes CJ 84 de Baer R 238 De Baere R 74, 283 de Barros Lopes M 475 de Beaulieu JL 404 de Berr C 239 De Graaff LH 324, 325 De Hoog GS 229, 231, 233, 236, 238, 240, 278 de Koning JRA 363 De la Peña M 32 de la Torre MA 279 de Morais PB 477 de Repentigny L 330 de Rijk P 86 De Salas MF 110 De Stefano M 170, 173 De Vries RP 324 De Wachter R 86, 238, 283, 356 de Waller J 326 Deak R 236 Deane JA 77, 80 Dearborn DG 239 Debets F 327, 331, 332 Debrunner-Vossbrinck BA 239 degli Ivanissevich S 383 Degrossi C 331 Degtyaryeva GV 456 Dehio C 32 Deichgräber G 84 del Tufo JP 362 Delabesse E 380 Deleault NR 32 Delneri D 278 DeLuca P 406 Delwiche CF 76, 80, 82, 83, 115 Demesure-Musch B 404 Deml G 235 Demoulin V 230 Denning D 327

Denning DW 323 Denniston KJ 38 Denny PW 80, 86 DePriest PT 230 D’Erchia A 401 Derelle E 111 DeRocher A 76 Derrick WB 32 Des Marais DL 401 deSantis-Maciossek G 428 Desikan A 278 Desjardins AE 238, 356 Dhardt DT 32 Di Battista C 356 Di Serio F 32 Diane N 401 Dieckmann T 33 Dierich MP 327 Diers BW 382 Dietrich F 239 Dietrich FS 229, 278 Dignani MC 361 Dimitrov K 236 Dinter-Gottlieb G 34, 37 Disney MD 429 Dixon MT 402 Dobrowolski MP 356 Dobson ADW 357 Dodd AP 475 Dodge JD 76, 111 Doebley J 428 Doermann AH 32 Doherty EA 32 Dohrmann J 430 Dolezel J 405 D’Ombrain MC 86 Dombrovska O 452 Donald RGK 86 Donaldson GC 356 Donis-Keller H 36 Donnetta AM 327 Donnini M 327 D’Onofrio G 111

489

Author Index

Donoghue MJ 231, 400 Doolittle WF 76, 79, 80, 86, 112, 227, 231, 283 Dooner HK 401 Dorey MW 231 Dörfler C 235 Dormer PG 330 Dorner JW 325 D’Orso I 84 Doster MA 323, 324 Doucette GJ 110 Doudna JA 32 Douglas AE 114 Douglas AW 429 Douglas SE 76, 77, 401 Dowd PF 332 Downing DL 330 Doyle JJ 399, 407 Drager RG 79 Draper DE 401 Drebes G 170 Drehwald U 454 Dressler DH 32 Driskell AM 402 Driver SE 33 Droop SJM 174 Drum RW 170 Druzhinina I 232 Dubois MP 355 Duchateau-Nguyen G 230 Duchense LC 358 Duffieux F 79 Dufresne A 77 Dugas M 452 Dujon B 228, 229, 278, 279 Dungan J 279 Dunham CM 32 Dunn-Coleman N 330 Durbin ML 428 Durley RC 324 Durnford DG 77, 78, 86 Durry E 327 Dyne K 327

Earnest TN 38 East PD 474, 478 Eathington SR 381 Ebbert MA 278 Ebizuka Y 329 Eckerlein B 235 Eckstein F 31, 37 Economou-Amilli A 170 Eddy SR 32 Edel V 356, 357 Edgar LA 170, 176 Edgar SM 170 Edlund MB 170 Edman JC 229, 234 Edwards SG 357 Edwards SV 380 Egea N 77 Egel DS 324 Eggink LL 77 Ehara M 171 Ehrendorfer F 429, 430 Ehrlich KC 324 Ehrman JM 172 Eichner RD 328 Eidell BR 231 Eilers RJ 429 Einax E 229 El-Kady I 324 El-Maraghy S 324 El-Minofi HA 329 El-Refai AMH 329 Elbraechter M 84 Elder JF 401 Elias KS 324 Eliceiri GL 32 Ellen R 111 Ellinger A 236 Ellmore GS 428 Elwood HJ 175, 229 Embley TM 77, 231 Endrizzi M 280 Eng C 357 Engelke DR 38

490

Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

Ennos R 404 Epplen J 360, 363 Epstein LM 32, 38 Erdner D 78 Erez T 476 Eriksson OE 229, 230, 232, 233, 240 Erlacher HA 35 Erlich HA 361 Ernster L 282 Espenshade MA 323 Esquível MG 456 Eujayl I 355 Evans EG 381 Exner LJ 281 Ezaki T 283 Fagan TF 77, 111 Fagard M 38 Fahrni JF 82 Falkowski PG 171 Faloona F 83, 233, 361 Fan J 33 Fang Z 85 Farmer VC 277 Farr DF 357 Fast NM 77, 111, 114 Fatima T 328 Faust MA 110 Federspiel NA 279 Fedor MJ 32, 34, 35 Feibelman TP 324 Feigon J 31 Fekete C 331 Feldberg K 454 Felden B 33 Feldmann H 230, 279 Feldstein PA 31, 37 Fell JW 232, 237, 277, 278, 280, 282 Felts D 328 Fennell DI 329 Fensome RA 111, 112 Ferbeyre G 32 Ferenczy L 329 Fernandez D 355, 357

Ferré D’Amaré AR 32, 36 Figge MJ 236 Figuera L 322 Figueras A 113 Fijisawa 406 Filgueiras TS 230 Filin VR 456 Filion M 363 Finch JT 36 Fineschi S 404 Fire A 33 Fischer E 454 Fischer SG 360 Fisher MC 239, 362, 383 Flaherty KM 36 Flannigan B 324, 327 Flavell RB 382 Flavier A 229, 278 Flinders J 33 Flo Jorgensen M 113 Flores R 32, 33, 35 Flossdorf J 279 Fogleman JC 475, 478 Folke C 171 Fonseca A 237, 278, 282 Fontdevila A 478 Fontinha S 456 Foreman DB 430 Forget L 229, 235 Forlani G 173 Forrest LL 428, 452 Forster AC 37 Fortas Z 355 Fortin JA 357 Fortini D 230 Föster H 357 Foth BJ 77 Fourie L 331 Fourier MJ 403 Fournier E 358 Fox GE 358 Fox MG 171 Fox RTV 360 Fradkin A 238

491

Author Index

Fraga S 113 Frahm JP 405, 431, 455 Francisco A de 171 Francisco-Ortega J 403 Franco JM 113 Franco MF 326 Frank DN 33 Frank JM 324, 329 Frank MR 475 Franks PJ 115 Franzot SP 230 Fraunholz M 77 Fravel DR 355 French FW III 171 Frey AL 33 Frey F 456 Frey ST 33 Frey W 405, 406, 430, 431, 455, 456 Fricke F 177 Friedel T 399 Friedl T 169 Friedlander TP 356 Friedman BA 239 Frijters A 362 Frisvad JC 324, 325, 328, 329, 330 Froenlich JE 76 Fryxell GA 171, 174 Fujimoto H 325 Fujita S 382 Fujita Y 79 Fujiu M 327 Fujiwara S 77 Fukagawa M 282 Fukatsu T 230, 279 Fukui K 404 Fukumasa-Nakai Y 231 Fukusaki E 279 Fukushima K 326 Fukuzawa H 404, 406 Fulkui K 406 Fulton B 278 Fulton L 278 Funes S 77 Fung E 77

Furman RM 230 Furukawa K 37 Gafa L 324 Gaffney TD 229, 278 Gago S 32 Gaiaschi A 323 Gaillardin C 228, 230 Galagan JE 230 Galibert F 279 Galitski T 236 Gall JG 32 Gallagher J 111 Galli CL 323 Galluzzi L 111 Gams W 229, 230, 232, 325, 355, 357, 363 Ganter PF 475, 477, 478 Gao XP 111 Garbelotto M 237 Garber RC 357 Garces E 111 Gardes M 357 Gardner C 170 Gardner MJ 77, 78, 86 Gareis M 359 Gargas A 230, 232, 236 Garibaldi A 356, 360 Garrett-Engele PW 232 Gasse F 177 Gast RJ 111 Gates K 229, 278 Gaucher GM 330 Gaut BS 400 Gautam SP 280 Gautheron N 357 Gauvreau H 237 Geiger HH 360, 361 Geiger J-P 355, 357 Geiser DM 230, 231, 239, 325, 330, 357, 362, 383 Geistlinger J 381 Geitler L 171 Gelfand DH 361 Gellatly DL 31

492 Gendron P 33 Gene J 325 Gené J 230 Geraud ML 113 Gerih JL 38 Gerlach W 357 Gernandt DS 401 Gershwin L 115 Gersonde R 171, 172, 175 Gessler C 383 Gesteland RF 33 Gherbawy Y 235 Gherbawy YAMH 357, 358 Ghignone S 356 Giammaria M 475 Gibas CFC 330 Gibbs SP 78, 81 Gibson A 475 Gibson RA 171 Gielkens MMC 325 Gilbert DG 475 Gilbert W 33 Gill F 38 Gillet R 33 Gillham, NW 78 Gillings M 381 Gillum AM 332 Gilson PR 78, 82 Giorni P 322 Giraud T 230, 358 Gissi C 401 Glasgow HB 81, 115 Glass NL 356 Gleeson MT 78 Glen AT 324 Glenn AE 230 Glenny D 453 Glezer ZI 171 Glöckner G 78 Gloer JB 332 Glusker DL 281 Gock MA 325 Gockel G 78 Godfrey JH 328

Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

Godhe A 175 Godoy P 325 Godwin ID 381 Goertzen LR 401, 403 Goetsch L 229 Goetz MA 327 Goffeau A 230, 279 Goffinet B 401, 405, 406, 428, 430, 431 Goheen DJ 230 Gold L 33 Goldman JC 171 Goldman N 77 Golenberg EM 428 Golinski P 323 Golubev WI 283 Gomes CM 477 Gomez MM 325 Gomi K 325 Gonzalez D 229 González U 176 Goodfellow M 278 Goodson MS 114 Gordon TR 227, 355, 356, 380 Goremykin VV 81, 401 Goris A 283 Goto T 326, 328 Gottschalk M 229 Gottsche ACM 452 Gottschling DE 34 Gottschling M 401 Gouka RJ 325 Grabowski PJ 34 Gradstein SR 402, 452-455, 457 Graham LE 78, 171, 235 Grajal-Martin MJ 358 Grammenoudi S 278 Granéli E 80 Grasby JA 38 Gräser Y 362 Grau CR 380-383 Gray J 237 Gray LE 380, 381, 383 Gray MW 75 Green BR 77, 78-80, 87, 112, 116

493

Author Index

Green DH 112 Greenwood AD 78 Gresshoff PM 356 Greuter W 230 Gribskov JL 322 Griffiths HB 78 Grimont PA 110 Grishok A 37 Gritton ET 382, 383 Grivet D 404 Grolle R 452-454, 456 Grossman L 358 Groth H 401, 402, 405, 453-455, 457 Growhurst RN 358 Grube M 230 Grum-Tokras V 33 Grzebyk D 111 Guadet J 358 Guadliana M 330 Guarro J 229, 230, 325 Gubler CJ 328 Gueho E 279 Guého E 233 Guerrier-Takada C 36 Guillard RRL 171 Guillebault D 111 Guillou L 113, 170, 171 Guldener U 228 Gullino ML 356 Gunde-Cimerman N 240 Günther F 171 Guo HC 33 Gupta RS 78 Gupta VS 355 Gusmao NB 326 Gustafsson P 232 Gutell RR 358, 401 Gutman GA 429 Gu ZJ 406 Gwilliam R 240 Haase G 230, 231, 238 Hachtel W 78 Hackett J 87

Hackett JD 74, 78, 79, 111, 116 Hadrys H 358 Hadziyannis SJ 33 Hagen CB 76 Hagler AN 279, 360, 476, 477 Hagopian JC 79 Hahn W 402 Hajós M 171 Hall BD 84, 85, 233 Hall GR 281 Hall N 77 Hallak C 324 Hallegraeff GM 110, 114, 115 Hallick RB 79, 428 Hallsteinsen H 176 Halmschlager E 231 Halpern AL 227 Hamaguchi M 113 Hamamoto M 279, 281 Hamari Z 325, 326, 330, 331 Hamdy AHA 329 Hamilton AJ 33 Hammann C 33 Hammann P 332 Hamory BH 327 Hampel A 31, 33 Hanley E 322 Hanlin RT 230 Hannaert V 79 Hannes AR 114 Hannukkala A 363 Hansel A 79 Hansen G 79, 110 Hansen HN 362 Hansen K 230 Hanson PM 381 Hanya Y 332 Hara S 325 Hargraves PE 171 Harper JT 79, 111 Harrington TC 231, 240, 381 Harris E 322 Harris SA 399 Harrison MA 325

494

Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

Hartl DL 278 Hartman GL 381 Hartmann FA 401, 402, 453, 457 Hartung U 281 Harwood DM 171, 172 Hase T 79 Hasegawa M 113 Haselkorn R 83 Haselkorn T 401 Hashimoto H 237 Hasle GR 172, 174 Hass H 239 Hässel de Menendez GG 453 Hastings JW 77 Hatefi Y 278 Hattori S 453 Hauben L 475 Haugland RA 238 Hauska G 235 Hausner G 231 Havanond S 428 Haware MP 355 Hawksworth DL 230, 231 Hawthorne BT 358 Hayes KA 238 Haywood AJ 111 He-Nygrén X 428, 453 Heale JB 358 Healy B 231 Heams T 37 Heard E 30 Heckman DS 231 Heckman JE 31 Hedenäs L 406, 429, 431 Hedgecock D 476 Hedges SB 231 Heed WB 475, 476, 478 Hegewald E 399 Hegg LA 35 Heinrichs J 401, 402, 405, 453-457 Heinrichs PJ 402 Heins L 79 Helbling A 231 Hellwig FH 401, 430

Helmchen T 74 Helms G 401, 453 Hendawy DS 228 Hendey NI 172 Hendriks I 356 Hennebert GL 231, 283 Henriksen P 76, 79 Henschke PA 475 Hensen O 322 Hensens OD 327 Henze K 79 Hepburn AG 381 Hepperle D 428 Hering O 358 Herman EM 110 Hernández C 33 Herrero E 279 Herrmann RG 428 Hershkovitz MA 402 Herth W 172 Herve G 37 Herzog M 114 Herzog T 455 Heslop-Harrison JS 402 Hess W 428 Hess WR 79, 83 Hester M 362 Heumann K 234 Hibberd DJ 79 Hibbett DS 229, 231, 239, 362 Hicks JS 75 Higgins KG 431 Higuchi R 361 Hiley SL 33 Hilger HH 401 Hill DRA 114, 176 Hillis DM 402 Hilu KW 428, 430, 455 Hilu W 400 Himes M 113 Himmler G 234, 360 Hinkle G 239 Hirano T 231 Hirata A 234, 282

495

Author Index

Hirooka K 279 Hirose T 428 Hirsch PR 355 Hirsch-Ernst KI 401 Hirshfield J 322 Hirt RP 77, 231 Hisashi I 85 Hishida H 330 Hjeljord L 231 Hobson LA 175 Hocking AD 325, 329 Hoef-Emden K 79 Hoek C van den 79 Hoekstra ES 326 Hoekstra RF 327, 331, 332 Hoffman C 322 Hoffmann B 33 Hofmann CJB 82, 86 Hoger R 362 Hogetsu T 113 Hoheisel JD 279 Hoiczyk E 79 Holley M 381 Hollingsworth PM 400 Hollins TW 382 Holmes MJ 112 Holst-Jensen A 231 Holtsford TP 400 Holz I 402, 453, 454, 455 Holzschu DL 476 Homma Y 355 Hommersand M 84 Honeycutt RL 406 Honeywill C 172 Hong L 79, 428 Hong SB 325 Hong TN 428 Hood EE 403 Hoogsteen K 322 Hoor-Suykerbuyk M 328 Hoot SB 402, 429, 430 Hopkin JM 235 Horak K 475 Hori H 279, 358

Horie Y 326, 330, 331, 332 Horiguchi K 331 Horiguchi T 112 Horio T 112 Horn BW 325, 328 Horn GT 361 Horner WE 231 Hornes M 362 Horrocks WDJ 33 Horvitz HR 36 Hou RF 278 Houbraken JAMP 324, 329 Houghton M 38 Houng JY 323 Houser-Scott F 38 Howard DH 279 Howarth DG 402 Howe AC 74 Howe CJ 110, 111, 112 Hoy JW 356, 381 Hsueh TG 38 Hu X 360 Huang CY 406 Huang HW 38 Huang KC 327 Huang S 428 Huang YS 38 Hubbard S 228 Hubbes M 358 Hudspeth M 358 Huff J 322 Hugall A 402 Hughes CE 399 Hughes CR 382 Hughes KW 402 Hughes TJ 381 Huitorel P 112 Hull CM 231 Humbert JF 380 Hung VKL 115 Hunt V 322 Hunter IL 278 Hunter N 476 Hunzikert JH 403

496 Hurtado LA 476 Hustedt F 172 Hutchins CJ 37 Hutchinson CR 326 Huttunen S 430 Hwang MJ 323 Hwu L 327 Hynes MJ 231 Hyun JW 358 Hyvonen J 361 Hyvönen J 363 Hywel-Jones NL 231 Ibl M 234, 236, 360 Ichinoe M 326 Idei M 174 Iglesias-Prieto R 110 Iitak Y 329 Ikeda M 325 Ilio de Dominicis R 403 Ilkiu-Borges AL 457 Inagaki J 79 Inagaki Y 79, 112, 171 Inokuchi H 404 Inoue H 455 Inouye I 86, 115 Ippolito A 400 Ishida K 79, 80, 85, 112 Ishida KI 115 Ishida T 80 Ishikawa H 279 Ishikura M 115 Isikawa Y 329 Isono K 429 Itakura S 113 Itezono Y 327 Ito M 402 Ito Y 231, 326, 328 Itoh M 281, 283, 326 Itoh TJ 112 Iwahashi H 77 Jackson CJ 381 Jacob Y 233

Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

Jacobs K 231 Jacobson DJ 239, 362, 383 Jacq C 279 Jaenisch R 38 Jahns HM 79 James SA 279, 358, 476 Jansen RK 401, 402, 403, 404 Janson S 80, 112 Jarv H 326 Jarvis P 80 Jeffrey SW 80 Jeng RS 358 Jenkinson P 361 Jensen KG 172 Jimenez-Diaz RM 358 Jimenez-Gasco MM 357, 358 John P 86 John U 112 Johnson AD 231 Johnson EA 328 Johnson JC 84 Johnson PJ 75, 115 Johnston M 278, 279 Jones D 277 Jones EW 455 Jones FD 33 Jones HM 172 Jones T 279 Jonsson P 231 Joosten V 326 Jorgensen R 35 Joshua H 322 Joyce GF 33 Burkholder JM Jr. 115 Juhász Á 331 Julien J 358 Juslén A 453 Kaboord BF 33 Kaczmarska I 172, 175 Kaden JE 476 Kadereit JW 403 Kaemmer D 363 Kahl G 363, 381

497

Author Index

Kaiser W 381 Kaji A 326 Kallas EG 325 Kalman S 279 Kanbe T 279 Kaneko T 80, 82 Kang S 357 Kangueane P 236 Kanzaki I 34 Kao R 327 Kardos NL 231 Karenina A 325 Karjalainen R 363 Karlsson-Borga A 231 Karol KG 80 Karpeisky A 35 Karuppayil SM 234 Kase N 332 Kasuga T 232, 238, 362 Kataoka A 328 Katayama Y 326 Katayama-Fujimura Y 279 Kato KH 112 Kato M 279 Katz ME 171 Kaufer NF 232 Kauff F 233 Kawachi M 110 Kawahara N 281 Kawai KI 328 Kawamura O 331 Kawamura Y 283 Kawano S 427 Kawasaki H 282, 283, 330, 479 Kawasugi S 328 Kaya HK 238 Kayamura T 360 Kayima S 331 Keegstra K 83 Keelin PJ 84 Keeling PJ 74, 79-81, 83, 84, 86, 111, 112, 114 Keese P 33, 37 Keiler KC 33

Keim P 382 Kejnovsky E 405 Kelch DG 402 Kelchner SA 402, 429 Keller B 229 Kellis M 280 Kellogg EA 405 Kelly A 358 Kendrew SG 326 Kenja J 332 Kennedy J 326 Keogh RS 232 Kerp H 239 Kerrigan J 280 Kerry BR 355 Kevei F 325, 326, 330, 331 Khalil MS 354 Ki-Hong C 172 Kiehn M 430 Kieslich K 326 Kikuchi A 279 Kilejian A 80 Kilham P 171 Kim HS 332 Kim YK 330 Kimbrough JW 228 Kindle KL 431 King J 85 Kinscherf TG 357 Kinsey JA 232 Kircher HW 476, 477 Kirisits T 235 Kiriyama N 326 Kirk PM 231, 232 Kishore R 80 Kiss I 331 Kissinger JC 77, 111 Kistler HC 235, 361 Kita Y 402 Kitaghawa S 329 Kitajima JP 79 Kitamoto K 325 Kjellberg F 383 Kjer KM 403

498

Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

Klassen GR 231 Klaubert DH 323 Klaveness D 115 Klein U 428 Klemsdal SS 363 Klich MA 323, 326 Klingberg M 81 Kloareg B 83 Klostermeier D 34 Klug A 36, 37 Knauf M 332 Knoll AH 171 Knoop V 400, 403, 405, 452 Koakutsu M 328 Kobayashi A 279 Kobayashi H 429 Kobayashi K 381, 383 Kobayashi T 329 Kobayashi Y 431 Koch A 360 Koch M 403 Kociolek JP 172, 173 Kodama K 234 Kodner R 236 Koenig H 326 Kofer W 428 Kofler G 327 Kogan N 359 Kogler H 332 Kohara Y 404 Kohchi T 404 Köhler S 80 Kohn LM 231, 380, 382 Koike K 85, 115 Koizumi M 34, 281 Komagata K 279, 281, 282 Komatsu Y 34 Kondo K 284 Kondo N 382 Kondrativea VI 476 König H 235 Kooistra W 110, 113 Kooistra WH 76 Kooistra WHCF 170, 173, 175, 177

Koop HU 428 Koptchinski A 232 Korthals HJ 75 Kosiak B 359 Koskinen S 363 Kostas SA 33 Köster J 176 Kotaki H 327 Kotani H 80 Koufopanou V 232, 330, 383 Koumandou VL 112 Kovács É 325 Kovacs JC 229 Kovacs W 232, 239 Kowallik KV 85, 112 Kowalski T 231 Kozakiewicz Z 322, 323, 325, 326, 328, 329, 331, 332 Kozlov G 84 Kozlowski M 358 Kramer CL 232 Krammer K 173 Krasske G 173 Kraus GF 232, 236 Krause K 427 Kreger-van Rij NJW 280 Krezy A 327 Krijger MC 355 Kroken S 239, 362 Kropf M 403 Kroth PG 80, 81 Kruger K 34 Krumbein WE 238 Krumlauf R 232 Krupinska K 427 Krzyosiak W 31 Kubicek CP 232, 239, 360, 363 Kubiek AR 363 Kuhls K 232, 239, 362 Kuhsel M 76, 82 Kuijpers AFA 324, 329 Kuiper M 362 Kuldau GA 357 Kullnig R 332

499

Author Index

Kullnig-Gradinger CM 363 Kumeda Y 326 Kun A 34 Kuo MYP 34, 37 Kupfer DM 232 Kupfer P 407 Kuraishi H 279, 326 Kuriyama A 175 Kuriyama H 230 Kurle JE 381, 382 Kuroiwa H 113 Kuroiwa T 404 Kuron G 322 Kurtzman CP 230, 232, 236, 279, 280, 282, 358-361, 476, 477 Kusters-van Someren MA 327 Kuzmine II 34 Kuzoff RK 405 Kwiecinska B 173, 177 Kwon-Chung KJ 277, 279, 280 Kydd GC 332 La Farge C 429 La Roche J 81 Laaser G 235 Laatsch T 80, 112 Lachance MA 280, 281, 475, 476, 477, 478 Laessøe T 230 Lafay JF 358 Lafontaine D 31 Lafontaine DA 34 Laforest MJ 235 Lagos-Quintana M 34 Laguerre G 356, 359 Lakatos M 173 Lambert R 332 Lammers JM 330 Lander ES 280 Landsberg J 112 Landvik S 232, 233 Lane DJ 228, 359, 403 Lanfranco L 359 Lang BF 81, 229, 233, 235

Lang M 81 Lang-Unnasch N 77 Lange-Bertalot H 173 Langridge P 475 Langseth W 359, 362 Languasco L 322 Lanoue KZ 403 Larsen J 110 Larsen TO 327 Lascoux M 404 Lasker BA 327 Lass-Florl C 327 Latgé JP 327 Latham BP 476 Lattanzi AR 355 Law CS 169 Lawlis JF Jr. 327 Lazarovits G 355 Le Guyader H 82 Lea A 328 Leadbeater BSC 78 Leander BS 81, 112, 114 Learn GH 400 Leatham T 116 Leblanc C 75 Lebruska LL 34 Leclerc JC 400 Leclerc MC 233 Lee J 380 Lee JA 74 Lee JJ 76, 170 Lee JS 37 Lee JY 38 Lee RC 34 Lee RE 81 Lee S 239, 363, 406 Lee SB 359 Legault P 33 Legrand A 114 Lehle L 235 Lehmann K 34 Lehmann PF 327 Lehmann R 37 Lehrer SB 231

500

Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

Lehtmaa J 326 Leigh J 229, 233 Lemieux C 35, 85 Lendeckel W 34 Leong SA 357 Lerch A 229, 278 Lerman LS 360, 362 Leroux P 230 Leskinen E 403 Leslie JF 382 Lester R 278 Levinson G 429 Levis C 230 Levy MG 112 Lewers KS 382 Lewin R 34 Lewin RA 81 Lewis BG 359 Lewis LA 81, 402 Lewitus AJ 81 Li C-W 173 Li JY 111, 112 Li N 283 Li TM 283 Li WH 429 Li Y 34 Li YF 406 Li ZY 283 Lia VV 403 Lian C 113 Liang WQ 403 Liaud M-F 81 Lichtenthaler FW 277 Lieberman K 38 Liebl S 234 Lieckfeldt E 232, 239, 360, 362 Liesch J 322 Liesch JM 327 Ligon JM 355 Lila Koumandou V 111 Lilley DMJ 33, 34, 38 Lin A 327 Lin D 327 Lindenberg JBG 452, 455

Linder CR 403 Lindlar A 431 Lindner M 402, 453, 454, 455 Ling KH 323 Linton EW 81 Lipari SE 360 Lippmeier JC 74 Lipson DA 237 Liston A 401, 403 Litaker RW 112 Liu JS 403 Liu YJ 233 Liu YL 233 Liu ZL 359 Livak KJ 363 Llorente B 228, 233 Lobanok AG 329 LoBuglio KF 233, 325, 359 Lockau W 235 Locke T 328 Loeblich AR 110 Loeffler W 234 Löffelhardt W 81, 84, 85 Logan KS 282 Logrieco A 359, 361 Lohse PA 37 Loiseaux-De Goër S 83 Lokman C 326 Lomer CJ 355 Long DG 430, 453 Longcore J 235 Longet D 74, 81 Longsdon JM 231 Lonial S 327 Lopandic K 233, 235, 236, 237 Lopez M 322 Lopez-Diaz TM 327 Lopez-Garcia P 112 Loppnau PA 233 Loret P 113 Louis EJ 278, 279, 476 Lowe DA 327 Lowen R 236 Lozano J 111

Author Index

Lu S-D 380 Lucassen RW 32 Ludwig M 81 Luedi P 229, 278 Luisetti M 327 Lumbsch HT 233 Lundholm N 173, 174 Luo YL 323 Luthy J 322 Lutzoni F 233, 403 Lynch M 75 Lyne R 233 Lysák MA 405 Ma DP 428 Mabry ML 429 Mach R 363 Macmillan J 324 Maddox JV 239 Maeda H 37 Maeda K 237, 284 Magee BB 279 Magee PT 279 Maggini F 403 Magnani M 111 Magnoli C 324 Maher A 228 Mahoney NE 323 Mahood AD 169 Mai JC 401, 403 Maier RM 428 Maier U 74 Maier UG 86, 112, 115 Majer D 359 Major F 33 Majors J 278 Makalowska I 357 Makarova IV 171 Malassezia Baillon 277 Malcomber ST 403 Maliga P 427, 429, 430 Malloch D 228 Maloney L 35 Malpertuy A 233

501 Malvick DK 382 Manabe M 328 Mandel M 281 Manen J-F 429 Mangnus J 329 Manhart JR 404 Maniatis T 360 Manley JL 37 Mann DG 79, 169, 170, 173-176, 178 Mann DG 174 Manos PS 404 Manulis S 359 Manzoni M 327 Maranda L 78, 111 Marasas WFO 238, 359, 361 Maraz A 236 Marchfelder A 35 Marciniec T 31 Marcon MJ 359 Marger N 359 Marienfeld JR 406 Marin B 79, 81 Marin I 113 Marinets A 76 Marino D 111, 113, 170 Markley JC 31 Markos S 399 Markow TA 476 Marlowe JL 278 Marmur J 281 Marr KA 322, 327 Marroco R 403 Martin AP 237 Martin F 356 Martin JP 404 Martin PM 110 Martin SJ 31 Martin W 81, 406, 456 Martin WF 456 Martinez del Pozo A 327 Martinez-Ruiz A 327 Maruyama T 85, 110, 115 Marzachi C 359 Marzluf GA 232

502

Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

Masclaux F 233 Massana R 113 Massey AP 36 Masur H 229 Mata J 233 Matheny PB 233 Mathews DH 429 Matsukuma S 327 Matsumoto S 406 Matsuyama Y 113 Matsuzaki M 82, 113, 404 Matthäi A 239 Matthews M 360 Maurais SC 401 Maurel MC 34, 37 Maxwell P 357 May G 362 Mayali X 115 Mayama S 175 Maynard Smith J 382 McCain AH 356 McCarroll R 360 McCarthy PJr 327 McClelland M 363 McCourt RM 80, 81 McCowen MC 327 McDaniel SF 429 McDonald SM 174 McEwan ML 114 McFadden GI 77, 78, 80, 82, 113 McGovern PE 278, 281 McGuire P 113 McKay DB 36 McLoughlin S 455 McNeill J 230 McQuoid MR 175 Medlin L 74, 175 Medlin LK 110, 112, 169-173, 175, 177 Meehan BM 362 Megnegneau B 327 Mehrotra B 401 Melkonian M 74, 79 Mello CC 33, 37

Melzer Krick B 230, 231 Menden-Deuer S 114 Mendonca-Hagler LC 360, 476, 477 Mendoza L 234 Meng X 382 Meng XQ 380 Mengistu A 382 Mereschkowsky C 82 Mergaert J 475 Messner K 362 Messner R 231, 234, 235, 360 Mesterházy Á 329 Metenier G 229 Meusel O 362 Mewes HW 228, 234, 279 Meyer EG 113 Meyer W 232, 278, 280, 360, 362 Miao VP 86 Michaels HJ 404 Michailides TJ 323, 324, 328 Michalowski C 85 Migheli Q 356, 360 Mikata K 281, 284, 479 Mikhailova RV 328 Milanovic M 33 Milgroom MG 360, 382 Millar DP 34 Miller MW 281, 282 Miller MWHJ 476 Miller WI 175 Millie D 402 Mills PR 362 Mims CW 227, 322 Minguez S 323 Minnikin DE 278 Minute A 356 Miranda M 281, 474, 476, 477 Mironenko NV 476 Mirza JH 328 Mishima M 404 Mishler B 402 Mishra PK 360 Misset MT 400

503

Author Index

Misumi O 113, 404 Mitchell A 356, 360 Mitchell GT 33 Mitchell TG 362 Mithen R 359 Mitsuhashi W 237 Mitter C 356, 360 Miyadoh S 332 Miyagishima S 404 Miyaji M 326, 332 Miyamoto K 326 Miyashita H 85 Moawad MK 228 Moeller M 430 Moestrup O 82, 110 Mohanty A 404 Mohmed IN 354 Mohr C 229, 239, 278 Molano J 281 Moldowan JM 113 Möller EM 360, 361 Moller M 404 Molnár J 331 Molnár O 233, 235-237 Monaghan R 322 Montalbano BG 324 Montegut-Felkner AE 82 Monteiro RF 477 Montgomery MK 33 Montgomery RT 175 Montresor M 111, 113 Moore ER 475 Moore RT 281 Moore TDE 234 Morais PB 476, 477 Moran VC 476 Morawetz R 360 Morden CW 82 Moreau H 113 Moreau RA 281 Moreira D 82, 112 Morgan DP 328 Morgan DR 404

Morgante M 383 Moritz C 402 Moriyama A 112 Morl M 35 Morse D 77, 82, 113 Mort ME 404, 429 Mortimer R 278 Morton BR 400 Mossink MH 38 Moukhamedov R 360 Muchhal U 85 Muchhal US 80 Muchlbauer FJ 358 Mueller UG 360 Mues R 453 Muhle H 400, 452 Mukherjee PK 356 Mukhtar T 78 Mulé G 323 Mullaney EJ 326 Müllbacher A 328 Mullenbach GT 38 Müller E 234 Müller J 455 Müller K 429, 430, 455 Muller KM 82 Müller O 175 Muller-Starck GM 404 Mullis KB 234, 361 Mullis KD 361 Munch J-C 356 Munholland J 82, 83 Munsterkotter M 228 Murakami Y 279 Murata K 382 Murchie AI 35 Murphy CA 76, 77 Murray D 430 Murray JB 32, 35, 37 Murray PR 234 Murray S 113 Murthy NBK 356 Musza LL 332

504

Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

Muyzer G 177, 360 Myers RM 360, 362 Naaber P 326 Nagahama T 234 Nagai S 113 Nagakawa S 406 Nagatsuka Y 479 Nagumo T 177 Najarian RC 38 Nakagawa Y 284 Nakahara H 114 Nakajima M 331 Nakamura Y 80, 82 Nakase T 235, 238, 278, 279, 281-283 Nakashima N 281 Nakayama N 327 Nakayama T 112 Namiki F 360 Nanney DL 113 Nannfeldt JA 234 Napoli C 35 Nash G 114 Nassal B 405 Nassoury N 82, 113 Nasu S 234 Natali A 429 Natvig DO 228 Naumov GI 476 Naumova ES 476 Navarro B 35 Naveau F 239 Nazar M 328 Nazar RN 360 Neefs JM 283, 356 Nees ab Esenbeck CG 452 Negishi H 381 Neinhuis C 400, 428 Nelissen B 283 Nelson MA 228 Nelson PE 359, 361 Nerad T 81 Nesbitt SM 35 Neto PRP 477

Neuveglise C 233 Neuvéglise C 228 New DC 115 Newell ND 176 Newman A 35 Newport G 279 Ng TB 328 Nicholson P 356, 361, 383 Nickell AD 380 Nickell CD 380, 381, 383 Nickerson DA 380 Nickle D 322 Nicklen S 84 Nickrent DL 404 Nicolson DH 230 Niederwieser D 327 Nielsen PV 329 Niemann C 360 Nikolaev SI 82 Nikolaev VA 172 Nilsen TW 31 Nilsson C 176 Nimer ND 328 Nirenberg HI 234, 357, 358, 362 Nirenberg I 357 Nisbet R 111 Nisbet RE 112 Nishi K 360 Nishida H 234, 238, 281, 282 Nishihara S 406 Nishimura K 326, 332 Nishiyama R 406 Niwa Y 429 Nixon KC 429 Noda H 234, 237, 281, 282 Noga EJ 112 Noguchi H 329 Noller HF 38 Nomura M 326 Norman DG 34 Norris RE 79 Norry FM 477 Novey HS 328 Nowell W 227

505

Author Index

Nozaki H 82, 113, 404 Nozawa K 328 Nudelman MA 81, 82 Nunez A 281 Nwakanma DC 404 Obata R 330 Oberwinkler F 228, 235 O’Brein PA 356 O’Callaghan J 357 Ochoterena H 430 O’Dell M 382 O’Donnell K 234, 235, 237, 238, 354, 355, 357, 361 Oestreich SC 31 Ognjanova-Rumenova N 176 Ogoshi A 381, 382, 383 O’Grady E 328 Ohama T 112 Ohmido N 404, 406 Ohno S 281 Ohta N 82, 404 Ohtsuka E 34 Ohtsuka T 327 Ohyama K 404, 406 Okada G 281 Okada K 85, 326 O’Kelly C 81 Okoli BE 404 Oldach DW 115 Oldach WD 115 Oliphant JM 403 Oliveira MC 82 Oliver RP 359 Oliver SG 235, 278, 279 Oliveri S 324 Olmstead RG 177 Olsen GJ 281, 359, 360, 403 Olson RJ 76 Omura S 330 O’Neill NR 355 Onions AHS 325 Ono H 328 Onofrio GD 113

Ooijen AJJ 324 Ooike M 328 Oplinger ES 380 Ordas A 113 Ordas M 113 Orgel LE 35 O’Rourke M 382 Orozco EM 429 Orsini L 176 Osafune T 84, 86 Osawa S 279, 358 Ostrowski M 327 Ostry V 328 Otis C 85 Otrosina WJ 237 Ou JH 38 Ouinten M 357 Oury B 383 Oyaizu H 281 Ozeki H 404, 406 Ozeki M 279 Paasche E 176 Paavanen-Huhtala S 361, 363 Pace B 359, 403 Pace NR 33, 281, 359, 403 Padhye AA 324, 327, 328 Page RDM 404 Palacio G 324 Palágyi A 331 Palenik B 83 Palm A 81 Palme A 404 Palmer JD 76, 83, 227, 404 Pan S 235 Panaud O 407 Panella L 356 Pang WM 281 Papes D 407 Paquin B 235 Parenicova L 328 Parikka P 363 Pariza MW 328 Park C 326

506 Park W 401 Parnell KM 36 Parry DW 361, 383 Parsons SJ 330 Partensky F 77, 79 Pasquinelli AE 36 Pasturenzi L 327 Pataky JK 381 Patchett A 322 Paterson E 404 Paterson RRM 328 Patriarca A 331 Patron NJ 83 Patterson DJ 75, 83 Patterson N 280 Patterson T 327 Patzoldt ME 382 Pauillac S 110 Paulus EF 332 Pawlowski J 74 Payne GA 329 Payri C 173 Peacock W 328 Pearce AR 324 Pearce DA 355 Pearson CAS 382 Peay KG 237 Peculis B 404 Pedersen N 429 Pedros-Alio C 112, 113 Pegg KG 355 Pegler DN 231 Peleman J 362 Pemberton RW 477 Pendola J 475 Peng CI 428 Peng FC 323 Penna A 111 Penno S 83 Penny SL 77, 401 Peona V 327 Perasso R 177 Perez-Artes E 358 Pérez-Artés W 358

Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

Pérez-Martínez X 83 Peris F 475, 478 Pernfuss B 235 Perreault JP 30, 31 Perrone G 323 Perrotta AT 31, 36 Pesole G 401 Peter B 238 Petersen RH 402 Peterson SW 235, 279, 323, 326, 328, 361 Petit RJ 404 Petrini O 232 Pfaller MA 234 Pfeifer F 234 Pflug IJ 325 Phaff HJ 278, 281, 282, 474-478 Phalen DN 282 Philipe H 82 Philippe H 233 Philippsen P 229, 238, 278, 279 Piccioni PD 327 Pickett-Heaps JD 83, 85, 169, 170, 176 Pietri A 322 Piippo S 428, 453, 455 Pillay M 404 Pin LC 113, 115 Pinero D 401 Pinner RW 327 Pinsker W 233, 430 Pinto G 87, 116 Pinto VF 331 Piola F 405 Pirseyedi M 228 Pirzadeh BS 330 Pitt JI 230, 233, 235, 325, 329, 359 Pixley FJ 235 Pley HW 36 Ploetz RC 235, 361 Pocock KL 176 Pócs T 454, 455 Podbielski A 230 Pöder R 235 Poggeler S 329 Pöhlmann R 229, 239, 278

507

Author Index

Polsinelli M 278 Pons A 323 Poole RW 356, 360 Poot GA 280, 282 Pore RS 282 Porter JM 400 Pot J 362 Potashkin J 232 Potter D 175 Poulos PG 325 Powell DA 359 Powers DA 113 Prade RA 232 Prado G 322 Prasad GS 280 Preiser DJ 86 Preiser P 86 Preisfeld A 75 Preparata RM 113, 401 Presber W 362 Price R 230 Price RA 399 Prillinger H 231-237, 357, 360, 361 Prinz S 236 Prior C 355 Procaccini G 113, 176 Pröchold T 402 Prody GA 36 Pröschold T 454, 455 Pruchner D 400, 405 Pryer KM 86, 401 Punt PJ 325, 326, 329 Qi ZT 330 Qiu YL 452 Quandt D 405, 406, 428, 430, 431, 455, 456 Quarles B 475 Queller DC 382 Quinn CJ 428 Radwanski ER 431 Rafalski JA 363 Ragan MA 83

Ragozzino A 32 Rahayu ES 330 Rahbaek L 329 Rainieri S 475 Rajandream MA 240 Ramakrishna NV 332 Rambold G 230 Ranieri RL 323 Ranjekar PK 355 RAPD–PCR 361 Raper KB 329 Rasmussen TP 38 Rassmann K 382 Rastogi T 36 Rath P 327 Rathjen PD 37 Rauhut R 34 Raven PA 83 Raymond O 405 Razafimandimbison SG 405 Redecker D 235 Redhead SA 235 Reeb V 403 Reece CA 232 Reed RE 36 Reel DC 84 Regier JC 356, 360 Rehner SA 236, 382 Reid J 231 Reijans M 362 Reiner-Drehwald ME 452 Reinhart BJ 36 Reis M 79 Reith M 82, 83 Remy W 239 Rendell S 404 Renker C 402, 454, 455 Renner SS 430 Rensing SA 86 Restani P 323 Retallack GC 236 Reuman S 83 Reuven M 359 Revoy F 359

508

Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

Reyes-Prieto A 77 Reynolds DR 236 Rezanoor HN 356, 361, 383 Rhode K 173 Richards MP 281 Richardson B 113 Richly E 81 Rieth H 236 Rigó K 329, 331, 332 Rikkerink EHA 358 Riley J 85 Rines JEB 176 Rinyu E 329, 330, 331 Rippka R 83 Ritz CM 430 Rivera A 355 Rivera P 176 Rizzo DM 231 Rizzo PJ 83 Robb J 360 Roberts IN 229, 278, 279, 476 Roberts KR 114 Roberts LN 358 Roberts M 110 Robertson DL 86 Robertson HD 31 Robinow CF 277 Robinson WA 403 Robnett CJ 232, 280, 359, 476 Rocap G 79, 85 Rodriguez N 232 Rodriguez-Valera F 112 Roe BA 232 Roeijmans HJ 236, 282 Roewer I 235 Roger AF 74 Roger AJ 74, 83, 84, 115, 227 Rogers CR 74, 110 Rogers JD 280 Rogers MB 83 Rogerson CT 236 Rolfsen W 231 Rollini M 327 Roos DS 77, 111

Ros RM 405 Rosa CA 476, 477 Rosa CAR 324 Rose F 455 Rosenstein SP 31, 36 Rosenthal A 78 Roshchin AM 176 Rospondek MJ 176 Rossen L 328 Rossi JJ 36 Rossi MS 82 Rossman AY 236, 357 Roth LE 171 Roth WB 428 Rothpletz A 176 Rothrock J 322 Rougvie AE 36 Round FE 176, 178 Rowan R 113 Rudi K 115 Rüdiger W 428 Ruffner DE 36 Ruiz-Gonzalez MX 113 Rujan T 81 Rundberget T 359 Rupe JC 381 Rupert PB 36 Rutherford MA 355 Ruvkun G 36 Ryburn JA 405 Rycroft DS 402, 405, 454, 455 Ryder SP 36 Ryocroft DS 402 Saavedra E 79 Sabbe K 170, 178 Sabourin M 358 Saccone C 401 Saenz GS 236 Sagbohan J 355 Saikawa M 238 Saiki R 361 Saiki RK 83 Sainani MN 355

509

Author Index

Saint-Louis D 75 Saito R 326 Sakharkar MK 236 Sako Y 111, 114 Salais MF 114 Salanoubat M 77 Saldarriaga JF 84, 111, 114 Salehi-Ashtiani K 36 Salkin IF 230 Sallam LAR 329 Salma N 173 Salvaggio JE 231 Samigullin TH 456 Sampaio JP 278 Samson RA 235, 323-329 Samuel R 430 Samuels GJ 232, 236, 239, 382 Sanderson MJ 400 Sands J 34 Sang JH 477 Sang T 405, 430 Sanger F 84 Sankawa U 329 Sanlaville-Boisson C 405 Sano T 327, 404 Santi DV 229 Santore UJ 78 Santos M 34 Santos SR 114, 477 Sapunova LI 328 Sarno D 176, 177 Sasa T 115 Sasorith S 111 Sato H 110, 112, 230, 234 Sato S 80, 82, 177, 331 Sato, Y 110 Satoh K 429 Sauer M 454 Sauer RT 33 Saunders GW 84, 114 Savage AM 114 Savile DBO 236 Saville BJ 36 Savonna C 81

Sawaguchi T 115 Saxena M 363 Schaal BA 400, 428 Schadt CW 237 Schardl L 403 Scharf S 83, 361 Scharf SJ 361 Scheetz TE 111 Scheidegger KA 329 Schell J 32 Scheper RJ 38 Scherer S 279 Scherzinger M 81 Scheuer C 235 Schiewater B 358 Schiffner V 456 Schill DB 430 Schilling AG 360, 361 Schimmel TG 330 Schimper AFW 84 Schindler M 405 Schlag-Edler B 477 Schlatter C 322 Schlick A 362 Schlötterer C 382 Schmedicke RA 478 Schmid A-MM 172, 175-177 Schmidt A 177 Schmidt M 177 Schmidt SK 237 Schmidt U 34 Schmitt R 282 Schmull M 453 Schneider EM 114 Schneider IR 36 Schneider RW 356, 381 Schnepf E 84, 114 Schoch S 428 Scholin CA 114, 115 Schönian G 362 Schroeder SJ 429 Schuette KP 404 Schuettpelz E 430 Schulz A 75

510

Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

Schumacher T 231 Schumacker R 453 Schupp JM 382 Schuster E 330 Schuster RM 455, 456 Schütt F 177 Schwan RF 322 Schwantes HO 237 Schwartz HF 356 Schwartzbach SD 80, 84, 86 Schweigkofler W 234-237, 239, 361 Schwenk FW 382 Scorzetti G 237, 282 Scotland RW 177 Scott DB 331 Scott JA 239 Scott WG 32, 35-37 Screen SE 330 Sears BB 431 Sebastian SA 381 Seelanan T 405 Sehgal OP 31 Seif E 233 Seila AC 36 Seki T 479 Sekiguchi J 330 Selker EU 232, 237, 357 Semina HJ 177 Sengco M 82 Seoighe C 232 Serra R 323 Sexton JP 114 Seyhan AA 35 Sgrosso S 113 Shailer NFJ 233 Shalchian-Tabrizi K 85, 115 Shank RC 323 Shankle AM 115 Shann C 282 Sharada K 362 Sharmeen L 34, 37 Sharp PA 37, 38 Shaw AJ 404-406, 428-431 Shearer TL 114

Sheath RG 82 Sheffield VC 362 Shen R 477 Sherkhane PD 356 Sherwood MA 237 Sherwood-Pike MA 237 Shi X 381 Shih I 36 Shih IH 37 Shiki Y 404 Shimamoto T 330 Shimazu M 234, 237 Shimma N 327 Shin C 37 Shin HD 325 Shin-i T 404 Shiomi T 360 Shirai H 327, 404 Shlacht S 360 Shoemaker RC 382 Shu S 283 Siddiqui N 84 Siegel AF 236 Sieminska J 177 Sigler L 235, 237, 239 Sigurdsson ST 31, 36 Silberman JD 115 Silbermann JD 84 Silhavy D 430 Siljak-Yakovlev S 400, 407 Sills GR 383 Silva PC 230 Sim-Sim M 456 Simmonds FJ 477 Simmons MP 430 Simmons RB 277 Simon CJ 358 Simon EM 113 Simon L 229 Simon LD 427 Simonsen R 177 Simpson AG 84 Simpson DR 383 Simpson GE 172, 174

Author Index

Simpson TJ 324 Sims PA 175 Sinclair JB 381 Singh B 78 Singler H 113 Sinninghe-Damsté JS 176, 177 Siroky J 405 Sjamsuridzal W 282 Skaida M 406 Skog JE 230 Skouboe P 325, 328 Slack FJ 36 Sluiman HJ 174 Small RL 401, 405 Smedsgaard J 325, 327 Smetacek V 171, 177 Smith AB 330 Smith AR 401 Smith EM 354 Smith JM 32 Smith LW 381 Smith ML 362 Smith MT 280 Smith MTh 282 Smith NH 382 Smith SL 237 Smits G 356 Smol JP 177 Smolich BD 362 Snowden KF 282 Snyder WC 362 So ML 453, 456 Soares MB 111 Sogin ML 84, 114, 229, 239, 359, 360, 403 Solga A 454 Solignac M 358 Soliman MH 478 Soll J 75, 79, 80 Soltis DE 400, 404-406, 429 Soltis PS 400, 404-406, 429 Someya A 330, 332 Someya J 77 Sone T 406 Song JT 37

511 Song SI 37 Sonneveld P 38 Sonntag L 230, 231 Sood VD 33, 37 Sorenson WG 239, 282 Sörhannus U 177 Sörhannus UM 171 Sorrell TC 278 Souciet JL 237 Sournia A 177 Soyer M-O 115 Soyer-Gobillard MO 113 Spagnuolo V 406 Spata G 324 Spatafora JW 237 Spencer DF 77 Spittstoesser DF 330 Spratt BG 382 Springer J 322 Spruce R 456 Squirrell J 430 Sreenivasaprasad S 362 Srinivasan A 236 St Germain G 330 St Arnaud M 363 St Leger RJ 330 Stahl DA 359, 403 Stahl YD 360 Stalpers JA 232 Stanton J 402 Stapley E 322 Starmer W 478 Starmer WT 474-478 Starr EM 404 Statzell AC 278 Statzell-Tallman A 282 Staudacher E 236 Stauffer RL 231 Stchigel A 325 Stchigel AM 230 Stech M 405, 406, 430, 431, 455, 456 Stechmann A 84, 237 Steele RH 478 Steenackers M 475

512

Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

Steenkamp ET 238, 362 Steidinger K 112 Steidinger KA 84, 111 Steimel J 381 Steinberg C 356, 357 Steiner JM 84 Steiner S 229, 239, 278 Steinhauser S 452 Stella G 324 Stephani F 456 Stephens PA 380 Stepien PP 358 Sterflinger K 236, 238 Stern DB 431 Sterrenburg FAS 173 Steven E 330 Stevenson JK 428 Steyn PS 331 Stibitz TB 84 Stickel S 175, 229 Stickel SK 239 Stickle AJ 172, 174 Stiller JW 84, 85 Stirewalt V 85 Stoddard BL 37 Stoebe B 81, 85, 115 Stoebe-Maier B 80, 112 Stoecker DK 113 Stoecker KD 115 Stoermer EF 170, 172, 173, 177 Stoffel S 361 Stormo GD 36 Stosch HA 178 Strassmann JE 382 Strauss J 239 Strel’nikova NI 178 Stringer JR 238 Strobel SA 33, 36 Stuessy T 430 Stuessy TF 405 Su V 86 Suda S 86, 115 Sudarsanam P 278 Suga H 37

Sugita C 431 Sugita M 431 Sugita T 238 Sugiura C 431 Sugiura M 428 Sugiura Y 331 Sugiyama J 80, 228, 234, 238, 279, 281, 282, 326, 330 Suh SO 238, 281, 282 Sulli C 85 Sullivan DJ 238 Summerbell RC 326, 330 Sun ZM 330 Sunazaka T 330 Sundback K 176 Supattapone S 32 Sutton BC 231, 238 Suzuki M 233, 238, 281 Svendsen A 327 Swamy KH 332 Swann EC 238, 239, 282 Swiezewska E 282 Swings J 475 Swofford DL 431 Symington HA 112 Symons RH 33, 37 Syvertsen EE 172 Szakacs G 232 Szaniszlo PJ 234 Szaro TM 229, 355 Szathmary A 34 Szencz S 331 Szostak JW 31, 36, 37 Tabara H 37 Tahvonen R 363 Taira K 37 Tajiri Y 282 Takada M 330 Takahara M 82, 113 Takahashi H 326 Takahasi H 332 Takaki GMC 326 Takeda Y 115

513

Author Index

Takematsu A 281 Takenada M 406 Takeo K 238 Takeuchi M 404 Takishita K 85, 115 Takishitam K 115 Talyzina NM 113 Tamura M 330 Tan CS 236 Tan S 77 Tanabe Y 238 Tanada Y 238 Tanaka A 85 Tanaka J 177 Tanaka K 328, 404 Tanaka NK 85 Tanaka R 85 Tang J 281 Tang RH 406 Tangney RS 405 Tanner NK 37 Tantaoui A 357 Tarlo SM 238 Tatum E 228 Taufratzhofer E 234 Tausch I 362 Tautz D 382 Tayler JM 37 Taylor F 114 Taylor FJ 114 Taylor FJR 84, 85, 111, 114, 115 Taylor J 34, 37, 363, 406 Taylor JW 228-230, 232, 233, 236, 238, 239, 277, 282, 325, 330, 356, 357, 359, 362, 380, 383 Taylor TN 239 Taylor WC 402 Teen LP 113, 115 Tehler A 230, 239 Tekaia F 233 Tekaia F 230 Temmis JN 406 Templeton MD 358 Tengs T 85, 113, 115

Tenkouano A 404 Tenover FC 234 Tenzer I 383 Teo SLM 112 Téren J 330, 331, 332 Teresa Gelati M 403 Terwey DP 35 Tester PA 112 Tettelin H 279 Thanh VN 279, 282 Thayer RM 38 Theriot EC 170, 173, 176, 178 Theron JJ 331 Thien LB 427 Thomson JB 35 Thorsteinsen KE 75 Thorstenson YR 279 Thuan TB 282 Tibayrenc M 383 Tietz H-J 362 Ting CS 79, 85 Tingey SV 362, 363 Tinoco IJ 31 Tippit DH 85 Tiu C 323 Tlachac K 235 Tobe S 37 Tobin RS 238 Tomas RN 85 Tomaszewski EK 282 Tomitani A 85 Tomoda H 330 Torp M 362 Torres RA 37 Tóth B 329, 331, 332 Tournas V 330 Toussoun TA 359, 361 Toyazaki N 330 Trapido-Rosenthal H 114 Trappe JM 234 Tredick J 476, 477 Tredick-Kline J 477 Trehane P 230 Trench RK 110

514 Triemer RE 82 Troitsky AV 456 Tronsmo A 231 Trop M 362 Truby EW 84, 111 Tsubouchi H 330, 331 Tsuda M 239 Tsuge T 360 Tsuneda A 231 Tsuzaki N 279, 326 Turgeon BG 357 Turland NJ 230 Turmel M 85 Turner BJ 401 Turner D 239 Turner DH 429 Turner S 86 Turner WB 324, 330 Tuschl T 34, 37, 38 Tzean SS 327 Uchida A 85 Uchiyama S 331 Udagawa S 326, 330-332 Udagawa SI 328, 332 Uebayasi M 37 Ueda Y 406 Uemura T 283 Ueno Y 328, 331 Ueyama A 239 Uhlenbeck OC 32, 36, 37 Uijthof JMJ 230 Umesono K 404, 406 Umile C 235 Unseld M 355, 406 Unterreiner WA 239 Urbach E 86 Usman N 35 Usup G 113, 115 Vaamonde G 331 Vacek DC 478 Vagi M 114 Vaillancourt RE 110

Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

Valadkhan S 37 Valentin E 279 Valentin K 78 Valiehoroman KM 456 Valiejo-Roman K 456 van Berkum P 355 van Biezen N 329 van de Lee T 362 Van de Peer Y 86, 283, 356 van de Vondervoort PJI 324 van den Broeck HC 324 van den Hondel C 329 van den Hondel CA 325, 326 van der Aart QJ 235 Van der Auwera G 74, 86 van der Merwe KJ 331 Van der Peer Y 230, 400 Van der Schoot J 400 Van der Staay GW 79 van der Staay GWM 81 Van der Walt JP 283 van Dijck PWM 330 Van Dooren GG 86 Van Dreven F 355 Van Dyk MS 282 van Eijk GW 282 Van Helden J 228 van Tol H 37 van Wyk PS 229 Van Zon A 38 Vanden Berghen C 457 Vanden Heuvel B 403 Vandenbempt I 356 Vanderpoorten A 406, 431 Varga J 325, 326, 329-332 Vaucheret H 38 Vauterin L 475 Vautrin D 358 Vázquez-Acevedo M 83 Vederas JC 326 Veenhuis M 75 Veeraraghavan N 357 Venancio A 323 Vendramin GG 404

515

Author Index

Vercken E 358 Verdonck L 475 Vergne J 37 Verreet JA 354 Vertesy L 332 Vesonder RF 332 Vesper SJ 239 Vida Z 332 Vijayakumar EK 332 Vijg J 357 Vila M 111 Vilardi JC 477 Vilgalys R 229, 362 Villareal TA 113 Visconti A 359 Visram S 114 Visser C 239 Visser J 323-325, 327, 328 Visvader JE 37 Vitt DH 429 Vivares CP 229 Voegeli S 229, 278 Vogel HJ 239 Voigt K 229, 239 Volcani BE 170, 173 von Arx JA 283 Vos P 359, 362 Vosman B 400 Vossbrinck CL 239 Vujanovic V 363 Vyskot B 405 Vyverman W 170, 178 Waalwijk C 355, 362, 363 Wagner D 431 Wagner P 403 Wainright PO 239 Wakazawa M 328 Wakefield AE 235 Walker WF 239, 283 Waller PRH 33 Waller RF 82, 86 Walsby AE 178 Walter F 35

Walter NG 35 Walter P 38 Wang CS 281 Wang KS 38 Wang L 332, 406 Wang Z 235 Wanner G 428 Ward TJ 357 Wasserman SA 479 Watanabe K 327 Watanabe KI 112, 171 Watanabe MM 80, 86, 115 Waterbury J 83 Waterston R 278 Watson AJ 169 Watson JD 283 Weber K 74 Weber NS 234 Webley DM 277 Wedekind JE 33 Wee JL 402 Wehrli E 282 Weigand F 363 Weigang F 235, 236 Weihe A 431 Weijman ACM 283 Weiner AJ 38 Weinreb SM 323 Weir A 239 Weisburg WG 229 Weising K 363, 381 Wellbrock U 76, 110, 170 Wells ID 328 Welsh J 363 Wendel JF 399, 401, 405, 429 Wendland J 239 Wenk-Seifert I 74, 227 Weresub LK 231 Werner O 405 Westerneng TJ 238 Wetherbee R 176 Wettern M 79 Whatley FR 86 Whatley JM 86

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Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

Wheals AE 322 Wheeler WC 406 Whelen S 233 White JJ 363 White TJ 229, 232, 239, 356, 357, 406 Whyte AC 332 Wickham GS 31 Wickliffe E 356 Wicklow DT 326, 332 Widmer C 278 Wiedenmann J 114 Wiemer EA 38 Wiewiorowska M 323 Wightman TF 112 Wilcox LW 78, 171 Wilde V 400, 428 Willard HF 31 Williams BAP 86 Williams DM 173, 175, 178 Williams JGK 363 Williams L 327 Williamson DH 78, 86 Willmot DB 383 Wilmotte A 283 Wilson R 401, 402, 453, 457 Wilson RJM 78, 86 Wilson TJ 34, 38 Wing RA 229, 278 Wingfield BD 238, 239, 240, 356, 362 Wingfield MJ 229, 231, 238-240, 282, 356, 362 Winka K 229, 240 Winter P 381 Wipf D 356 Witthuhn RC 240 Woese CR 38, 86, 239, 281, 360, 363 Wogan GN 323 Wojciechowski MF 400 Wolf G 332 Wolf M 401 Wolf PG 403 Wolf S 401 Wolfe GR 86 Wolfe K 232

Wolfson R 431 Wolk CP 80 Wolters J 115 Wong FTW 116 Wong JCW 115 Wong JTY 115, 116 Wong SM 332 Wood AM 116 Wood V 240, 283 Workneh F 381 Wöstemeyer J 239, 360 Wright SW 80, 114 Wu HN 38 Wu S 406 Wurtz J 111 Wurzner R 327 Wutz A 38 Wyss P 359 Xiao S 38 Xu HX 283 Xu S 33 Xu Z 401 Xue L 111 Xypolyta A 178 Yacentyuk SP 456 Yaguchi T 330, 332 Yahara T 404 Yamada Y 281, 283, 284, 330, 479 Yamamoto H 381, 383 Yamamoto K 325, 331 Yamamoto M 112 Yamamoto Y 329 Yamaoka S 406 Yamato KT 406 Yamazaki M 325 Yan X 401 Yang XB 381 Yarrow D 283, 284 Yaser M 358 Yashiro K 332 Yeo SH 332 Yergeau E 363

517

Author Index

Yeung PKK Wong 116 Yike I 239 Yin WS 323 Yli-Mattila T 361, 363 Yoder OC 357 Yoder WT 363 Yokota A 282 Yokoyama K 326, 332 Yokoyama T 326 Yolken RH 234 Yoon HS 74, 78, 79, 87, 111, 116 Yoshida K 85 Yoshimura A 228 Yoshizawa-Ebata J 112 Young KJ 38 Yu TS 332 Yuan YM 407 Yusupov MM 38 Yusupova GZ 38 Zabeau M 362 Zafari D 232 Zalar P 240 Zamore PD 37, 38 Zang Mu 277 Zare R 232 Zaslavkaia L 74

Zaug AJ 34 Zauner S 77, 80, 112 Zechman FW 178 Zelger T 240 Zettler ER 76 Zhang J 281 Zhang N 357 Zhang Y 38 Zhang Z 87, 116, 281 Zhao L 283 Zhao SW 356 Zhao Z 281 Zhao ZY 34 Zhou K 32 Zhuze AP 171 Ziegler K 235 Zieve GW 38 Ziman PA 115 Zimmer EA 178, 402 Zimmermann NFA 430 Zink DL 327 Zohri AN 324 Zoldos V 407 Zoller S 403 Zondervan RL 112 Zotti M 332 Zuck RK 240 Zuker M 429

PLANT GENOME: BIODIVERSITY AND EVOLUTION Volume 2, Part B Lower Groups

Detailed Contents of Volume 2, Part B

Preface to the Series v Preface to this Edition vii List of Contributors xi 1. From Ancient to Modern RNA World: A Metabolic Story 1 Introduction 2 Non-coding RNAs 4 The Small Nuclear RNAs (snRNAs) 4 The Small Nucleolar RNAs (snoRNAs) 4 The Transfer Messenger RNAs (tmRNAs) 5 The RNA Signal Recognition Particule (RNA-SRP) 6 Vault RNA 6 The Small Interfering RNAs (siRNA) 6 The Micro RNAs (miRNAs) 7 The Small Temporal RNAs (stRNA) 7 The X Inactive-Specific Transcript RNAs (Xist RNA) 8 Major Naturally Occurring Ribozymes 9 The Ribonuclease P (RNase P) 9 The Class I Introns 9 The Class II Introns 11 The Small Self-Cleaving RNAs Family 11 The Hairpin Ribozyme 14 Structure 14 Catalytic mechanism 16 Varkud Satellite Ribozyme 18 Structure 18

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Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

Catalytic Mechanism 21 The Hepatitis Delta Virus Ribozyme Structure 22 Catalytic mechanism 24 The Hammerhead Ribozyme 26 Structure 26 Catalytic mechanism 28 Conclusion 30 Acknowledgments 30 References 30

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2. Plastid Origin: A Driving Force for the Evolution of Algae Introduction 40 Assessing the Algal and Protists Evolution 44 Primary Endosymbiosis: the Birth of Plastids 48 Secondary endosymbiosis: spreading the plastids 57 The Green Secondary Plastids 59 The ‘Red’ Secondary Plastids 62 The ‘chromalveolate’ hypothesis 66 Plastids Replacements in Dinoflagellates 69 Conclusion 73 Acknowledgments 73 References 74

39

3. Evolution and Diversity of Dinoflagellates: Molecular Perspectives 117 Introduction 90 Early Evolution of Dinoflagellates 90 The Dinoflagellate “Mesokaryotic” Nature, Histone-like Proteins and Evolution of the Liquid Crystal Genome 91 Molecular Phylogeny of the Dinoflagellates 93 Overview 93 Description of the Major Orders of Dinoflagellates 96 Multi-species Complexes 99 Intra-specific Variation 101

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Plastid Genomes of Dinoflagellates Minicircles 107 Conclusion 109 Acknowledgments 109 References 110

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4. Evolution of the Diatoms 117 Introduction 118 Morphology of the Silica Frustule 119 Taxonomy Based on Characteristics of the Silica Frustule 123 Phylogenies Based on Characteristics of the Silica Frustule 133 A Phylogeny Inferred from Nuclear SSU rDNA Sequences 134 Basal centrics 138 The Radial Centrics 138 The Multipolar Centrics 139 The Araphid Pennates 144 The Raphid Pennates 148 Phylogenetic Signal in the Life Cycle and Auxospore Ontogeny 151 Gamete Formation 153 Auxospore Development 155 Phylogenetic Signal in Cytoplasmic Ultrastructural Features 157 Palaeontology and Phylogeny 161 New Directions in Diatom Phylogeny 165 Acknowledgements 169 References 169 5. Ascomycota: Introduction to Biodiversity, Evolutionary Genomics and Systematics Introduction 180 Fossil Record of the First Fungi 182 Fungal Biodiversity 182 Anamorph, Teleomorph, Holomorph 183 Species Concepts Used in Fungal Taxonomy 185 Systematics Based on Morphological Characters 187

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Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

Molecular Approaches to Fungal Phylogenetics 188 Comparative Genomics of Ascomycetous Fungi 192 Genome Analysis of Saccharomyces cerevisiae 192 Genome Analysis of Ashbya gossypii 193 Comparative Genomics among Hemiascomycetes 196 Organization of Introns of Hemiascomycetous Yeasts 197 Genome Analysis of Schizosaccharomyces pombe 199 Genome Analysis of Euascomycetes 200 Ascomycete-specific Genes 203 Phylogenetic Relationships among the Ascomycota and their Anamorphs 204 Hemiascomycetes 207 Archiascomycetes 209 Euascomycetes 211 Plectomycetidae 212 Pyrenomycetidae 215 Loculoascomytidae 221 Discomycetes 223 Acknowledgments 226 References 226 6. Yeast Biodiversity and Evolution Definition of Yeast 242 Morphological Diversity 243 Ultrastructure of the Cell Wall and Hyphal Septa 247 Sexual Cycle 249 Life Cycle of Ascomycetous Yeasts 249 Life Cycle of Basidiomycetous Yeasts 251 Cell Wall Composition 254 G+C Content 256 CoQ Type 257 Yeast Diversity and Evolution as revealed by RNA/DNA Sequencing 259 5S rRNA 259 5.8S rRNA 260

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18S rDNA 260 26S RDNA 262 Other House-keeping Genes 263 Sequencing of the whole Genome 263 Genome Sequencing and the Theory of whole Genome Duplication 265 Overview of Important Groups of Yeasts 266 Ascomycetous yeasts 266 Basidiomycetous yeasts 272 Conclusion 276 References 277 7. Evolutionary Relationships among Economically Important Species of Aspergillus Subgenera Aspergillus and Fumigati 285 Introduction 286 Aspergillus Species as Pathogens of Plants and Animals 286 Mycotoxins Produced by Aspergilli 287 Aspergillus Species in the Food and Pharmaceutical Industries 288 Taxonomic outline of the Aspergillus genus 289 Aspergillus subgenus Aspergillus 289 Aspergillus section Aspergillus 289 Aspergillus section Nigri 295 Aspergillus section Flavi 298 Aspergillus section Circumdati 306 Aspergillus section Terrei 307 Aspergillus sections Cervini, Candidi and Cremei 310 Aspergillus subgenus Fumigati 311 Aspergillus section Fumigati 311 Aspergillus section Clavati 319 Conclusion and Future Prospects 321 Acknowledgments 322 References 322

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Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

8. Polymerase Chain Reaction-based Methods in Fusarium Taxonomy 333 Introduction 334 Taxonomy 334 PCR-based Methods 336 1. Random Amplified Polymorphic DNA (RAPD-PCR) 337 2. Restriction Fragment Length Polymorphism (RFLP) 341 3. Ribosomal RNA (rRNA) and Ribosomal DNA (rDNA) Sequence Comparisons 343 ITS regions 347 4. Micro- and Minisatellites 350 5. DGGE 351 6. Amplified Fragment Length Polymorphism (AFLP) 352 Conclusion 354 References 354 9. Use of Molecular Markers to Study Host-Pathogen Co-evolution: A Case Study of the Fungal Pathogen Phialophora gregata 365 Introduction 366 The Brown Stem Rot Pathosystem 367 Molecular Markers for Different Taxonomical Levels 368 Molecular Markers for Separating the Pathogen from Other co-colonizing Fungi 368 Molecular Markers for Separating Formae Speciales of P. greagta 369 Molecular Markers for Separating Pathotypes of P. gregata f. sp. sojae 370 Nuclear rDNA marker 370 Microsatellite markers 373 Anonymous DNA markers 374 Correspondence of Molecular Markers with Pathotypes 375 Correlation of Molecular Markers with Host Cultivar Preference 376 Association of Independent Genetic Traits and Clonal Reproduction 378

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Acknowledgments References 380

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10. Utility of the Internal Transcribed Spacers of the 18S-5.8S-26S Nuclear Ribosomal DNA in Land Plant Systematics with Special Emphasis on Bryophytes 385 Introduction 386 Location and General Features of the ITS 388 ITS Amplification, Sequencing, and Analysis 392 Amplification and Sequencing 392 Sequence Alignment 393 Phylogenetic Analysis 394 Phylogenetic Utility and Limitations 396 Conclusion 398 References 399 11. Molecular Evolution of the atpB-rbcL Intergenic Spacer in Bryophytes Introduction 410 Material and Methods 411 Sampling of sequences 411 Alignment, Secondary Structures and Statistics 412 Phylogenetic analyses 413 Results and Discussion 414 Sequences 414 Length and GC-content 415 Alignments, repeats, and Sequence Variation 417 Promoter Elements 419 Hairpin Secondary Structures 421 Phylogenetic Inference 423 Intraspecific Sequence Variation 426 Conclusion 426 Acknowledgments 427 References 427

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Plant Genome: Biodiversity and Evolution—Vol. 2, Part B

12. Molecular Phylogeny and Biogeography of Plagiochila (Jungermanniidae: Plagiochilaceae) 433 Introduction 434 Species Ranges: Traditional concepts 434 Supraspecific Classification of Plagiochila Based on Morphology 435 Implications from Molecular Phylogenies 436 Molecular Markers 436 Current Hierarchical Subdivisions of Plagiochila are not Reflected in Molecular Topologies 437 Natural Species Groups Span Several Floristic Kingdoms 437 Molecular Topologies and Morphology: the Case Study Plagiochila sect. Superbae 443 Morphological Datasets of Plagiochila have a High Content of Homoplasy 447 Taxonomic Decisions Based on Molecular Topologies: Plagiochila detecta, Plagiochila maderensis, Plagiochila rutilans and Rhodoplagiochila 449 Phylogeography 450 Acknowledgments 452 References 452 13. Genome Studies of Cactus Yeast The Cactus-Yeast—Drosophila Community 459 The Host Cacti 459 Cactophilic Insects 460 The Cactophilic Yeast 462 Cactus-Yeast Interactions 463 Drosophila-Yeast Interactions 465 Yeast-Yeast Interactions 466 Genome Studies of Cactophilic Yeast 466 The Pichia cactophila Species Complex 467 The Pichia kluyveri complex 468 Candida Sonorensis 471 The Sporopachydermia cereana Species Complex

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Clavispora opuntiae 472 Other Cactophilic Yeast Species Conclusion 473 Acknowledgments 474 References 474

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