Advances in Parasitology Volume 56

Advances in PARASITOLOGY VOLUME 56 Editorial Board M. Coluzzi, Director, Istituto de Parassitologia, Universita` Deg...

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Advances in PARASITOLOGY

VOLUME 56

Editorial Board M. Coluzzi, Director, Istituto de Parassitologia, Universita` Degli Studi di Roma ‘La Sapienza’, P. le A. Moro 5, 00185 Roma, Italy C. Combes, Laboratoire de Biologie Animale, Universite´ de Perpignan, Centre de Biologie et d’Ecologie Tropicale et Me´diterraneénne, Avenue de Villeneuve, 66860 Perpignan Cedex, France D.D. Despommier, Division of Tropical Medicine and Environmental Sciences, Department of Microbiology, Columbia University, 630 West 168th Street, New York, NY 10032, USA J.J. Shaw, Instituto de Cieˆncias Biome´dicas, Universidade de Saõ Paulo, av. Prof. Lineu Prestes 1374, 05508-900, Cidade Universita´ria, Saõ Paulo, SP, Brazil K. Tanabe, Laboratory of Biology, Osaka Institute of Technology, 5-16-1 Ohmiya Asahi-Ku, Osaka, 535, Japan P. Wenk, Falkenweg 69, D-72076 Tu¨bingen, Germany

Advances in PARASITOLOGY Edited by

J.R. BAKER Royal Society of Tropical Medicine and Hygiene, London, England

R. MULLER London School of Hygiene and Tropical Medicine, London, England and

D. ROLLINSON The Natural History Museum, London, England

VOLUME 56

Amsterdam Boston Heidelberg London New York Oxford Paris San Diego San Francisco Singapore Sydney Tokyo

ELSEVIER Ltd The Boulevard, Langford Lane Kidlington, Oxford OX5 1GB UK ß 2004 Elsevier Ltd. All rights reserved. This work is protected under copyright by Elsevier Ltd, and the following terms and conditions apply to its use: Photocopying Single photocopies of single chapters may be made for personal use as allowed by national copyright laws. Permission of the Publisher and payment of a fee is required for all other photocopying, including multiple or systematic copying, copying for advertising or promotional purposes, resale, and all forms of document delivery. Special rates are available for educational institutions that wish to make photocopies for non-profit educational classroom use. Permissions may be sought directly from Elsevier’s Rights Department in Oxford, UK: phone (þ44) 1865 843830, fax (þ44) 1865 853333, e-mail: [email protected]. Requests may also be completed on-line via the Elsevier homepage (http://www.elsevier.com/locate/permissions). In the USA, users may clear permissions and make payments through the Copyright Clearance Center, Inc., 222 Rosewood Drive, Danvers, MA 01923, USA; phone: (þ1) (978) 7508400, fax: (þ1) (978) 7504744, and in the UK through the Copyright Licensing Agency Rapid Clearance Service (CLARCS), 90 Tottenham Court Road, London W1P 0LP, UK; phone: (þ44) 20 7631 5555; fax: (þ44) 20 7631 5500. Other countries may have a local reprographic rights agency for payments. Derivative Works Tables of contents may be reproduced for internal circulation, but permission of the Publisher is required for external resale or distribution of such material. Permission of the Publisher is required for all other derivative works, including compilations and translations. Electronic Storage or Usage Permission of the Publisher is required to store or use electronically any material contained in this work, including any chapter or part of a chapter. Except as outlined above, no part of this work may be reproduced, stored in a retrieval system or transmitted in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, without prior written permission of the Publisher. Address permissions requests to: Elsevier’s Rights Department, at the fax and e-mail addresses noted above. Notice No responsibility is assumed by the Publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. Because of rapid advances in the medical sciences, in particular, independent verification of diagnoses and drug dosages should be made. First edition 2004 British Library Cataloguing in Publication Data A catalogue record is available from the British Library. Academic Press An imprint of Elsevier 84 Theobald’s Road London WC1X 8RR, UK http://www.academicpress.com

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CONTRIBUTORS TO VOLUME 56 E. U. CANNING, Department of Biological Sciences, Imperial College London, Silwood Park Campus, Ascot, Berkshire, SL5 7PY, UK N. B. CHILTON, Department of Veterinary Science, The University of Melbourne, Werribee, Victoria 3030, Australia G. A. DOSREIS, Instituto de Biofı´sica Carlos Chagas Filho, Centro de Cieˆncias da Sau´de, Bloco G, Universidade Federal do Rio de Janeiro, 21 944 970–Ilha do Fundaõ, Cidade Universita´ria, Rio de Janeiro, RJ, Brazil R. E. FOWLER, Membrane Biology Group, Division of Biomedical Sciences, University of Edinburgh, George Square, Edinburgh EH8 9XD, UK R. B. GASSER, Department of Veterinary Science, The University of Melbourne, Werribee, Victoria 3030, Australia N. HEISE, Instituto de Biofı´sica Carlos Chagas Filho, Centro de Cieˆncias da Sau´de, Bloco G, Universidade Federal do Rio de Janeiro, 21 944 970–Ilha do Fundaõ, Cidade Universita´ria, Rio de Janeiro, RJ, Brazil M. HU, Department of Veterinary Science, The University of Melbourne, Werribee, Victoria 3030, Australia C. JONES, Laboratory for Molecular Structure, National Institute for Biological Standards and Control, Potters Bar, Hertfordshire, EN6 3QG, UK G. MARGOS, Malaria Laboratory, Peter Gorer Department of Immunobiology, Guy’s, King’s College and St Thomas’s Hospitals’ School of Medicine, New Guy’s House, Guy’s Hospital, London, SE1 9RT, UK L. MENDONÇA PREVIATO, Instituto de Biofı´sica Carlos Chagas Filho, Centro de Cieˆncias da Sau´de, Bloco G, Universidade Federal do Rio de Janeiro, 21 944 970–Ilha do Fundaõ, Cidade Universita´ria, Rio de Janeiro, RJ, Brazil ADVANCES IN PARASITOLOGY VOL 56 ISSN: 0065308X DOI: 10.1016/S0065-308X(03)56005-5

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CONTRIBUTORS TO VOLUME 56

G. H. M ITCHELL , Malaria Laboratory, Peter Gorer Department of Immunobiology, Guy’s, King’s College and St Thomas’s Hospitals’ School of Medicine, New Guy’s House, Guy’s Hospital, London, SE1 9RT, UK B. OKAMURA, School of Animal and Microbial Sciences, University of Reading, Whiteknights, Reading, Berkshire, RG6 6AJ, UK J. O. PREVIATO, Instituto de Biofı´sica Carlos Chagas Filho, Centro de Cieˆncias da Sau´de, Bloco G, Universidade Federal do Rio de Janeiro, 21 944 970– Ilha do Fundaõ, Cidade Universita´ria, Rio de Janeiro, RJ, Brazil A. R. TODESCHINI, Instituto de Biofı´sica Carlos Chagas Filho, Centro de Cieˆncias da Sau´de, Bloco G, Universidade Federal do Rio de Janeiro, 21 944 970–Ilha do Fundaõ, Cidade Universita´ria, Rio de Janeiro, RJ, Brazil R. WAIT, Kennedy Institute for Rheumatology Division, Faculty of Medicine, Imperial College London, 1 Aspenlea Road, London, W6 8LH, UK

PREFACE This volume opens with a piece of international collaboration, a review by Jose Previato and colleagues from the Federal and State Universities of Rio de Janeiro in Brazil and Imperial College London and the National Institute for Biological Standards and Control in the United Kingdom. This team of authors comprehensively reviewed the structure and biosynthesis of the unusual glycoinositolphospholipids (GIPLs) present on the surface of Trypanosoma cruzi and other trypanosomatids. These compounds occur in two different forms on different isolates of T. cruzi, depending on the nature of the substitution of one of the mannosyl residues. Although the precise function of these molecules is unknown, it is clear that they can modulate the immune response and hence may well be involved in the pathogenesis of T. cruzi infection. Purified GIPL from T. cruzi suppresses T cell activation in mice, particularly of Th1 cells. It also blocks the ability of dendritic cells and macrophages to interact fully with T cells. These immunosuppressive effects presumably aid the parasite’s evasion of the host’s immune response to infection. The authors conclude that a complete understanding of the GIPL biosynthetic pathway in T. cruzi could lead to the development of a new class of chemotherapeutic agents. The second paper in the volume reviews a group of parasitic organisms which were for many years regarded by most workers as protozoa, although this was already being questioned at the very end of the nineteenth century. The evolutionary development of the group in question, first accorded the status of phylum Myxozoa in 1970 and now generally recognized as being degenerate metazoans, is reviewed by Elizabeth Canning and Beth Okamura from Imperial College London and the University of Reading in the UK. Both these authors have played a significant role in the recent work elucidating ADVANCES IN PARASITOLOGY VOL 56 ISSN: 0065308X DOI: 10.1016/S0065-308X(03)65006-2

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PREFACE

the full life cycle of several species of Myxozoa and in determining their metazoan status and phylogenetic relationships. This work revealed, to the surprise of many parasitologists, a link between two hitherto ‘distinct’ groups of parasites, the Actinosporea infecting invertebrates and the Myxosporea of vertebrates, both forms now being known to be stages in a single dioecious life cycle. The evolutionary origin of the Myxozoa, and consequently their phylogenetic relationships, are still disputed subjects. The resemblance of myxozoan polar capsules to the nematocysts of Cnidaria has led many to believe that the two groups are related. However, studies using techniques such as electron microscopy and DNA analysis tend to the idea that the Myxozoa are related to the Bilateria, a group of ‘primitive’ metazoa including animals such as nematodes and flatworms. This topic is discussed at length by Canning and Okamura, who conclude that ‘the balance of evidence is now heavily weighted in favour of Myxozoa as a [‘degenerate’] phylum within the Bilateria.’ The vexed question of the homologies, if any, of the polar capsules remains unanswered; Canning and Okamura suggest three possibilities, the last of which they regard as unlikely: acquisition by endosymbiosis, convergent evolution, and gene transfer. Ruth Fowler, Gabrielle Margos and Graham Mitchell of the Department of Immunology at Guy’s Hospital, London, UK, then review the molecular mechanisms of locomotion associated with the invasive zoite stages of apicomplexans; most of the available information relates to the species of medical or veterinary importance. They all possess a characteristic complex of apical organelles (hence the phylum name) used to invade the host cell. It is believed that there is a cell-surface linear acto-myosin motor with actin filaments anchored to a microtubule or some other cytoskeletal structure responsible for three types of locomotion including invasion of host cells. Many different myosins are described, some found only in this group. Developments in DNA sequencing technology have facilitated the rapid generation of full mitochondrial DNA sequences for a variety of animals. Well over 200 complete mitochondrial (mt) genomes have now been determined and are available from the public databases.

PREFACE

ix

Mitochondria are indeed fascinating organelles, which are believed to originate from free-living eubacteria undergoing endosymbiosis, so it is particularly relevant that their role and evolution in parasitic groups should be considered. Mitochondria evolve independently from the nuclear genome and much can be learnt by detailed study of gene arrangements, coding and non-coding regions, replication and transcription and mutations. Significant progress has been made with unravelling the complexities of mt genomes in the Nematoda. In the final chapter of this volume, Min Hu, Neil Chilton and Robin Gasser from the University of Melbourne, Australia present a comprehensive review of this topic, paying particular attention to implications for population genetics and systematic studies. The authors emphasize how studies of mt genomes can contribute to understanding the ecology and epidemiology of parasitic nematodes. John Baker Ralph Muller David Rollinson

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CONTENTS CONTRIBUTORS TO VOLUME 56 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . PREFACE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

v vii

Glycoinositolphospholipid from Trypanosoma cruzi: Structure, Biosynthesis and Immunobiology Jose O. Previato, Robin Wait, Christopher Jones, George A. DosReis, Adriane R. Todeschini, Norton Heise and Lucia Mendonça Previato

1. 2. 3. 4. 5.

Abstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Structural Characterization of T. cruzi Gipl . . . . . . . . . . . . . . . . . . . . . GPI Biosynthesis in T. cruzi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . T. cruzi GIPL and Host Immune Responses . . . . . . . . . . . . . . . . . . . . . Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 2 7 24 28 31 32 32

Biodiversity and Evolution of the Myxozoa Elizabeth U. Canning and Beth Okamura

1. 2. 3. 4. 5. 6. 7.

Abstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 43 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 44 Classification of the Phylum Myxozoa . . . . . . . . . . . . . . . . . . . . . . . . . . 46 Diagnostic Characters of Myxozoa . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 47 Diagnostic Characters of the Class Myxosporea . . . . . . . . . . . . . . . . . 54 Diagnostic Characters of the Class Malacosporea . . . . . . . . . . . . . . . 62 Phylogenetic Relationships of Myxozoa . . . . . . . . . . . . . . . . . . . . . . . . . 94 Myxozoan Lineages and Evolution of Parasitism . . . . . . . . . . . . . . . . 112 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117

ADVANCES IN PARASITOLOGY VOL 56 ISSN: 0065308X DOI: 10.1016/S0065-308X(03)65007-4

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CONTENTS

The Mitochondrial Genomics of Parasitic Nematodes of Socio-Economic Importance: Recent Progress, and Implications for Population Genetics and Systematics Min Hu, Neil B. Chilton and Robin B. Gasser

1. 2. 3. 4. 5.

Abstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Background on the Mitochondrial Genomics of Animals . . . . . . . . The Mitochondrial Genomes of Parasitic Helminths . . . . . . . . . . . . . Mitochondrial Gene Markers for Studying the Molecular Systematics and Population Genetics of Parasitic Nematodes . . . . Conclusions, Perspectives and Prospects . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

134 134 136 150 163 179 187 188

The Cytoskeleton and Motility in Apicomplexan Invasion Ruth E. Fowler, Gabriele Margos and Graham H. Mitchell

1. 2. 3. 4. 5.

Abstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Basic Description of the Apicomplexan Zoite and Organisation of the Cytoskeleton . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . External Proteins Implicated in Motility . . . . . . . . . . . . . . . . . . . . . . . . . Cytoskeletal and Motor Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Models of Motility . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

214 214 215 219 221 242 249 249

INDEX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 265

CONTENTS


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Glycoinositolphospholipid from Trypanosoma cruzi: Structure, Biosynthesis and Immunobiology Jose O. Previato1, Robin Wait2, Christopher Jones3, George A. DosReis1, Adriane R. Todeschini1, Norton Heise1 and Lucia Mendonça Previato1,* 1

ˆ Instituto de Biofı´sica Carlos Chagas Filho, Centro de Ciencias da Sau´de, Bloco G, Universidade Federal do Rio de Janeiro, 21 944 970–Ilha do Fundaõ, Cidade Universita´ria, Rio de Janeiro, RJ, Brazil; 2 Kennedy Institute for Rheumatology Division, Faculty of Medicine, Imperial College London, 1 Aspenlea Road, London, W6 8LH, UK; 3 Laboratory for Molecular Structure, National Institute for Biological Standards and Control, Potters Bar, Hertfordshire, EN6 3QG, UK

Abstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Structural Characterization of T. cruzi GIPL . . . . . . . . . . . . 2.1. Methodology. . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Chemical Composition of T. cruzi GIPL . . . . . . . . . . . 2.3. Sample Preparation for Structural Studies of T. cruzi GIPL 2.4. Characterization of the PI-glycan Moiety of T. cruzi GIPL 2.5. Characterization of the Lipid Moiety of T. cruzi GIPL . . . 3. GPI Biosynthesis in T. cruzi . . . . . . . . . . . . . . . . . . . . . . 4. T. cruzi GIPL and Host Immune Responses . . . . . . . . . . . . 5. Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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

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

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

2 2 7 7 9 11 12 20 24 28 31 32 32

*Author for correspondence.


Copyright ß 2004 Elsevier Ltd. All rights of reproduction in any form reserved

2

JOSE O. PREVIATO ET AL.

ABSTRACT The pathogenic protozoan parasite Trypanosoma cruzi expresses on its surface an unusual family of glycoinositolphospholipids (GIPLs) closely related to glycosylphosphatidylinositol (GPI) anchors. Different parasite isolates express distinct GIPLs which fall into two series, depending on the substitution of the third mannosyl residue in the conserved glycan sequence Man4-(AEP)GlcN-InsPO4 by ethanolamine phosphate or -galactofuranose. Although the exact role of these molecules in the cell biology and pathogenicity of T. cruzi remains unknown, the lipid and glycan moieties impart distinct responses to host T and B lymphocytes and phagocytes, overall favouring an immune response permissive to the parasite. The biosynsthesis of GIPLs follows a pathway similar to that observed for GPI anchors. However, a more detailed understanding might enable the development of specific inhibitors of parasite-specific enzymes and lead to novel drugs to ameliorate Chagas disease.

1. INTRODUCTION The flagellate protozoan Trypanosoma cruzi, the aetiological agent of Chagas disease, belongs to the family Trypanosomatidae and is transmitted to humans by blood-sucking triatomine insects in endemic areas of Latin America. The acute phase of Chagas disease often includes parasitaemia, prior to onset of a chronic phase that may have varying clinical features. Of the 12 million people infected (Moncayo, 1999), a proportion develop myocarditis or pathological abnormalities of the digestive and peripheral nervous systems, whereas others remain asymptomatic (WHO, 2000). The basis for this variable pathology remains unknown, but must result from the interaction of host- and parasite-specific factors. This is consistent with well-established strain-to-strain variations in morphology, virulence, tissue tropism, and pathology (Melo and

GLYCOINOSITOLPHOSPHOLIPID FROM TRYPANOSOMA CRUZI

3

Brener, 1978; Murta et al., 1998), as well as in several genetic markers1. The complex interactions between parasite and host cells are likely to be mediated by the molecular architecture of the cell surface, which is rich in glycoconjugates. Since a range of hostile environments confronts T. cruzi during its life cycle (Brener, 1973), it is likely that its unusual surface glycomolecules play a crucial role in its survival strategies. In protozoa, cell surface proteins are frequently attached to the membrane via glycosylphosphatidylinositol (GPI) anchors (McConville and Ferguson, 1993). In parasites of the family Trypanosomatidae GPI anchors are also used to attach cell surface polysaccharides, for example the lipophosphoglycan (LPG) of Leishmania spp. (Turco and Descoteaux, 1992); or mono- or oligosaccharides, such as the glycoinositolphospholipids (GIPLs) of T. cruzi (see Previato et al., 1990; Lederkremer et al., 1991) and Leishmania spp. (McConville and Ferguson, 1993). All GPI-anchored molecules contain the conserved core motif Man(1 ! 4)GlcN (1 ! 6)Ins1-PO3 , linked to either glycerolipid or to sphingolipid. McConville and Ferguson (1993) showed that, in Leishmania species, the conserved core motif is further modified by the addition of mannose (Man), or Man and galactose (Gal) residues, to form three distinct lineages of GIPL, designated type 1, type 2 and hybrid. This classification is not restricted to Leishmania, since glycoconjugates from other parasites appear to conform to the same architecture. In type 1 the Man of the conserved core is substituted with a (1 ! 6)-linked Man, whereas in type 2 a Man is (1 ! 3)-linked to this residue. The hybrid type is a branched structure in which two Man substituents are present, linked (1 ! 3) and (1 ! 6) (Figure 1). A noteworthy feature is that, of all parasite GPI anchored molecules characterized, only the T. cruzi GIPL contains a type 1 conserved glycan core, similar to that of the host GPI-linked proteins (Figure 1).

1

Macedo, A.M., Oliveira, R.P. and Pena, S.D.J. (2002). Chagas’ disease: role of parasite genetic variation in pathogenesis. Expert Reviews in Molecular Medicine. 5 March, http://www-ermm. cbcv.cam.ac.uk/02004118.htm.


4 Conserved core motif of GPI-family molecules

Type

Model

1

GPI- protein anchors (Ferguson, 1999)

1

T. cruzi GIPL (Previato et al., 1990, Lederkremer et al., 1991)

HO O

HO

OH HO O O

HO

OH

O

HO

HO

HO O

O HO

P

O

LIPID

RO O NH2

O

OH

HO HO HO O

HO

OH HO

O HO

O O

HO

OH

NH2

P

O

O

HO

HO O HO

O

P

O

LIPID

OH O NH2

O

OH

HO HO OH O

HO

OH

O

O

O HO

HO OH

HO

HO

HO

O

O

O

LIPID

2

RO O NH2

HO

P

O

OH

Leishmania spp. GIPL and LPG (McConville and Ferguson, 1993); Leptomonas samueli GIPL; (Previato et al., 1992, 1994); Endotrypanum spp. GIPL (Xavier da Silveira et al., 1998)

HO HO O

HO

OH HO O O

HO

OH O

OH O HO

HO

HO O HO

HO HO

O O

P

O RO

O NH2

O

LIPID

Hybrid

Leishmania spp. GIPL (McConville and Ferguson, 1993; Previato et al., 1997) and Herpetomonas samuelpessoai, (Routier et al., 1995)

OH

HO HO

Figure 1 Structures of the conserved core motif of glycosylphosphatidylinositol (GPI)-family molecules; GIPL ¼ glycoinositolphospholipids, LPG ¼ lipophosphoglycan.

In T. cruzi, these unusual phosphoinositol-containing glycolipids, originally called lipopeptidophosphoglycan (LPPG) (Lederkremer et al., 1976), are major surface components of the experimentally accessible epimastigote forms, and thus have been the subject of many investigations (Lederkremer et al., 1977, 1978, 1980, 1985, 1991;


5

Ferguson et al., 1982; Mendonça-Previato et al., 1983; Mendonça Previato and Previato, 1985; Previato et al., 1990; Carreira et al., 1996). The primary structure of the GIPL from T. cruzi Y strain was the first complete structure reported for a member of the GIPL family (Table 1) (Previato et al., 1990; Lederkremer et al., 1991). The glycan is composed of a tetramannose sequence additionally substituted by two terminal galactofuranosyl units, with a 2-aminoethylphosphonate (AEP) substituent on O-6 of the glucosamine (GlcN) in the conserved core motif. The lipid moiety is a lignoceroylsphinganine. Subsequent study of a wider range of strains revealed additional structural diversity. Although all T. cruzi derived-GIPLs share the same conserved core, they can be classified into two series, which differ in the substitution of the third Man of the tetramannosyl sequence. In series 1, this substituent is ethanolaminephosphate (EtNP) linked to O-6, whereas in series 2 a galactofuranosyl (Galf ) unit is present on O-3 (Table 1). The biosynthesis of protein-linked GPI anchors and GIPLs involves glycosylation reactions that have been characterized in several models (Ferguson, 1999), and were recently validated as targets for drugs against human African trypanosomiasis (Ferguson, 2000). In T. cruzi, the difference in lipid composition of proteinlinked GPI anchors (Gu¨ther et al., 1992; Heise et al., 1995; Previato et al., 1995; Almeida et al., 2000) and GIPLs (Previato et al., 1990; Lederkremer et al., 1991), together with the presence of unique structural features, may represent additional targets for new chemotherapeutic agents against Chagas disease. The function of GIPLs in the biology of T. cruzi, or their role as a virulence factor, have not been unambiguously defined, although the molecules are antigenic. The non-reducing terminal Galf residues are crucial, since oxidation and reduction, which convert them to arabinofuranose, destroy antigenicity. Furthermore, soluble -D-Galf(1 ! 3)--D-Manp-OMe inhibited precipitation by 75%. This was the first demonstration of the antigenic activity of specific sugar units in T. cruzi (see Mendonça-Previato et al., 1983). More recently, studies on cells of the host immune system have shown that T. cruzi GIPLs are bifunctional molecules, the lipid and glycan elements of which elicit different biological responses (DosReis et al., 2002). Whilst the

6

Table 1 Structure of the major phosphatidylinositol glycan (PI-glycan) species present in T. cruzi glycoinositolphospholipids (GIPLs)a GIPL

PI-glycan EtNP AEP # # 6 6 -Manp-(1 ! 2)--Manp-(1 ! 2)--Manp-(1 ! 6)--Manp-(1 ! 4)--GlcN-(1 ! 6)-Ins-P

T. cruzi strain

Tulahuen

Series 1 EtNP AEP # # 6 6 -Galf-(1 ! 3)--Manp-(1 ! 2)--Manp-(1 ! 2)--Manp-(1 ! 6)--Manp-(1 ! 4)--GlcN-(1 ! 6)-Ins-P

G

Galf AEP # # 6 3 -Galf-(1 ! 3)--Manp-(1 ! 2)--Manp-(1 ! 2)--Manp-(1 ! 6)--Manp-(1 ! 4)--GlcN-(1 ! 6)-Ins-P Y and CL

a

Abbreviations are explained in the text.


Series 2


7

ceramide-containing moiety modulates T lymphocytes and phagocytes, the glycan chain stimulates NK cell activity and antibody production. In this review we describe advances in the study of T. cruzi GIPLs, with an emphasis on their molecular structure, since understanding of the structural features of GIPLs is important for explaining both their biological function and their role in virulence.

2. STRUCTURAL CHARACTERIZATION OF T. CRUZI GIPL 2.1. Methodology The characterization of glycoconjugates such as parasite GIPLs is one of the most challenging problems in structural biochemistry. Whereas protein primary structure may be specified by determination of the identities and sequence of the constituent amino acids, for carbohydrates, in addition to establishing the monosaccharide sequence, it is also necessary to determine ring geometry, absolute stereochemistry, the positions and anomeric configurations of the glycosidic bonds, and the branching pattern. In the case of glycolipids the lipid moiety must also be identified. GIPL molecules from T. cruzi present further difficulties because they frequently contain unusual constituents such as Galf (Lederkremer and Colli, 1995), noncarbohydrate substituents, e.g. AEP (Ferguson et al., 1982; Lederkremer et al., 1985; Mendonça-Previato and Previato, 1985) and EtNP (Carreira et al., 1996). Moreover, because oligosaccharide biosynthesis is not template-directed, GIPLs are intrinsically heterogeneous. Thus, the structural elucidation of the GIPLs from T. cruzi has been heavily dependent on the development of sophisticated phy sicochemical methods and the use of controlled degradation (Figure 2). Mass spectrometry (MS) and nuclear magnetic resonance (NMR) spectroscopy are sensitive and powerful techniques for biopolymer characterization, which, together with chemical derivatization, have played a key role in the structure determination of the GIPLs from T. cruzi. Mass spectrometric analysis of a GIPL may provide either a molecular mass only (from which it is possible to deduce its


8

Glycan moiety

Methanolysis

Methyl glycosides, FAMEs, long chain base, alkyl glycerol, N-acetylation, trimethylsilylation

GC/MS

Sugar and lipid compositions

Conserved core motif

Methylation analysis

Partially Omethylated methyl glycosides

Lipid moiety

Alkaline degradation

Nitrous acid deamination

Isolation of phosphoinositol glycans

Inositolphospholipids

FAB -MS spectrometry

1D/2D NMR spectroscopy

FAB/MS spectrometry

GC/MS

Linkage between sugars

Molecular mass, sequence of monosaccharides, position and configuration of glycosidic bonds, location of substituents

Phospholipid components

Figure 2 Approaches for structural characterization of T. cruzi glycoinositolphospholipids; FAB ¼ fast atom bombardment, FAMEs ¼ fatty acid methyl esters, GC ¼ gas chromatography, MS ¼ mass spectrometry, NMR ¼ nuclear magnetic resonance.

composition in terms of monosaccharides and substituents), or a fragmentation pattern, which can define residue sequence, branching pattern and the location of substituents. NMR spectroscopy potentially enables complete structure determination, including identification of sugars and their ring forms, anomeric configuration and the sequence of, and linkages between, residues. It also reveals the absolute configurations of the sugar residues and the positions of substituents. Because NMR data are quantitative, the relative amounts of different glycan species in mixtures can be estimated by integration of appropriate resonance signals. Gas chromatography/mass spectrometry (GC/MS) finds extensive use in GIPL characterization for the identification of monosaccharides,


9

fatty acids and long chain bases. GC/MS is also a key element of the methylation analysis strategy for the determination of linkage positions of the sugar residues. Specific fragmentations in the mass spectra of O-methyl monosaccharide derivatives enable the position of linkages in the intact oligosaccharide to be deduced (Bjorndal et al., 1967). The requirement for thermal vaporization generally precludes analysis of intact glycoconjugates or oligosaccharides by chemical ionization (CI), electron ionization (EI) or GC/MS. Such compounds became routinely amenable to direct MS characterization with the development of ‘soft ionization’ methods, in which sample molecules are ionized directly from the condensed phase. The first high-masscapable soft ionization technology to find widespread application in carbohydrate analysis was fast atom bombardment (FAB) (Barber et al., 1981). While much gentler than EI, FAB ionization usually imparts sufficient excess energy to induce structurally informative fragmentation, though molecular ions tend to predominate. Glycosidic bond cleavages are usually the dominant mode of spontaneous fragmentation in the spectra of native and derivatized protonated oligosaccharides, and enable assignment of the sequence of residues and identification of the nature and location of branches and substituents. The various possible fragmentation processes are unambiguously described by a system of nomenclature proposed by Domon and Costello (1988) (Figure 3).

2.2. Chemical Composition of T. cruzi GIPL The prototypical T. cruzi GIPL could be considered to be the major component derived from Y strain (Camargo, 1964). Early structural studies of this glycolipid established the presence of Gal, Man, glucose (Glc), and traces of GlcN (Lederkremer et al., 1976). Mild periodate oxidation and reduction yielded L-arabinofuranosyl residue, indicating that the Gal unit was in the furanosyl configuration (Lederkremer et al., 1980). The presence of an unusual phosphonate substituent, AEP, was first demonstrated by paper chromatography after strong acid hydrolysis (Ferguson et al., 1982),


10

Y 2

1,5X 1

Z2

HO CH 2

Y1 Z1

HO CH2 O

Y0 Z0 HO CH2

O

O O

OH

OH

O

R

OH

O

HO OH

OH

B1

0,2A 1

C1 2,4A 2

OH

B2 C2

2,5 A 3

B 3

C 3

HO CH 2 4 3

5

OH

O

o 1

HO

4

3 OH

OR HO

2

OH

O

o OR 1

2

OH

Figure 3 Nomenclature of oligosaccharide fragment ions, and the numbering system used for bonds in pyranose and furanose rings in the Domon and Costello (1988) system; note that the numbers refer to bonds, not atoms.

and was confirmed by 31P NMR (Lederkremer et al., 1985) and 13C NMR (Mendonça-Previato and Previato, 1985). The lipid moiety was shown to be a phosphoinositolceramide apparently containing hexadecanoic and tetracosanoic acids, and two long chain bases, sphinganine and 17-methylsphinganine, the latter being the major component (Lederkremer et al., 1978), though the presence of methylsphinganine was not confirmed in subsequent studies. It is now known that the long chain bases of ceramide-linked GIPL from the Y strain are sphinganine and sphingosine in a molar ratio of 3:1 (Lederkremer et al., 1990; Previato et al., 1990) (Table 2).


Table 2

11

Lipid composition of T. cruzi glycoinositolphospholipidsa T. cruzi strain Tulahuen

G

Y

CL

100 –

80 20

85 15

60 tr

Alkyl of alkyl glycerol 16:00

tr

tr

–

40

Fatty acid 16:00 18:00 24:00

17 13 70

30 5 65

16 4 80

70 tr 30

Long chain base Sphinganine Sphingosine

a

Determined by GC/MS (values are expressed in mol%); tr ¼ trace.

2.3. Sample Preparation for Structural Studies of T. cruzi GIPL Mass spectra obtained from intact GIPL are complex, because alkyl and acyl group heterogeneity in the lipid anchor degrades signalto-noise ratio by distributing the ion current over several molecular species (McConville et al., 1990). It is therefore desirable to remove the lipid moiety before MS analysis. Although NMR spectra can be obtained from intact GIPL in aqueous dimethyl sulphoxide solution, the water-soluble phosphatidylinositol glycan (PI-glycan) obtained after delipidation is more amenable to further purification. Removal of the lipid moiety may be achieved enzymatically with phosphatidylinositol-specific phospholipase C (PIPLC), which cleaves the lipid/ phosphate bond of most GPIs, including diacylglycerol-, plasmanyl-, lyso-plasmanyl- and ceramide-containing species, except when the inositol (Ins) ring is acylated (Ferguson, 1992; Menon, 1994). Other methods of delipidation include treatment with aqueous hydrofluoric acid, which dephosphorylates the sample, and deamination with nitrous acid. Such drastic chemical degradation procedures are not the ideal method of sample preparation for GIPLs containing labile substituents. Hydrogen fluoride treatment results in the loss of all

12


phosphate-linked groups and (possibly) cleavage of susceptible glycosidic linkages such as those to Galf. Deamination of GIPLs with nitrous acid results in specific cleavage between GlcN and Ins, producing a water-soluble glycan terminating in 2,5-anhydromannose and an inositolphospholipid (Gu¨ther and Ferguson, 1993; Menon, 1994; Schneider and Ferguson, 1995). Deamination is a useful method if the primary objective is recovery of the lipid moiety, but is less suitable if the aim is subsequent characterization of the liberated glycan. Substituents containing primary amino groups are also deaminated under the usual reaction conditions; ethanolamine, for example, is converted to ethene-ol or to ethylene glycol (Mayor et al., 1990), increasing sample heterogeneity. Subsequent glycan analysis by MS may be compromised because sensitivity is reduced by loss of basic amino groups, and several informative fragmentations are less easy in their absence. Base treatment (Figure 2) is a better method to liberate glycan chains from GIPLs before MS and NMR analyses. Most substituents and glycosidic linkages are stable to this procedure, the main artefact being limited phosphate migration from O-1 to O-2 of inositol (Previato et al., 1992).

2.4. Characterization of the PI-glycan Moiety of T. cruzi GIPL Previato et al. (1990) first published a negative ion FAB spectrum of the liberated underivatized PI-glycan from GIPL. Positive and negative ion FAB spectra of T. cruzi Y strain-derived GIPL are shown in Figure 4. Abundant protonated (M þ H) and deprotonated (M H) molecules are present at m/z (mass/charge) 1501 in the positive and at m/z 1499 in the negative ion spectra, respectively, which were consistent with Hex6-(AEP)-HexN-InsPO4 structure. The signals at m/z 1339 and 1177 (in the positive ion spectrum) are consistent with Y6 and Y5 fragments and are accompanied by the expected 1,5Xn signals at m/z 1367 and 1205 respectively. In the negative ion spectrum, the corresponding deprotonated fragments were observed. A prominent signal at m/z 529 (527 in the negative


13

Figure 4 Positive (A) and negative (B) fast atom bombardment/ mass spectrometry spectra of the major phosphatidylinositol glycan from T. cruzi (Y strain) glycoinositolphospholipid; abbreviations are explained in the text.

14


mode), assigned as Y2, provides evidence of an AEP substituent on GlcN; there is no indication of additional non-carbohydrate substitutions elsewhere in the molecule. Ions at m/z 853 and 691, respectively consistent with Y3 and Y4 fragments, are accompanied by Zn and Xn ions at m/z 835, 673, 881 and 719. In the positive ion spectrum no glycosidic cleavage ions are observed between m/z 853 and 1177, indicating the presence of a hexose (Hex) substituent on the third Hex; a linear phosphoinositol-heptasaccharide, by contrast, would have a Y5 fragment at m/z 1015. In the negative ion spectrum a weak ion at m/z 1013 is observed, but at substantially lower abundance than m/z 851 and 1175, reflecting the fact that its formation requires the cleavage of two glycosidic bonds. This structure was partially confirmed by Lederkremer et al. (1991) using FAB-MS of the high performance liquid chromatography (HPLC)purified, N-acetylated (NAc) permethylated glycan liberated by PIPLC. The spectrum of the major HPLC fraction had a protonated molecule at m/z 1828, which was consistent with permethylated Hex6HexNAc-InsPO4, indicating that the N-acetylated AEP had been eliminated under the methylation conditions. An easy fragmentation between N-acetylglucosamine (GlcNAc) and Ins resulted in an ion at m/z 1484, which underwent additional Y type cleavages giving double cleavage fragments at m/z 1266, 1062, 654 and 450. The absence of a fragment at m/z 696 indicated that the Hex substituent was located on the third Hex distal to GlcN. Methylation analysis of the intact and partially acid hydrolysed GIPL enabled characterization of linkage positions between sugar residues and provided valuable confirmation of the structure (Previato et al., 1990). Permethylation of the intact GIPL resulted in 2 moles of terminal Galf and 1 mole of each of 2-O, 3-O, 6-O and 2,3-di-O-mannopyranosyl (Manp) substituted residues, consistent with a structure containing six Hex units. The location of -D-Galf linked 1 ! 3 to 1 ! 2 substituted Manp was inferred by methylation analysis of mild acid hydrolysed GIPL, which showed that 2,3substituted Manp was replaced by 2-substituted Manp, and that the 3-substituted Manp was converted to a non-reducing end-unit (Previato et al., 1990).


15

Examination of GIPLs from a wider range of T. cruzi strains (Tulahuen, G and CL) revealed additional structural diversity, specifically the existence of a second family of GIPLs in the Tulahuen and G strains, lacking the Galf branch on the third Man (Table 1). Some of the mass spectrometric properties of these molecules, and their distribution between strains, are summarized in Table 3. Figure 5 shows the positive ion FAB spectrum of a PI-glycan liberated by alkaline hydrolysis of the major GIPL from the Tulahuen strain. The signal at m/z 1300.5 represents the protonated molecule. In contrast to the Y strain GIPL (Figure 4) described above (and also to the CL strain GIPL), these are observed at even mass number, implying the presence of an odd number of nitrogen atoms (McLafferty and Turecek, 1993). However, the ions at m/z 529 (Y2), 557 (1,5X2), 691 (Y3), 719 (1,5X3) and 853 (Y4) are also observed in Y and CL GIPLs and support the common presence of the structure (Hex)2-(AEP)-GlcN-InsPO4. Thus the remaining portion of the molecule must contain an odd number of nitrogen atoms, which requires the presence of either a second nitrogen-containing substituent or an amino sugar. The ion at m/z 1138 is consistent with the loss of an unsubstituted Hex from the non-reducing terminal by a Y-type fragmentation. The increment of m/z 285 between m/z 853 and m/z 1138 does not fit any known monosaccharide, but is consistent with an EtNP-substituted Hex. Although the GIPLs of several trypanosomatid species contain either AEP (Previato et al., 1992, 1994; Routier et al., 1995) or EtNP (Winter et al., 1994;

Table 3 Summary of the fast atom bombardment/mass spectrometry data on the major phosphatidylinositol glycan of T. cruzi glycoinositolphospholipidsa Strain Tulahuen G Y and CL a

[M þ H] þ 1300.5 1462.4 1501.8

[M

H]

1298.3 1460.3 1499.7

Composition

Mr

(Hex)4-HexN-InsP-EtNP-AEP (Hex)5-HexN-InsP-EtNP-AEP (Hex)6-HexN-InsP-AEP

1299.332 1461.38 1500.429

[M þ H] þ and [M H] indicate protonated and deprotonated molecules, respectively; other abbreviations are explained in the text.

16


Figure 5 Positive ion fast atom bombardment/mass spectrometry spectra of the major phosphatidylinositol glycan from T. cruzi (Tulahuen strain) glycoinositolphospholipid; abbreviations are explained in the text.

Previato et al., 1997; Xavier da Silveira et al., 1998), this is the first instance of both substituents occurring in the same molecule. The GlcN appears to be exclusively substituted by AEP, because Y2 glycosidic cleavage fragments were observed only at m/z 529, and not at m/z 545 (Figures 4 and 5). The presence of an EtNP-substituted Hex in the subterminal position is supported by the prominent B2 fragment at m/z 448, also observed in the spectrum of a GIPL from Leishmania adleri (see Previato et al., 1997), which likewise has an EtNP substituent in this position. Other non-reducing terminal fragments present at m/z 610 (B3), 772 (B4) and 1040 (B5) provide further confirmation of the sequence (Figure 5). Other further GIPL related to this species (Table 1) has been characterized by MS (Table 3). The FAB spectrum of the major GIPL from the G strain contained


17

a protonated molecule at m/z 1462, corresponding to an EtNPsubstituted homologue of the Tulahuen GIPL containing an additional Hex residue. Fragments at m/z 529 (Y2), 691 (Y3), 853 (Y4) and m/z 1300 (Y6) and their associated X and Z ions indicate an AEP substituent located on GlcN.

2.4.1. Characterization of PI-glycan from T. cruzi GIPL by NMR Spectroscopy Detailed structural analysis of PI-glycans from T. cruzi Tulahuen, G, Y and CL strains has been performed by one- and two-dimensional homo- and heteronuclear NMR spectroscopy (Previato et al., 1990; Carreira et al., 1996). The different peaks in the NMR spectra were assigned with respect to the atom giving rise to them in the particular glycan structure (Figure 6) by the standard range of correlation experiments (see Dabrowski, 1994 for a review). Essentially, assignments within a sugar residue were made using correlations mediated through chemical bonds (coupling constants) and, because the magnitude of coupling constant depends on stereochemical factors, these also allowed the configuration of the sugar to be determined. The linkages between the sugars and their sequence were determined from through-space correlations (nuclear overhauser enhancements or NOEs) or long-range 1H–13C correlations across the glycosidic linkages. Additional information on the linkage positions is available from chemical shifts (substitution of an oxygen almost invariably causes a downfield shift in the chemical shift of the carbon to which it is attached and the hydrogen(s) attached to that carbon; see Bock et al., 1984). The anomeric configurations of the sugar residues were determined from the magnitude of the coupling constant between H-1 and H-2, the coupling constant between H-1 and C-1, the chemical shifts of H-1 and of C-1, and the patterns of NOEs from the H-1 to other hydrogens in the ring. The 13C resonances from sugars in the furanose ring form present in the GIPL structures of G, Y, and CL strains (Table 1) are

18


Figure 6 Partial 500 MHz 1H nuclear magnetic resonance spectra of the major phosphatidylinositol glycans of glycoinositolphospholipids from T. cruzi strains Tulahuen (A), G (B), and Y and CL (C). The Figure shows the spectral region containing resonances from anomeric protons. Peak labelling is as follows (see Table 1 for structures; abbreviations are explained in the text): 1, ! 4)-GlcN-(1 ! 6)-Ins-1-PO4; 2, ! 4)-GlcN-(1 ! 6)Ins-2-PO4; 3, ! 6)-Man-(1 ! 4); 4, ! 2)-Man-(1 ! 6); 5, ! 2,6)-Man(1 ! 2); 6, terminal -Man-(1 ! 2); 7, ! 3)-Man-(1 ! 2); 8, terminal -Galf-(1 ! 3); 9, ! 2,3)-Man-(1 ! 2) and 10, side chain -Galf-(1 ! 3).


19

Figure 7 Partial 500 MHz 1H nuclear magnetic resonance spectra of the major phosphoinositol glycans of glycoinositolphospholipids from T. cruzi strains Tulahuen (A), G (B), and Y and CL (C), showing the resonances from 2-aminoethyl phosphonate (AEP) and ethanolamine phosphate (EtNP). The integration of these resonances is a measure of the relative proportions of these two substitutents.

downfield from those in the pyranose ring form (with anomeric resonances below 105 ppm, for example, and C-4 below 84 ppm). The spectra of mixtures are invariably more complex than those of pure compounds but, by using data from the different PI-glycan samples, the intensity of the resonances can be used to assign specific structural features to particular PI-glycans. The assignments for the anomeric protons are shown in Figure 6, and Figure 7 shows the resolved resonances from the –CH2–NH2 resonances of the AEP and EtNP residues. Integration of these two resonances allows quantification of the relative amounts of these two substituents.

20


Combining data from NMR spectroscopy, MS and methylation analyses established the major glycan chain structures summarized in Table 1.

2.5. Characterization of the Lipid Moiety of T. cruzi GIPL 2.5.1.

GC/MS Characterization of the Lipid Components

GC/MS remains the method of choice for identification and quantification of the fatty acid and long chain base components of T. cruzi GIPLs, and early investigations established the presence of sphinganine, sphingosine, tetracosanoic, octadecanoic and hexadecanoic acids in GIPL hydrolysates (Lederkremer et al., 1990; Previato et al., 1990). Subsequently hexadecanoylsphinganine and tetracosanoylsphinganine were detected by GC/MS after PIPLC cleavage of LPPG from Y strain (Lederkremer et al., 1993). After treatment of GIPLs with aqueous methanolic HCl, fatty acid methyl esters and long chain bases can be conveniently separated by extraction with hexane (at low pH) and diethyl ether (at basic pH), respectively, as described by Gaver and Sweeley (1966). The resulting long chain bases are amenable to GC/MS analysis after N-acetylation and trimethylsilylation. Two peaks, in an approximate 3:1 ratio, are observed in methanolysates of GIPLs from the Y strain. Their mass spectra (Figure 8) are consistent with sphinganine ((2S,3R)-2-amino1,3-octadecanediol) and sphingosine ((2S,3R,4E)-2-amino-4-octadecene-1,3-diol), respectively. These are also present in methanolysates of GIPLs from strains G and CL, but the proportion of sphingosine is lower, being about 10% compared to sphinganine in G strain, and only a trace constituent in the CL strain (Table 2). Molecular ions are not observed in the EI spectra of either compound, but the molecular mass is easily deducible from M-15 fragments at m/z 472 (for sphinganine) or 470 (for sphingosine) and from the M-103 peaks (loss of CH2OSi(CH3)3), observed at m/z 384 (in sphinganine) and 382 (in sphingosine). A useful cleavage between C-3 and C-2 provides


21

Figure 8 Electron ionization/mass spectrometry spectra of the N-acetylO-trimethylsilyl derivatives of sphinganine (A) and sphingosine (B) from the major glycoinositolphospholipid of T. cruzi strain Y.

evidence of both the chain length and the degree of unsaturation. In sphinganine this process results in the fragment at m/z 313, whereas in sphingosine, which has a double bond in position four, the analogous fragment is observed at m/z 311 (Figure 8).

22


Fatty acid methyl esters (FAMEs) are also highly amenable to analysis by EI/MS and display the low mass CnH2n 1O2 ion series (m/z 73, 87, 101 etc.) characteristic of saturated FAMEs, together with the well-known McLafferty rearrangement product at m/z 74 as base peak. In all strains except CL, the most abundant component has a molecular ion m/z 382, and fragment ions at M-29, M-31 and M-43, consistent with the methyl ester of tetracosanoic acid. Methyl hexadecanoate is abundant in all cases, and is the major FAME of the CL strain (Table 2). Methanolysates of GIPLs from strain CL contain an additional component, the spectrum of which does not resemble that of a fatty acid methyl ester. The EI and CI spectra of this component (Figure 9) are consistent with a 1-O-alkyl-2,3-di-Otrimethylsilyl glycerol (Myher et al., 1974). Molecular ions are absent from the EI spectrum, but the M-15 peak at m/z 445 implies a relative molecular mass of 460. This value was confirmed by a CI experiment which revealed abundant protonated molecules at m/z 461 (Figure 9), consistent with 1-O-hexadecyl-2,3-di-O-trimethylsilyl glycerol. The base peak at m/z 205 in the EI spectrum originates by cleavage between carbons 1 and 2 of glycerol (with charge retention on the di-trimethylsilylated fragment), and provides evidence that the two trimethylsilyl groups are contiguous (Figure 9).

2.5.2.

MS Characterization of the Intact Lipid Components of T. cruzi GIPL

Hydrolysis of GIPLs to determine their fatty acid and long chain base composition precludes identification of the individual ceramide molecular species. This may be achieved by MS analysis of the phosphoinositolceramides liberated by deamination with nitrous acid (Figure 2). The negative mode FAB spectrum of the mixtures of phospholipids obtained by nitrous acid deamination of GIPLs from strain CL is shown in Figure 10. The major species was observed at m/z 795, and m/z 780 and 892 were each present at about 65% of its intensity. The signal at m/z 795 cannot be a deprotonated phosphoceramide since, being at odd mass number, it contains either zero or an even number


23

Figure 9 Electron ionization/mass spectrometry (A) and chemical ionization/mass spectrometry (B) spectra of the 1-O-hexadecyl-2,3-di-Otrimethylsilylglycerol from methanolysed glycoinositolphospholipid of T. cruzi strain CL.

of nitrogens. A plausible identification is 1-O-hexadecyl-2-O-hexadecanoyl-sn-glycero-3-phosphoinositol (calculated [M H]– ¼ 795.5), because GC/MS detected significant levels of hexadecylglycerol. Interestingly, the pattern of acyl substitution differs from that of the phosphoinositolceramides, the alkylglycerol being almost exclusively linked to hexadecanoate. In conjunction with the GC/MS results described in Section 2.5.1, the signals at m/z 780 and 892 can be assigned to deprotonated molecules of phosphoinositolceramides in

24


Figure 10 Partial negative ion fast atom bombardment/mass spectrometry spectrum of deaminated glycoinositolphospholipid of T. cruzi strain CL.

which the long chain base is sphinganine, and the fatty acyl groups are hexadecanoate and tetracosanoate respectively.

3. GPI BIOSYNTHESIS IN T. CRUZI GPI anchors are assembled in a subcompartment of the endoplasmic reticulum by sequential transfer of monosaccharides and EtNP to phosphatidylinositol, as reviewed by McConville et al. (2002, and references therein). We have shown that assembly of GPI-protein anchor precursors is broadly similar in T. cruzi and other eukaryotes (Heise et al., 1996), but that some specific details remain to be clarified. In T. cruzi GPI-anchor biosynthesis, GlcNAc is transferred from uridine diphosphate-GlcNAc to an alkyl-acyl-glycerol-containing phosphatidylinositol to yield GlcNAc-phosphatidylinositol (Heise et al., 1996). Although direct experimental evidence is lacking, it is believed that the GlcNAc-phosphatidylinositol is first


25

de-N-acetylated and four Man residues are added forming Man4GlcN-phosphatidylinositol. The Man residues are derived from dolichol-phosphomannose that is synthesized from guanosine diphosphate-Man and dolichol-phosphate (Heise et al., 1996). Finally, as in sialoglycoprotein GPI anchors (Acosta et al., 1995; Previato et al., 1995), either EtNP or AEP is transferred to the third Man residue distal to inositol forming the protein GPI-anchor precursors known as glycolipids A-like 1 and 2 (Heise et al., 1996). Acylation of the Ins ring of some intermediate(s) in the pathway may occur before addition of EtNP during the synthesis of a glycolipid C-like intermediate (Heise et al., 1994, 1996). Although all sialoglycoprotein GPI anchors (Acosta et al., 1995; Previato et al., 1995; Almeida et al., 2000) and GIPLs from various strains of T. cruzi (Carreira et al., 1996) (Table 1) have an AEP substituent on O-6 of GlcN, it is not known if AEP addition is an early or late biosynthetic event (Heise et al., 1996). GPI biosynthesis is thought to utilize endogenous phosphatidylinositol as initial acceptor. Recent investigations of L. mexicana identified two major phosphatidylinositol pools with distinctive alkyl-acyl-glycerols used for biosynthesis of GPI protein anchors or GIPLs repectively (Ralton and McConville, 1998). In T. cruzi, the protein anchor precursors are also assembled on alkyl-acyl-glycerolcontaining phosphatidylinositol (Heise et al., 1996) but, in contrast to Leishmania, the ether–lipid chains are homogeneous and are always sn-1-O-hexadecyl-2-O-acyl-glycerol. The presence of 1-O-hexadecylglycerol as the alkyl chain was observed for all T. cruzi-derived GPI protein anchors (Gu¨ther et al., 1992; Schenkman et al., 1993; Almeida et al., 1994, 2000; Acosta et al., 1995; Heise et al., 1995; Previato et al., 1995) and GIPLs so far described (Lederkremer et al., 1993; Carreira et al., 1996). In place of 1-O-hexadecyl-glycerol, T. cruzi may use sphingolipids composed of dihydrosphingosine long-chain base N-acylated with palmitic (C16:0) or lignoceric (C24:0) acids in both GPI protein anchors (Acosta et al., 1995; Heise et al., 1995; Bertello et al., 1996; Agusti et al., 1998) and GIPLs (Previato et al., 1990; Lederkremer et al., 1991; Bertello et al., 1995; Carreira et al., 1996; Uhrig et al., 1996).

26


The mechanism of ether-lipid biosynthesis in T. cruzi is currently unknown. However, in other trypanosomatids such as T. brucei and L. mexicana, the initial steps are similar to those of mammals, and follow the dihydroxyacetonephosphate (DHAP) pathway (Opperdoes, 1984; Heise and Opperdoes, 1997). The pathway of sphingolipid biosynthesis in T. cruzi is likewise unknown. In mammals and fungi, the first step in de novo sphingolipid biosynthesis is the serine-palmitoyl transferase (SPT) catalysed condensation of serine and palmitoyl-coenzymeA to yield 3-ketodihydrosphingosine, which is rapidly reduced to dihydrosphingosine (DHS) and acylated to N-acyl-sphinganine (dihydroceramide). In animals, dihydroceramide is transformed into N-acyl-sphingosine (ceramide) by introduction of a 4,5-trans double bond (Merril and Jones, 1990). The hydroxyl group at ceramide C-1 then receives a phosphocholine head group forming sphingomyelin (Merril and Jones, 1990; Dickson, 1998) or sugar residues to give various glycosphingolipids (Ichikawa and Hirabayashi, 1998). In fungi, however, dihydroceramide is hydroxylated at C-4 giving phytoceramide. Alternatively, C-4 hydroxylation of DHS forms phytosphingosine that is then transformed into phytoceramide. It is not clear whether DHS or dihydroceramide is the preferred substrate for C-4 hydroxylation in fungi (Dickson and Lester, 2002). In contrast to animals, the major head group attached to the phytoceramide C-1 hydroxyl group is phosphoinositol, forming inositolphosphorylceramide (IPC), a reaction catalysed by an IPC synthase unique to fungi (Dickson and Lester, 1999). T. cruzi resembles fungi (Lester and Dickson, 1993) in that it expresses high levels of Ins-containing sphingolipids, an important observation because it has recently been shown that the sphingolipid biosynthetic pathway is the target for several anti-fungal drugs (Nagiec et al., 1997). Although sphingolipids represent a low percentage of fungal phospholipids, they are essential for viability (Dickson and Lester, 1999). Mutants of Saccharomyces cerevisae defective in sphingolipid synthesis are non-viable and pathogenic fungi such as Cryptococcus neoformans, Candida albicans, Aspergillus fumigatus and Histoplasma capsulatum are killed by inhibitors of


27

sphingolipid biosynthesis, especially compounds which target IPC synthase, including aureobasidin A, khafrefungin and rustmicin (Takesako et al., 1993; Mandala et al., 1997, 1998). In contrast to C. neoformans and A. fumigatus, no other fungal sphingolipid biosynthetic inhibitor we tested was toxic to T cruzi. Even high concentrations (50 mg/mL) of viridiofungin A, australifungin, rustmicin, fumonisin B1 and aureobasidin A had no effect on the viability of epimastigote and metacyclic forms of T. cruzi strain Y (Lima et al., 1999; Figueiredo et al., 2001). Since the viability of T. cruzi epimastigotes was unaffected by either rustmicin or aureobasidin A, possible interpretations are that either IPC synthase is absent, or alternatively T. cruzi IPC synthase is not susceptible to these inhibitors because of differences in substrate specificity. Possibly the explanation is that fungi contain phytoceramide whereas T. cruzi does not. However, the trypanosomatids Crithidia luciliae and Herpetomonas samuelpessoai, which contain only phytoceramides in their GIPLs (Routier et al., 1993; McConville, 1996), are not affected by rustmicin (N. Heise, unpublished results). Recently, it has been shown that aureobasidin A resistance in Aspergillus species is due to increased efflux of the drug, since the minimum inhibitory concentrations were lowered 100-fold by mammalian multidrug resistance modulators (Zhong et al., 2000). Similar experiments with T. cruzi indicated that aureobasidin A was indeed toxic to the epimastigotes in a dose-dependent manner when the parasites were co-incubated with the multidrug resistance modulator verapamil (Figueiredo et al., 2001). The fact that GPI-anchored glycoproteins (Acosta et al., 1995, Heise et al., 1995) and GIPLs (Lederkremer et al., 1993) have structurally heterogeneous lipid anchors might imply the presence of at least two separate phosphatidylinositol pools for anchor biosynthesis: one containing ceramide and the other hexadecyl-glycerol. However, our previous work indicated only the presence of alkyl-acyl-glycerol in all precursors for GPI anchors in T. cruzi (see Heise et al., 1996). Studies on T. brucei and S. cerevisae demonstrated that after the GPI anchors have been synthesized the initial lipid moieties are replaced either before (T. brucei) or after (S. cerevisae) transfer of the GPI to proteins

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(Masterson et al., 1990; Sipos et al., 1994). Therefore, another hypothesis in the case of T. cruzi might imply a remodelling operation similar to the phenomenon observed with some GPI-anchored proteins (Reggiori et al., 1997; Sipos et al., 1997) and IPCs (Becker and Lester, 1980; Reggiori and Conzelmann, 1998) of S. cerevisae. In this respect, it is curious to note that in yeast ceramide is absent from the complete GPI anchor precursors (Sipos et al., 1994), a situation analogous to that reported for T. cruzi by Heise et al. (1996). Moreover, recent studies identified phospholipases from the parasite that were active both on phosphaditylinositol and IPC and could therefore be involved in remodelling processes (Bertello et al., 2000; Furuya et al., 2000; Salto et al., 2002).

4. T. CRUZI GIPL AND HOST IMMUNE RESPONSES Engagement of GPI-anchored surface proteins transmits regulatory signals to host macrophages and lymphocytes (Robinson et al., 1989; Lund-Johansen et al., 1993). In addition, several studies demonstrated that purified GPI anchors and GIPLs from protozoan parasites can modify leucocyte function (Proudfoot et al., 1995; Gomes et al., 1996; Camargo et al., 1997; Tachado et al., 1997; Kierszenbaum et al., 2002). These studies suggest that parasite glycolipids could play a role in immune regulation and pathology of protozoal infections. GIPL from T. cruzi (see DosReis et al., 2002), and GPI anchors from other protozoan parasites (Tachado et al., 1997), are bifunctional molecules that elicit differing biological effects through structurally unrelated glycan and lipid elements. This functional dichotomy contributes to the multiplicity of effects and the range of cell types affected by T. cruzi GIPL (DosReis et al., 2002). Infection of mice with T. cruzi leads to T cell immunosuppression and to B cell polyclonal activation and polyclonal immunoglobulin secretion (DosReis, 1997). We investigated the roles of T. cruzi GIPL in host T and B cell activation. Purified T. cruzi GIPL suppressed primary mouse CD4 þ and CD8 þ T cell activation in vitro, induced by T. cruzi antigen, superantigen, or anti-CD3 monoclonal antibody


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(mAb) (Gomes et al., 1996). Suppression resulted from polyclonal cell cycle arrest, and a blockade in interleukin (IL)-2 production, but no apoptosis was induced. The protein kinase C activator phorbol ester prevented suppression, ruling out non-specific toxic effects. No reduction in IL-4 production was seen, suggesting that T. cruzi GIPL targets Th1, but spares Th2, T cell function. Importantly, T. cruzi GIPL also blocked naive T cell activation in vivo. Three days after anti-CD3 mAb injection into the footpad, T cells from draining lymph nodes become responsive to recombinant IL-2, and express the IL-2R chain marker CD25. GIPL blocked both events when co-injected with anti-CD3 (Gomes et al., 1996). A control GIPL purified from Leptomonas samueli (see Previato et al., 1992) had no suppressive effect. Purified PI-glycan chain from T. cruzi GIPL had no suppressive effect on T cells, but the purified lipid moiety of T. cruzi GIPL retained all the suppressive effects of the intact molecule (Gomes et al., 1996). T. cruzi GIPL lipid moiety is mainly lignoceroylsphinganine (Previato et al., 1990), a dihydroceramide. Ceramides are pleiotropic second messengers for lymphocytes and other cell types, and are critically involved in cellular stress responses, cell cycle arrest and apoptosis (Hannun and Luberto, 2000). Therefore, GIPL appears to effect T cell activation by mimicking an intracellular ceramide mediator. In a model in vitro using a mouse T cell hybridoma, T. cruzi GIPL induced cycle arrest, but also acted as a co-stimulatory signal. It potentiated IL-2 mRNA transcription and IL-2 secretion driven by activating mAbs, by Ca2þ ionophore, and by phorbol ester (Bellio et al., 1999). Purified GIPL induced a sustained and oscillatory rise in cytosolic free Ca2þ levels, which resulted in activation of NFAT-1 (nuclear factor of activated T cells1) and nuclear translocation (Bellio et al., 1999). Purified GIPL ceramide, but not the glycan chain, reproduced the effects of the intact molecule (Bellio et al., 1999). These results indicate that T. cruzi GIPL modifies host T cell function through its ceramide moiety. While the outcome of the effect depends on the differentiation stage of the responding cell, its dominant effect on host T cells is suppressive, and also Th2-biasing. These effects could potentially help the parasite to evade the host immune response.

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Polyclonal B lymphocyte activation and immunoglobulin secretion is characteristic of T. cruzi infection (D’Imperio Lima et al., 1985). We investigated whether T. cruzi GIPL modulates B cell proliferation and immunoglobulin secretion in vitro. GIPL induced IgM production, even in the absence of a B cell proliferative response. Total immunoglobulin secretion was increased by a combination of IL-2, IL-4 and IL-5 (Bento et al., 1996). GIPL induced IgG3 secretion and co-stimulated IgG3 secretion induced by lipopolysaccharide (LPS) (Bento et al., 1996). In the presence of IL-4, secretion of IgG1 was also stimulated by GIPL. Interestingly, these stimulatory effects on B cell function were mediated by the glycan chain of GIPL, and not by the ceramide portion (Bento et al., 1996). GIPL modulated B lymphocyte activation in vivo, as measured by an increase in total serum IgM levels (Bilate et al., 2000). Moreover, B cells isolated after intravenous injection of GIPL are hyperresponsive to B cell activators, and can produce 100-fold more immunoglobulin than naive B cells (Bilate et al., 2000). GIPL activates purified B cells directly, but it can also activate B cells indirectly, through activation of NK cells. For example, B cells derived from NK and T celldeficient mice secreted immunoglobulin in vitro when stimulated with GIPL, and this response was increased by the addition of NK cells to the culture (Arruda Hinds et al., 1999). These results suggest that T. cruzi GIPL co-stimulates both B lymphocytes and NK cells, and that GIPL-activated NK cells enhance B cell responses to GIPL (Arruda Hinds et al., 1999). Together, the results suggest that GIPLs could be one class of the parasite molecules responsible for polyclonal B cell activation and antibody production in the course of Chagas disease. Macrophages and dendritic cells (DCs) are important host cells and regulators of immune responses against T. cruzi. Although intact GIPL had little or no effect on cytokine production by resting mouse macrophages, the GIPL ceramide moiety induced macrophage apoptosis in the presence of the proinflammatory cytokine interferon (IFN)-c (Freire-de-Lima et al., 1998). When macrophages are pre-treated with excess GIPL glycan chain, intact GIPL induces apoptosis in the presence of IFN-c (Freire-de-Lima et al., 1998). These results suggest that the biological effects of GIPL depend on


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the endocytic delivery of the molecule by specialized receptors, or on direct insertion into the phospholipid bilayer. Using human monocyte-derived macrophages and DCs activated by LPS or by CD40L, the roles of T. cruzi GIPL on cytokine secretion and expression of co-stimulatory molecules were investigated. Addition of GIPL reduced LPS-induced secretion of tumour necrosis factor (TNF)-, IL-10 and IL-12 by human macrophages and DCs (Brodskyn et al., 2002). GIPL downmodulated expression of CD80, CD86, and HLA-DR in LPS-stimulated macrophages, and blocked DC maturation responses to LPS and to CD40L, as measured by CD83, CD80, CD86, and HLA-DR expression (Brodskyn et al., 2002). The deactivating effects of GIPL mapped to the ceramide moiety (Brodskyn et al., 2002). Taken together, these results indicate that GIPL blocks the ability of macrophages and DCs to interact properly with T cells. In this way, effector lymphocytes would not be alarmed by the presence of T. cruzi inside host phagocytes.

5. CONCLUDING REMARKS Molecules belonging to the GPI family are ubiquitous amongst the eukaryotes and are most abundant in the protozoa, where they appear to be the major group of glycomolecules. T. cruzi strains contain a unique class of inositolphosphate-containing glycolipids. Two series of GIPL structures were identified based on the same conserved core sequence. The series 1 glycan chain is substituted at the third Man distal to Ins by EtNP, as found in T. cruzi Tulahuen strain. The PIglycan can be further substituted with a terminal (1 ! 3)-linked Galf unit, as in the G strain. The series 2 GIPL structure is present mainly in the Y and CL strains. The glycan lacks the additional phosphorus-containing group on the third Man, which instead is substituted by a single Galf residue. The lipid moiety in GIPLs from the Tulahuen, G and Y strains is predominantly ceramide, whereas that from the CL strain is a mixture of ceramide and alkyl-acylglycerol species. The function of the T. cruzi GIPL is unknown, however the unusual structural features of these molecules may

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indicate a role in parasite–host interactions, as a virulence determinant and/or as a modulator of the host’s immune response. T. cruzi GIPLs suppress Th1 responses, block DC maturation and reduce surface display of co-stimulatory molecules and of major histocompatibility complex class II in host macrophages. These effects are beneficial to the parasite and detrimental to the host immune response. Furthermore, these biological effects are mediated by the ceramide moiety. The glycan chain, conversely, co-stimulates B cells and polyclonal immunoglobulin production, which could be important for autoantibody production. Overall, the results suggest an important role for T. cruzi GIPLs in the pathogenesis of Chagas disease. Future studies need to be directed towards a better understanding of the biosynthesis of these T. cruzi-specific molecules. Disruption of this pathway could provide a route to novel anti-T. cruzi drugs which could reduce the severity of Chagas disease, if not eliminate the parasite completely. ACKNOWLEDGEMENTS This work was supported by grants from the Conselho Nacional de Desenvolvimento Cientı´ fico e Tecnolo´gico (CNPq/Pronex/Padct), Conselho de Ensino para Graduados (CEPG/UFRJ), Fundaçaõ Universita´ria Jose´ Bonifaćio, and Fundaçaõ de Amparo a` Pesquisa do Estado do Rio de Janeiro (FAPERJ). G.A.D.R. and L.M.P. are Howard Hughes International Research Scholars. We thank Mrs Lucy Jacinto do Nascimento and Mr Orlando Augusto Agrellos for their excellent technical assistance, and Dr Suzanne Mandala (Merck, USA) for gifts of viridiofungin A, australifungin and rustmicin. R.W. thanks the Centre for Applied Microbiology and Research, Porton, UK, where some of these studies were carried out. REFERENCES Acosta, A.S., Schenkman, S., Yoshida, N., Mehlert, A., Richardson, J.M. and Ferguson, M.A.J. (1995). The lipid structure of the


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Mayor, S., Menon, A.K. and Cross, G.A. (1990). Glycolipid precursors for the membrane anchor of Trypanosoma brucei variant surface glycoproteins. II. Lipid structures of phosphatidylinositol-specific phospholipase C sensitive and resistant glycolipids. Journal of Biological Chemistry 265, 6174–6181. McConville, M.J. (1996). Glycosyl-phosphatidylinositols and the surface architecture of parasitic protozoa. In: Molecular Biology of Parasitic Protozoa (D.F. Smith and M. Parsons, eds.), pp. 205–228. Oxford: IRL Press. McConville, M.J. and Ferguson, M.A.J. (1993). The structure, biosynthesis and function of glycosylated phosphatidylinositols in the parasitic protozoa and higher eukaryotes. Biochemical Journal 294, 305–324. McConville, M.J., Homans, S.W., Thomas-Oates, J.E., Dell, A. and Bacic, A. (1990). Structures of the glycoinositolphospholipids from Leishmania major. A family of novel galactofuranose-containing glycolipids. Journal of Biological Chemistry 265, 7385–7394. McConville, M.J., Mullin, K.A., Ilgoutz, S.C. and Teasdale, R.D. (2002). Secretory pathway of trypanosome parasites. Microbiology Molecular Biological Review 66, 122–154. McLafferty, F.W. and Turecek, F. (1993). Interpretation of Mass Spectra, edition 4. Mill Valley: University Science Books. Melo, R.C. and Brener, Z. (1978). Tissue tropism of different Trypanosoma cruzi strains. Journal of Parasitology 64, 475–482. Mendonça-Previato, L. and Previato, J.O. (1985). Glycoconjugates from epimastigotes of Trypanosoma cruzi. Anais da Academia Brasileira de ˆ Ciencias 37, 135–136. Mendonça-Previato, L., Gorin, P.A.J., Braga, A.F., Scharfstein, J. and Previato, J.O. (1983). Chemical structure and antigenic aspects of complexes obtained from epimastigotes of Trypanosoma cruzi. Biochemistry 22, 4980–4987. Menon, A.K. (1994). Structural analysis of glycosylphosphatidylinositol anchors. Methods in Enzymology 230, 418–442. Merril, A.H. and Jones, D.D. (1990). An update of the enzymology and regulation of sphingomyelin metabolism. Biochimica et Biophysica Acta 1044, 1–12. Moncayo, A. (1999). Progress towards the elimination of transmission of Chagas’ disease in Latin America. Memo´rias do Instituto Oswaldo Cruz 94, 401–404. Murta, S.M., Gazzinelli, R.T., Brener, Z. and Romanha, A.J. (1998). Molecular characterization of susceptible and naturally resistant strains of Trypanosoma cruzi to benznidazole and nifurtimox. Molecular and Biochemical Parasitology 93, 203–214.


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Myher, J.J., Marai, L. and Kuksis, A. (1974). Identification of monoacyland monoalkylglycerols by gas–liquid chromatography–mass spectrometry using polar siloxane liquid phases. Journal of Lipid Research 15, 586–592. Nagiec, M.M., Nagiec, E.E., Baltisberger, J.A., Wells, G.B., Lester, R.L. and Dickson, R.C. (1997). Sphingolipid synthesis as a target for antifungal drugs: complementation of the IPC synthase defect in a mutant strain of Saccharomyces cerevisiae by the AUR1 gene. Journal of Biological Chemistry 272, 9809–9817. Opperdoes, F.R. (1984). Localization of initial steps in alkoxyphospholipid biosynthesis in glycosomes. FEBS Letters 169, 35–39. Previato, J.O., Gorin, P.A.J., Mazurek, M., Xavier, M.T., Fournet, B., Wieruszesk, J.M. and Mendonça-Previato, L. (1990). Primary structure of the oligosaccharide chain of lipopeptidophosphoglycan of epimastigotes. Journal of Biological Chemistry 265, 2518–2526. Previato, J.O., Mendonça-Previato, L., Jones, C., Wait, R. and Fournet, B. (1992). Structural characterization of a novel class of glycophosphosphingolipids from the protozoan Leptomonas samueli. Journal of Biological Chemistry 267, 24279–24286. Previato, J.O., Wait, R., Jones, C. and Mendonça-Previato, L. (1994). Structural analysis of novel rhamnose-branched oligosaccharides from the glycophosphosphingolipids of Leptomonas samueli. Glycoconjugate Journal 11, 23–33. Previato, J.O., Jones, C., Xavier, M.T., Wait, R., Travassos, L.R., Parodi, A.J. and Mendonça-Previato, L. (1995). Structural characterization of the major GPI membrane anchored glycoprotein of epimastigote forms of Trypanosma cruzi. Journal of Biological Chemistry 270, 7241–7250. Previato, J.O., Jones, C., Wait, R., Routier, F., Saraiva, E. and MendonçaPreviato, L. (1997). Leishmania adleri, a lizard parasite, expresses structurally similar glycoinositolphospholipids to mammalian Leishmania. Glycobiology 7, 687–695. Proudfoot, L., O’Donnell, C.A. and Liew, F.Y. (1995). Glycoinositolphospholipids of Leishmania major inhibit nitric oxide synthesis and reduce leishmanicidal activity in murine macrophages. European Journal of Immunology 25, 745–750. Ralton, J.E. and McConville, M.J. (1998). Delineation of three pathways of GPI biosynthesis in Leishmania mexicana: precursors from different pathways are assembled on distinct pools of PI and undergo fatty acid remodeling. Journal of Biological Chemistry 273, 4245–4257. Reggiori, F. and Conzelmann, A. (1998). Biosynthesis of inositol phosphoceramides and remodeling of GPI anchors in S. cerevisae are mediated by different enzymes. Journal of Biological Chemistry 273, 30550–30559.

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Reggiori, F., Canivenc-Gansel, E. and Conzelmann, A. (1997). Lipid remodelling leads to the introduction and exchange of defined ceramides on GPI proteins in the ER and Golgi of Saccharomyces cerevisiae. EMBO Journal 16, 3506–3518. Robinson, P.J., Millrain, M., Antoniou, J., Simpson, E. and Mellor, A.L. (1989). A glycophospholipid anchor is required for Qa-2-mediated T cell activation. Nature 342, 85–87. Routier, F., Previato, J.O., Jones, C., Wait, R. and Mendonça-Previato, L. (1993). Glycoinositolphospholipids from members of the Trypanosomatidae family: investigation on the lipid moiety. Journal of Brazilian Association and Advance of Science 45, 66–68. Routier, F.H., Xavier da Silveira, E., Wait, R., Jones, C., Previato, J.O. and Mendonça-Previato, L. (1995). Chemical characterization of glycosylinositolphospholipids of Herpetomonas samuelpessoai. Molecular and Biochemical Parasitology 69, 81–92. Salto, M.L., Furuya, T., Moreno, S.N., Docampo, R. and Lederkremer, R.M. (2002). The phosphatidylinositol-phospholipase C from Trypanosoma cruzi is active on inositolphosphorylceramide. Molecular and Biochemical Parasitology 119, 131–133. Schenkman, S., Ferguson, M.A.J., Heise, N., Cardoso de Almeida, M.L., Mortara, R.A. and Yoshida, N. (1993). Mucin-like glycoproteins linked to the membrane by glycosylphosphatidylinositol anchor are the major acceptors of sialic acid in a reaction catalyzed by trans-sialidase in metacyclic forms of Trypanosoma cruzi. Molecular and Biochemical Parasitology 59, 293–304. Schneider, P. and Ferguson, M.A.J. (1995). Microscale analysis of glycosylphosphatidylinositol structures. Methods in Enzymology 250, 614–630. Sipos, G., Puoti, A. and Conzelmann, A. (1994). GPI membrane anchors in Saccharomyces cerevisae: absence of ceramides from complete precursor glycolipids. EMBO Journal 13, 2789–2796. Sipos, G., Reggiori, F., Vionnet, C. and Conzelmann, A. (1997). Alternative lipid remodelling pathways for GPI membrane anchors in Saccharomyces cerevisae. EMBO Journal 16, 3494–3505. Tachado, S.D., Gerold, P., Schwarz, R., Novakovic, S., McConville, M. and Schofield, L. (1997). Signal transduction in macrophages by glycosylphosphatidylinositols of Plasmodium, Trypanosoma and Leishmania: activation of protein tyrosine kinases and protein kinase C by inositolglycan and diacylglycerol moieties. Proceedings of the National Academy of Sciences of the USA 94, 4022–4027. Takesako, K., Kuroda, H., Inque, T., Haruna, F., Yoshikawa, Y. and Kato, I. (1993). Biological properties of aureobasidin A, a cyclic depsipeptide antifungal antibiotic. Journal of Antibiotics 46, 1414–1420.


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Turco, S.J. and Descoteaux, A. (1992). The lipophosphoglycan of Leishmania parasites. Annual Review of Microbiology 46, 65–94. Uhrig, M.L., Couto, A.S., Colli, W. and Lederkremer, R.M. (1996). Characterization of inositolphospholipids in Trypanosoma cruzi trypomastigote forms. Biochimica et Biophysica Acta 1300, 233–239. WHO (2000). Chagas’ disease, Brazil. Weekly Epidemiology Record 75, 153–155. Winter, G., Fuchs, M., McConville, M.J., Stierhof, Y.D. and Overath, P. (1994). Surface antigens of Leishmania mexicana amastigotes: characterization of glycoinositol phospholipids and a macrophage-derived glycosphingolipid. Journal of Cell Science 107, 2471–2482. Xavier da Silveira, E., Jones, C., Wait, R., Previato, J.O. and MendonçaPreviato, L. (1998). Glycoinositol phospholipids from Endotrypanum species express epitopes in common with saccharide side chains of the lipophosphoglycan from Leishmania major. Biochemical Journal 329, 665–673. Zhong, W., Jeffries, M.W. and Georgopapadakou, N.H. (2000). Inhibition of inositol phosphorylceramide synthase by aureobasidin A in Candida and Aspergillus species. Antimicrobial Agents and Chemotherapy 44, 651–653.


Biodiversity and Evolution of the Myxozoa Elizabeth U. Canning1 and Beth Okamura2 1

Department of Biological Sciences, Imperial College London, Silwood Park Campus, Ascot, Berkshire, SL5 7PY, UK; 2 School of Animal and Microbial Sciences, University of Reading, Whiteknights, Reading, Berkshire, RG6 6AJ, UK Abstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Classification of the Phylum Myxozoa. . . . . . . . . . . . . . . . 3. Diagnostic Characters of Myxozoa. . . . . . . . . . . . . . . . . . 3.1. Biology and Life Cycles . . . . . . . . . . . . . . . . . . . . 3.2. Hosts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4. Diagnostic Characters of the Class Myxosporea . . . . . . . . . . 4.1. Life Cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5. Diagnostic Characters of the Class Malacosporea . . . . . . . . . 5.1. Buddenbrockia plumatellae Schro¨der, 1910 . . . . . . . . . 5.2. Malacosporean Stages in Carp, Cyprinus carpio . . . . . . 5.3. Tetracapsuloides bryosalmonae (Canning, Curry, Feist, Longshaw and Okamura, 1999), the PKX Organism 6. Phylogenetic Relationships of Myxozoa. . . . . . . . . . . . . . . 6.1. Inferred Higher Level Phylogeny . . . . . . . . . . . . . . . 6.2. Myxozoan Polar Capsules and Cnidarian Nematocysts . . 7. Myxozoan Lineages and Evolution of Parasitism. . . . . . . . . . 7.1. Morphological Simplification due to Parasitism . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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ABSTRACT Myxozoans (phylum Myxozoa) are metazoan parasites utilizing invertebrate and (mainly) aquatic vertebrate hosts. They have in common with cnidarians the possession of virtually identical, highly ADVANCES IN PARASITOLOGY VOL 56 ISSN: 0065308X DOI: 10.1016/S0065-308X(03)56002-X


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ELIZABETH U. CANNING AND BETH OKAMURA

complex organelles, namely the polar capsules in myxozoan spores, serving for attachment to new hosts and the nematocysts in surface epithelia of cnidarians, serving for food capture. Although myxozoan spores are multicellular, the simple trophic body forms of almost all species, reduced to syncytial plasmodia or single cells, reveal no clues to myxozoan ancestry or phylogenetic relationships. The myxozoan genus Buddenbrockia is one of only two known genera belonging to a clade which diverged early in the evolution of the Myxozoa. Today the Myxozoa are represented by two classes, the Myxosporea, containing all the better-known genera, which alternate between fish and annelids, and the Malacosporea, containing Buddenbrockia and Tetracapsuloides, parasitising bryozoans. The latter genus also infects salmonid fish, causing proliferative kidney disease (PKD). The enigmatic Buddenbrockia has retained some of its ancestral features in a body wall of two cell layers and a worm-like shape, maintained by four longitudinally-running muscle blocks, similar to a gutless nematode and suggestive of a bilaterian ancestry. Although some analyses of 18S rDNA sequences tend towards a cnidarian (diploblast) affinity for myxozoans, the majority of these studies place them within, or sister to, the Bilateria. The latter view is supported by their possession of central class Hox genes, so far considered to be synapomorphic for Bilateria. The simple body form is, therefore, an extreme example of simplification due to parasitism. Various hypotheses for the occurrence of identical complex organelles (nematocysts and polar capsules) in diploblast and triploblast phyla are evaluated: common ancestry, convergent evolution, gene transfer and, especially, endosymbiosis. A theory of the evolution of their digenetic life cycles is proposed, with the invertebrate as primary host and secondary acquisition of the vertebrate host serving for asexual population increase.

1. INTRODUCTION Myxozoans are best known as parasites of poikilothermic vertebrates, especially fish, but, since the discovery of tubificid annelids

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as alternate hosts for Myxobolus cerebralis by Markiw and Wolf (1983) and Wolf and Markiw (1984), it has been established that there may be an obligatory or facultative invertebrate host in most, if not all, myxozoan life cycles. The discovery of the role of invertebrates has raised important questions about the evolution of myxozoans, especially with respect to their phylogeny and whether they originated in aquatic invertebrates and spread to fish or vice versa. The most characteristic feature of myxozoans are organelles known as polar capsules, which are rarely present in somatic cells but are omnipresent in spores. Each polar capsule contains a coiled, eversible polar filament which, on coming into contact with a suitable host, is everted to anchor the spore and allow the infective agent to emerge from the spore valves and enter the host. Spores were first recorded from fish by Mu¨ller (1841) who called them Psorospermien. By the end of the 19th century spore development, by endogeny within plasmodia, had been observed in many different species and it became clear that several cells contributed to spore formation. The multicellular nature of spores led Sˇtolc (1899), Emery (1909) and Ikeda (1912) to propose that myxozoans were in fact Metazoa, although they had previously been classified as Protozoa. Myxozoan spores differ fundamentally from the environmentally resistant stages (cysts or spores) of protistan organisms, including those of the former taxon Protozoa. In protists the cyst or spore wall is a non-living protective coat, secreted around itself by the living cell, which may or may not divide within it before emerging to infect a new host. In myxozoan spores, the infective stage (sporoplasm) is surrounded by several living cells which play a protective role and are discarded when infection is achieved by the emergent sporoplasm. Attention was again drawn to the multicellularity of myxozoan spores by Weill (1938), who made the comparison between myxozoan polar capsules and coelenterate (cnidarian) nematocysts. The remarkable similarity between polar capsules and nematocysts during development and in their mature structure, suggesting a cnidarian origin for myxozoans, has dominated phylogenetic studies of myxozoans in recent years. We address this issue using the most recent ultrastructural and molecular data. Although myxozoans were

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accorded the status of phylum Myxozoa by Grasse´ (1970), many texts continued to classify them as Protozoa. There is now compelling evidence that this needs to be rejected once and for all and their true status as a metazoan phylum recognized, as proposed previously by Grell (1956) and Lom (1969). When actinosporean parasites of oligochaete annelids were identified as stages in the life cycle of myxozoans (Markiw and Wolf, 1983), the class Actinosporea was suppressed, leaving a single class Myxosporea in the phylum (Kent et al., 1994). However, a new class Malacosporea was established (Canning et al., 2000) after the enigmatic myxozoan known as the PKX organism was shown by phylogenetic analyses of 18S rDNA sequences to be a new species of Tetracapsula, the single known genus representing a distinct clade that had diverged early in myxozoan evolution before the radiation that had given rise to the numerous better known genera (Anderson et al., 1999a,b). The Malacosporea, with the single order Malacovalvulida and single family Saccosporidae, were characterized by soft-walled spores, special sporoplasmosomes with a bar-like invagination, bryozoans as invertebrate hosts, and spore formation within a sac-like body form. This review is mainly concerned with myxozoans of the class Malacosporea, and we discuss how knowledge of the Malacosporea has thrown light on the origins of myxozoans, both in terms of their phylogenetic relationships and the evolution of their life cycles. However, differentiation from the class Myxosporea requires that background data on the phylum and both classes be included.

2. CLASSIFICATION OF THE PHYLUM MYXOZOA The system of classification adopted in this review is based on that of Kent et al. (2000b) with the addition of the Class Malacosporea. Phylum: Myxozoa Grasse´, 1970. Class: Myxosporea Bu¨tschli, 1881. Order: Bivalvulida Schulman, 1959.


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Suborders: Sphaeromyxina Lom and Noble, 1984 with one family and one genus. Variisporina Lom and Noble, 1984 with ten families and 33 genera. Platysporina Kudo, 1919 with one family and eleven genera. Order: Multivalvulida Schulman, 1959 with six families and seven genera. Class: Malacosporea Canning, Curry, Feist, Longshaw and Okamura, 2000. Order: Malacovalvulida Canning, Curry, Feist, Longshaw and Okamura, 2000. Family: Saccosporidae Canning, Okamura and Curry, 1996. Genus: Buddenbrockia Schro¨der, 1910. Genus: Tetracapsuloides Canning, Tops, Curry, Wood and Okamura, 2002.

3. DIAGNOSTIC CHARACTERS OF THE PHYLUM MYXOZOA 3.1. Biology and Life Cycles All Myxozoa are metazoan parasites of vertebrates and invertebrates, many life cycles involving an alternation between the two. Cell organization is typically eukaryotic with nucleus, endoplasmic reticulum and Golgi cisternae. Mitochondria tend to have tubular rather than plate-like cristae and, unusually for metazoans, centrioles are not involved in nuclear division. Cilia are absent. Myxozoans are commonly extracellular and may be coelozoic or histozoic. When histozoic, the trophic phases often lie in an intercellular position and, even when they grow to macroscopic plasmodia, many lie in spaces between compressed host cells. However, numerous species have truly intracellular stages (Stehr and Whitaker, 1986; Lom et al., 1989; Yokoyama, 2002). Extrasporogonic stages (trophozoites, trophic stages) may be pseudoplasmodia (uninucleate), syncytial plasmodia (multinucleate), or organized as a cellular layer around a syncytium or as hollow sacs

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or vermiform stages. Growth of plasmodia is accompanied by an increase in the number of nuclei, some somatic, some generative. Generative nuclei are destined for spore formation. Although cell division by binary fission is known, particularly in the bryozoan phase of malacosporean myxozoans, where cells form into tissues proliferation by endogeny is highly characteristic of the phylum. After nuclear division, one of the nuclei and its surrounding cytoplasm become enveloped by endoplasmic reticulum, so that a secondary cell is formed. The secondary cell, limited by its plasma membrane, lies in a membrane-bound vacuole in the primary cell. Endogeny may be taken to tertiary or quaternary levels and is responsible for proliferation when two or more endogenous cells are formed within the cytoplasm of another and are released by breakdown of the parent cell. In the vertebrate host myxosporean sporogony may be initiated in plasmodia or pseudoplasmodia. In plasmodia, endogenous cells formed around generative nuclei are of two types: sporogonic cells which give rise to spores, and pericytes which surround the sporogonic cells and presumably assist in the nutrition of the developing sporogonic cells. The sporogonic and pericytic cells are collectively known as a pansporoblast. In pseudoplasmodia the primary cell is the equivalent of the pericyte and functions as a pansporoblast. Plasmodia produce many spores (polysporic) and pseudoplasmodia normally produce two spores (disporic). Each sporogonic cell divides to give the requisite number of capsulogenic and valvogenic cells protecting one or two sporoplasms. In most genera spores harbour a single binucleate sporoplasm but in a few cases (e.g. Sphaerospora, Unicapsula) there are two uninucleate sporoplasms. Morphogenesis of the polar capsules occurs in the capsulogenic cells. Typically, a bulb-like capsular primordium is formed together with an external tube, both surrounded by a corset of longitudinally running microtubules. The polar filament is assembled in the external tube and, as the tube is gradually resorbed, the polar filament enters the capsule as a folded or coiled structure, with its base attached to the mouth of the capsule, which is sealed. Junctions between capsulogenic and valvogenic cells are complexes of septate or


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adherens junctions and gap junctions. The type of spore formed in vertebrate hosts is known as myxosporean (Figure 1) and the cycle of development as outlined above is known as the myxosporean phase. All cells in the myxosporean phase of myxozoans are believed to be diploid (El-Matbouli et al., 1998). The sporogonic sequence of malacosporeans in vertebrates is not known. In annelids the cycle of development is known as the actinosporean phase. After proliferation, a sexual cycle within pericytes culminates in spore formation. The actinosporean spores (actinospores) differ from myxosporean spores in having a tri-radial symmetry with three polar capsules and three valves, which often have elongated or bulbous extensions (Figure 2). Also, the sporoplasm is multicellular. In bryozoans, sac-like or vermiform stages have inner and outer cell layers. The inner layer proliferates to form a population of cells in the central cavity, some of which divide or aggregate to make sets of cells which differentiate into the four polar capsules, four valves and two sporoplasms of malacosporean spores (Figure 3). There are no pericytes or pansporoblasts. Nuclear division by meiosis occurs in the invertebrate phase of both myxosporean and malacosporean cycles. The developmental sequence following meiosis differs between the classes. In the myxosporeans the spores are formed from zygotes and all cells of the spore are diploid, whereas in the malacosporeans fusion of nuclei has not been observed and the restoration of diploidy is presumed to occur in the next host. When polar capsules and/or endogeny are found in a new organism a diagnosis of Myxozoa must be considered, although endogeny without polar capsules might also be indicative of organisms in the phylum Paramyxea (see Desportes and Perkins, 1990).

3.2. Hosts 3.2.1. Invertebrates Alternation of hosts in myxosporean life cycles was first reported by Markiw and Wolf (1983) and Wolf and Markiw (1984) for

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pc

1

2

S

S 3


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M. cerebralis, the parasite responsible for ‘whirling disease’ in trout and, since then, alternation of hosts has been described for other freshwater myxosporeans. Known invertebrate hosts are mostly aquatic oligochaete annelids with more than 25 species now linked with myxosporeans infecting fish (summarized by Kent et al., 2001). A freshwater polychaete, Manayunkia speciosa, is involved in the life cycle of Ceratomyxa shasta, a parasite of salmonid fish (Bartholomew et al., 1997). Actinosporean stages have been found in numerous marine oligochaetes but none of these has yet been linked to a particular myxosporean cycle (Hallett and Lester, 1999; Hallett et al., 1999, 2001). Also in the marine environment a sipunculid worm, Nephrostoma ( ¼ Petalostoma) minuta, has been listed as the host of Tetractinomyxon intermedium and Tetractinomyxon irregulare (see Ikeda, 1912), and tetractinomyxon stages have been identified in the polychaetes Nereis diversicolor (see Køie, 2000), Spirorbis spirorbis and Hydroides norvegica (see Køie, 2002), but fish hosts for these actinosporeans have not been identified. In the single malacosporean species, Tetracapsuloides bryosalmonae, for which two hosts are known, bryozoans are utilized as invertebrate hosts (Anderson et al., 1999a,b; Okamura et al., 2001; Okamura and Wood, 2002). Bryozoans also harbour the malacosporean Buddenbrockia plumatellae (syn. Tetracapsula bryozoides) but a vertebrate host is not known for this species (Canning et al., 2002; Morris et al., 2002; Okamura et al., 2002a). Myxozoans have been reported from a few other invertebrates. Thus, Chloromyxum diploxys, in the lepidopteran Tortrix viridana (see The´lohan, 1895), is probably a myxozoan, although unlikely to be a species of Figures 1–3 Comparison of myxozoan spores. Figure 1 Myxosporean-type spore represented by Myxobolus ellipsoides from chub, Leuciscus cephalus, showing two polar capsules (pc) and specific shape maintained by strengthened valves. Bar ¼ 10 mm. Photograph provided by Dr M. El-Matbouli. Figure 2 Actinosporean-type spore represented by Myxobolus cerebralis from Tubifex sp. showing position of sporoplasm (arrow) with germ cells, internal to three polar capsules (arrowhead) and extensions of the valves conferring triradial symmetry. Bar ¼ 50 mm. Photograph provided by Dr M. El-Matbouli. Figure 3 Malacosporean spores represented by Buddenbrockia plumatellae from Cristatella mucedo, showing either four polar capsules (arrows) or two sporoplasms (s), depending on the plane of focus. The spherical shape is maintained by turgor pressure, as the valve cells are not strengthened. Bar ¼ 10 mm. Photograph provided by Ms Sylvie Tops.

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Chloromyxum. Myxobolus sp. in the polychaete worm Nais lacustris (see Gurley, 1894) is unusual in forming myxosporeantype, rather than actinosporean-type, spores in an invertebrate. Trematodes have twice been found infected with myxosporean-type spores, apparently in the absence of myxozoan infections in the fish harbouring the flatworms: both of these myxozoan species, Fabespora vermicola (see Overstreet, 1976) and Fabespora sp. (see Siau et al., 1981), were parasites of trematodes in fish belonging to the family Sparidae. 3.2.2.

Vertebrates

By far the commonest vertebrate hosts of Myxosporea are teleost fish but some species are known from amphibians (see, e.g., Upton et al., 1992 and McAllister et al., 1995) and chelonid reptiles (see Lom, 1990). Although it has long been considered that myxozoans are restricted to poikilothermic vertebrates, recent data suggest that birds and mammals may also be susceptible. Friedrich et al. (2000) described a myxozoan-like parasite causing xenoma formation (gross host cell hypertrophy) in the brains of moles, Talpa europaea. The nuclei of the host capillary pericytes were encircled by primary parasitic cells, each with secondary cells and, in two cases, with tertiary cells (Figure 4). In one giant host cell, a primary parasite cell contained 42 secondary cells. No spore was seen, so that a firm assignment to the Myxozoa was not possible but, as stated above, endogenous proliferation is an almost uniquely myxozoan character. The presence of myxozoan spores in human faeces has been reported several times (McClelland et al., 1997; Boreham et al., 1998; Lebbad and Willcox, 1998; Moncada et al., 2001). In most cases, the abdominal pain and/or diarrhoea for which the patients were investigated was attributed to other causes and the myxozoan spores were thought to have come from eating fish and to have passed unchanged through the intestine. The patient described by Moncada et al. (2001) was infected with human immunodeficiency virus (HIV) and the diarrhoea had persisted for 18 months. Myxobolus-like


53

cp

4

Figure 4 Numerous cells of a myxozoan-like parasite undergoing endogeny in a grossly enlarged capillary pericyte (cp) in the brain of the European mole, Talpa europaea. Bar ¼ 10 mm. Reproduced by permission of Cambridge University Press from Friedrich et al., 2000, Parasitology 121, 483–492.

spores were found first in association with the coccidium Isospora belli (which was presumed to be the cause of the diarrhoea) and again two months later. The persistence of the spores suggested to the authors that there was a true infection. The immunocompromised status of the patient might have permitted infection but the source is obscure unless the infection came directly from fish, contrary to known life cycles. True infections have recently been found in birds. Lowenstine et al. (2002) reported six cases in anatid ducks collected from

54


enclosures in zoological gardens. Developmental stages and mature myxosporean-type spores were found in inflammatory lesions in the bile ducts and in hepatic parenchyma after liberation from ruptured bile ducts. The ability of myxozoans to survive in these homeothermic vertebrates suggests that the host range could well include mammals and awareness of this possibility might eventually establish whether or not humans are truly susceptible. A life cycle incorporating anatid ducks and aquatic invertebrates is concordant with the known pattern of myxozoan life cycles. However, a life cycle involving a wholly terrestrial vertebrate such as the mole would presumably involve a terrestrial invertebrate such as an earthworm, probably without release of spores until death of the host, and it is difficult to see how an infection in humans could be part of a continuing cycle.

4. DIAGNOSTIC CHARACTERS OF THE CLASS MYXOSPOREA 4.1. Life Cycle The class comprises about 55 genera and over 1300 species. Alternation of hosts is known for about 25 species (see Kent et al., 2001) and it is likely that these species reflect the general life cycle plan for the majority, divided into a myxosporean phase in the vertebrate and actinosporean phase in the invertebrate. Myxosporean spores are infective to annelids and actinosporean spores are infective to fish. Annelids and sipunculids, but not bryozoans, serve as alternate hosts. So far, direct transmission between fish, using natural routes as opposed to inoculation, has been established experimentally for three species only, all being intestinal parasites. Diamant (1997) achieved infection of Myxidium leei in specific pathogen-free sea bream (Sparus auratus) by co-habitation with infected fish, exposure to water discharged from a tank of infected fish and gavage of infected gut tissue. Highly successful transmission was effected in a similar series of experiments with Enteromyxum scophthalmi, an enteric


55

parasite of turbot, Scophthalmus maximus (see Redondo et al., 2002) and with Myxidium fugu and Myxidium sp. TP, parasites of tiger puffer fish, Takifugu rubripes (see Yasuda et al., 2002). Assuming that myxosporean spores are infectious to invertebrates only, the investigators in these studies proposed that vegetative stages were the sources of infection to the experimental fish. The trophic plasmodia would have had to survive passage through stomach and small intestine to reach the posterior intestinal mucosa if ingested or administered by canula to the stomach. In the case of M. fugu, the stomach should not be a problem as it can actually be a site of infection, albeit rarely. The trophic stages would also have to survive exposure to sea water in the effluent transmission experiment. The direct transmission between fish does not preclude the possibility of a facultative actinosporean stage in an alternate host but might represent an evolutionary short cut, i.e., a reduction in life cycle complexity by omitting what was probably the primary host, as occurs with some trematodes (see Poulin and Cribb, 2002). So far, a complete life cycle involving two hosts has not been demonstrated for any myxozoan parasitizing marine fish, although fish hosts for species undergoing actinosporean development in polychaetes must certainly exist. 4.1.1. Characters of the Myxosporean Phase Trophic stages are pseudoplasmodia or plasmodia but not hollow sacs or worm-like forms. Coelozoic forms may be attached uninucleate pseudoplasmodia or free multinucleate plasmodia. Attachment of pseudoplasmodia to the lining of hollow organs may be by the insertion of long projections between host cells (e.g. Myxidium trachinorum; Figure 5) (see Canning et al., 1999a), by interdigitation of cytoplasmic processes with host cell villi (e.g. Hoferellus gilsoni; Trouillier et al., 1996), by surfaces flattened against the host cells (e.g. Myxidium gadi; Feist, 1995), or by means of septate junctions between them (e.g. Zschokkella pleomorpha; Lom and Dykova´, 1996). After proliferation by endogeny, pseudoplasmodia themselves become pansporoblasts, generating two sporogonic cells which differentiate into spores. Multinucleate coelozoic forms, which may increase in

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5

p

6

Figure 5 Myxidium trachinorum: pseudoplasmodia attached by filose processes to gallbladder epithelium of lesser weever fish, Echiichthys vipera. Reproduced by permission of Springer-Verlag from Canning et al., 1999, Parasitology Research 85, 910–919. Bar ¼ 5 mm. Figure 6 Part of a plasmodium of Henneguya striolata from gill of black tailed piranha, Serrasalmus striolatus, showing an external zone of trophozoites and early sporogonic stages (arrows) within the plasma membrane (p) and an internal zone of maturing and mature spores. Bar ¼ 5 mm. Micrograph provided by Dr Graça Casal and Professor C. Azevedo.


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sc

sg

7

Figure 7 Superficial zone of a large plasmodium of Myxobolus pendula infecting the gill of creek chub, Semotilus atromaculatus. A palisade layer of secretory cells (sc) separates a zone containing the sporogonic cells (sg) from an extracellular matrix (arrow). Bar ¼ 10 mm. Reproduced by permission of the Society of Protozoologists from Martyn et al., 2002, Journal of Eukaryotic Microbiology 49, 175–182.

number by plasmotomy, usually have an ectoplasmic layer free of organelles. In sporogony, one of two endogenously formed cells surrounds the other. The outer cell is the pericyte which becomes the pansporoblast and the inner cell is the sporogonic cell which will divide further into the requisite number of cells to form (usually) two spores (disporic sporogony). Histozoic forms are often macroscopic plasmodia (Figure 6) enclosed by reactive host tissue consisting of fibroblasts and collagen. Most histozoic plasmodia, like the coelozoic plasmodia, are syncytial, with non-nucleated ectoplasm surrounding endoplasm harbouring somatic and generative nuclei. However, the plasmodia of Myxobolus pendula (Figure 7) have the central, syncytial endoplasm surrounded by a palisade layer of secretory cells (Martyn et al., 2002). This is an interesting body form which may represent a stage along the

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degenerative path from the triploblast structure of the ancestral myxozoans (represented today only by the malacosporeans) to completely syncytial plasmodia. Some myxosporeans undergo a period of proliferation in body organs other than the final sporogonic site. Thus, periods of extrasporogonic proliferation of Sphaerospora renicola occur in the blood (Csaba, 1976; Lom et al., 1983), swim bladder (Dykova´ et al., 1990) and kidney tubule epithelium of common carp, Cyprinus carpio (see Lom et al., 1982), before sporogony in the kidney tubule lumina (Molna´r and Kovaćs-Gayer, 1986). Development in organs other than the final sporogonic site is likely to be common in myxosporean life cycles, accounting for an increase in numbers before the sporogonic phase is initiated, as is known for other species of Sphaerospora and in the migration of M. cerebralis from peripheral sites of invasion to the cartilage (El-Matbouli et al., 1995). Myxosporean spores have 2–7 valves and 1–7 polar capsules (Figure 1), according to genus, and there is usually a single sporoplasm. The valvogenic cells become greatly attenuated with little, if any, residual cytoplasm but are strengthened by secreted material which forms the mature valves and maintains the shape characteristic of each species. The valves may be smooth or ridged, of simple outline or bear a variety of broad or filamentous appendages and are typically thickened along the sutural line. The apices of the polar capsules are subterminal in the spore and are enclosed by the valves (Figure 8). The opening in the capsule wall is sealed with a plug which may extend over the anterior face of the capsule. On discharge, the filament must break not only through the capsule plug but also the valve, and the position of a potential orifice has been identified in the valve close to the sutural ridge (El-Matbouli et al., 1990). In almost all myxosporean genera the spore encloses a single sporoplasm which may be binucleate or uninucleate. In Sphaerospora there are two separate uninucleate sporoplasms, and in Kudoa and Unicapsula one envelops the other. Two uninucleate sporoplasms have sometimes been reported in species of Leptotheca, Ceratomyxa and Chloromyxum, where they may represent stages prior to, or derived from, the more common binucleate sporoplasm.


59

v

st 9

I

8

10

Figures 8–10 Comparison of myxozoan polar filament extrusion points from polar capsules. Figure 8 Myxosporean-type spore of Myxobolus sp.: extrusion point closed with a stopper (st), positioned well inside the spore valves (v). Bar ¼ 1.0 mm. Reproduced by permission of the Society of Protozoologists from Desser and Brockley Patterson, 1978, Journal of Protozoology 25, 314–326. Figure 9 Actinosporean spore, exemplified by aurantiactinomyxon-type from Branchyiura sowerbi with extrusion point overlain by a prominent conical stopper with a microtubular coat, protruding through an opening in the shell valves. Bar ¼ 0.5 mm. Reproduced by permission of Urban & Fischer Verlag from Lom et al., 1997, Archiv fu¨r Protistenkunde 148, 173–189. Figure 10 Malacosporean-type spore of Buddenbrockia plumatellae with umbrella-shaped lid (l) overlain by a pad (arrow) containing tubules. The polar capsule lies superficially in its host cell, overlain only by the cell membrane. Bar ¼ 0.5 mm. Reproduced by permission of Cambridge University Press from Okamura et al. (2002), Parasitology 124, 215–223.

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Intracellular bodies of unknown composition and function have been described in myxosporeans variously as haplosporosomes or sporoplasmosomes, the latter present only in sporoplasms. In the majority of myxosporean and actinosporean stages these are fairly large cytoplasmic organelles, mostly greater than 150 nm in diameter and sometimes greater than 300 nm. Although many appear membrane-bound and of uniform density, others show an outer, denser cortical region around a less dense region and even a central dense core. Yet others appear as lucent rings around a dense core. These structures may be randomly distributed in the cytoplasm, or have a largely peripheral distribution in the sporoplasm (Desser and Brockley Patterson, 1978; Lom et al., 1982), or be inconspicuous or absent. Their function is unknown. Over 55 genera of Myxosporea have been named primarily on the basis of myxosporean spore structure (see Lom and Dykova´, 1992a and Kent et al., 2000b). However, variation in spore shape and position of polar capsules in species within genera sometimes makes the distinction between genera difficult, e.g., with Myxidium and Zschokkella. Furthermore, while spores of genera such as Henneguya and Myxobolus are distinguished morphologically (Henneguya spores possess long caudal processes which are absent from Myxobolus spores), these genera are widely distributed in phylogenetic trees based on analyses of 18S rDNA sequences (Smothers et al., 1994; Kent et al., 2001). It is likely that myxosporean-type spore characters are not wholly reliable for allocation of species to genera. 4.1.2.

Characters of the Actinosporean Phase

In contrast to the myxosporean phase of the life cycle, in which all nuclei are thought to be diploid (El-Matbouli et al., 1998), the actinosporean cycle of development includes a sexual phase leading to spore production. There are several excellent studies of these developmental cycles, including those culminating in triactinomyxontype spores (Lom and Dykova´, 1992b; El-Matbouli and Hoffmann, 1998), in raabeia-type and aurantiactinomyxon-type spores (Lom et al., 1997; Oumouna et al., 2002), and in sphaeractinomyxon-type


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spores (Hallett et al., 1998). Repeated proliferative cycles prior to, and concurrent with, sporogony, must be necessary for the long term infections and prolonged periods of spore release which are commonly observed (e.g., Gilbert and Granath, 2001). However, presporogenic stages have rarely been described. Recently, early stages of two actinosporeans have been described with proliferation through gametogony (Oumouna et al., 2002). The actinosporean cycle has been most fully described for Myxobolus cerebralis in Tubifex tubifex (see El-Matbouli and Hoffmann, 1998), which begins with proliferation in an intercellular position in the gut epithelium. Although the early stages have been described as multinucleate single cells, the illustrations suggest that there are several secondary cells within a primary cell, so that division is by endogeny not multiple fission. Sporogony in the same site occurs within pansporoblasts, which are formed by envelopment of two sporogonic cells by two pericytes. The enveloping cells become attenuated and divide to enlarge the pansporoblast. Each sporogonic cell undergoes meiosis and divides to give eight slightly anisogamous and gametes, which fuse with . Each of the eight zygotes (sporoblasts) divides into three valvogenic and three capsulogenic cells and a sporoplasm, and then matures into a triradially symmetrical spore, with three polar capsules, three valves and the sporoplasm in the form of a primary cell (endospore) containing several secondary cells (germ cells) (Figure 2). Abundant sporoplasmosomes have been reported in primary sporoplasm cells of, for example, aurantiactinomyxon-type spores (Lom and Dykova´, 1997), but not in their secondary (germ) cells and have not been specifically noted in M. cerebralis actinospores even in the primary sporoplasms. In most known actinosporean collective groups (previously considered as genera—see Kent et al., 1994), eight zygotes are formed giving eight spores in the pansporoblast. In two recently described actinosporean species, Tetraspora discoidea and Tetraspora rotundum, the pansporoblasts are tetrasporoblastic (Hallett and Lester, 1999) and it is likely that further variations will be revealed when more extensive studies of invertebrate hosts are carried out.

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The three valve cells which eventually form the spore wall become progressively thinner during spore maturation until little remains other than membrane doublets. The three valves fuse with each other to enclose the three capsulogenic cells except for apertures over the polar capsule apices. In most spore types the doublets extend posteriorly as pointed or globular caudal processes (absent from sphaeractinomyxon and tetractinomyxon types), which are folded when packed into the pansporoblast cavity. A layer of granular material may be secreted on to the outer surfaces of the doublets, giving strength to the valves especially over the anterior apertures and caudal processes (Lom et al., 1997). The mature polar capsules, formed via a capsule primordium and external tube, are plugged with a prominent stopper (Hallett et al., 1998), which may be covered with fibres or microtubules (Lom and Dykova´, 1997) (Figure 9). An especially interesting feature of actinosporean development, recorded several times, is the development of the sporoplasms separately in the pansporoblasts and their late incorporation within the valves (Lom et al., 1997; El-Matbouli and Hoffmann, 1998; Hallett et al., 1998). On release of the spores by passage through the gut or by death of the annelid host, the caudal processes inflate and probably serve to maintain the spores in the water at a level facilitating contact with the host fish. Spores of M. cerebralis have been shown to attach to the skin, allowing the sporoplasm to penetrate between cells of the epithelium and begin their division and migration to the cartilage (El-Matbouli et al., 1995).

5. DIAGNOSTIC CHARACTERS OF THE CLASS MALACOSPOREA The class comprises only one order—Malacovalvulida Canning, Curry, Feist, Longshaw and Okamura, 2000, one family— Saccosporidae Canning, Okamura and Curry, 1996, two genera— Buddenbrockia Schro¨der, 1910 and Tetracapsuloides Canning, Curry, Feist, Longshaw and Okamura, 2002, and two species— Buddenbrockia plumatellae Schro¨der, 1910 (syn. Tetracapsula


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bryozoides Canning, Okamura and Curry, 1996) and Tetracapsuloides bryosalmonae (Canning, Curry, Feist, Longshaw and Okamura, 1999) (syn. Tetracapsula bryosalmonae Canning, Curry, Feist, Longshaw and Okamura, 1999; Tetracapsula renicola Kent, Khattra, Hedrick and Devlin, 2000). Additionally, organisms clearly identifiable as malacosporeans have been described in pillar cells and endothelial cells of common carp, Cyprinus carpio, by Voronin (1993) and Voronin and Chernysheva (1993). The transfer of infection from bryozoans to salmonid fish has been successful for T. bryosalmonae (see Feist et al., 2001; Longshaw et al., 2002) but, so far, transmission between fish has been achieved only by inoculation of proliferative stages, and transmission between bryozoans or from fish to bryozoans has also not been achieved with certainty (see Section 5.3.7). Freshwater bryozoans (Phylum Bryozoa, Class Phylactolaemata) are the only known hosts for B. plumatellae but it is possible that fish host(s) exist for this species. Marine bryozoans may be involved in the life cycle of some marine myxozoans. There are five principal diagnostic characters of malacosporeans in the bryozoan hosts: (i) trophic phases are in the form of closed sacs or hollow ‘worms’ (vermiform stage), in the central cavity of which the spores lie free without envelopment by pericytes; (ii) spores have four capsulogenic and four valve cells surrounding two sporoplasms containing endogenous cells (Figure 3); (iii) the spore valve cells are not strengthened; (iv) openings of the polar capsules lie at the surface of the spore and are sealed by an umbrella-shaped cap or lid overlain by a pad with tubular or fibrous components (Figure 10); and (v) sporoplasms have highly characteristic, peripherally-distributed sporoplasmosomes which are dense, membrane-bound structures with a lucent invagination. In T. bryosalmonae, the species for which a fish host is known, the parasites occur extracellularly in a variety of tissues, including kidney interstitium, as single cells proliferating by endogeny, and in kidney tubule lumina, as spores with two polar capsules. In the proliferative phase in fish all primary cells have sporoplasmosomes characteristic of the class but sporoplasmosomes have not been described in the spores that develop in kidney tubule lumina.

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5.1. Buddenbrockia plumatellae Schro¨der, 1910 5.1.1.

History

The first record of B. plumatellae was that of Dumortier and van Beneden (1850), who observed worm-like creatures, about 0.1 mm long and filled with cells, moving in the body cavity of Alcyonella (now Plumatella) fungosa in Belgium. The ‘worms’ were subsequently studied in Plumatella repens and P. fungosa in Germany by Schro¨der (1910), who named the species. He first proposed that they were mesozoans but, after determining that they had four blocks of longitudinal muscle responsible for their movements, he reconsidered their status and suggested that they might be derived from nematodes (Schro¨der, 1912). They were seen at about the same time in Germany and Turkestan by Braem (1911), whose observations on Buddenbrockia in P. fungosa were in agreement on most points with those of Schro¨der (1910), although Braem proposed that Buddenbrockia was a trematode sporocyst. A wide host range and broad geographical distribution have subsequently been established by reports of Buddenbrockia from the bryozoans Stolella evelinae in Brazil (Marcus, 1941), Hyalinella punctata in Bulgaria (Grancarova, 1968), Austria (Wo¨ss, 2000) and Ohio, USA (Okamura et al., 2002a), Lophopodella carteri in Japan (Oda, 1972), Plumatella fungosa in Austria (Wo¨ss, 2000) and England (Canning et al., 2002), Plumatella repens in France (Okamura et al., 2002a), Plumatella sp. in Turkestan (Braem, 1911), and Fredericella sultana in Germany (Canning et al., 2002). Recently, studies of Buddenbrockia ‘worms’ have provided information on their ecology and pathogenicity in bryozoans and revealed their malacosporean nature through observations on fresh material (Morris et al., 2002) and their ultrastructure (Okamura et al., 2002a; Canning et al., 2002). Furthermore, 18S rDNA sequence analysis (Monteiro et al., 2002) has confirmed the assignment of B. plumatellae to the Malacosporea and revealed sequence identity with T. bryozoides which had been described previously in its sac-like form from Cristatella mucedo (Canning et al., 1996). T. bryozoides has subsequently been synonymized with


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B. plumatellae on the basis of both ultrastructural and sequence information (Canning et al., 2002). Sacs and ‘worms’ have not been found to co-exist in any host. This enigmatic parasite thus appears to be the ‘coelacanth’ of the myxozoan world, representing the ‘missing link’ in the evolution of myxozoans from worm-like bilaterian ancestors to the degenerate plasmodia and pseudoplasmodia of most present day myxozoans. Buddenbrockia also gives visual credence to the early molecular phylogenies which, almost unbelievably at the time, placed myxozoans as triploblast organisms close to the phylum Nematoda (Smothers et al., 1994; Schlegel et al., 1996).

5.1.2. Morphology and Development of the Vermiform Stage (a) Light microscopy. Publications on Buddenbrockia record it as worm-like, the smallest measuring 500 300 mm and the largest 3.6 0.1 mm (Figure 11). Irregular, club-shaped or branched specimens have been seen rarely (Morris et al., 2002). Most worms move freely in the coelomic cavity of their hosts but attached stages, presumed to be temporary, have been reported (Schro¨der, 1912; Canning et al., 2002; Morris et al., 2002). Schro¨der (1910, 1912) studied Buddenbrockia in P. repens and P. fungosa, first observing the stages which culminated in the formation of ‘embryos’ and later tracing the development of younger stages. The structures that he described as freely circulating in the coelom of the bryozoans can easily be equated with those seen by electron microscopy (Section 5.1.2.b). The youngest stage, like a ball of cells, consisted of an outer layer (Aussenzellen) around a solid core of cells. The core cells resolved into an inner layer (Innerzellen) around a central cavity. After organization of the inner cells as four longitudinal muscle blocks, a complete new inner cell layer was formed, which was linked to the muscle blocks by special cells with alveolar cytoplasm, the Z-lineage (Z-Linean). The new inner cell layer (which he called the oogonium layer) then fragmented, releasing ‘ova’ into the central cavity and these were converted through ‘blastula

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11

12

13

Figure 11 Part of the fully mature vermiform stage of Buddenbrockia plumatellae emerging from a living zooid of P. fungosa which has retracted its lophophore. Bar ¼ 100 mm. Figure 12 Ellipsoid sacs of spores of B. plumatellae (arrows) in coelomic cavity and within the tentacles of the lophophore of a living zooid of Cristatella mucedo. Bar ¼ 100 mm. Figure 13 A branch of a living colony of Fredericella sultana containing spherical sacs of Tetracapsuloides bryosalmonae. Bar ¼ 100 mm. Photographs provided by Ms Sylvie Tops.


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stages’ into ‘embryos’. The ‘embryos’ consisted of large polar cells, each containing a ‘lenticular capsule’ (linzenformige Kapsel ), and flattened cells surrounding two large and two small cells. As well as the ‘ova’ there were small rounded bodies 2.0–3.0 mm in diameter, often, but not always, with chromophilic granules. Schro¨der (1912) suggested that special disc-like cells with a tail at one end of the ‘worm’ might be spermatocytes, but he did not observe fertilization. Morris et al. (2002) observed the sudden appearance and growth of ‘elongate sacs’ in a culture of P. repens, which had been exposed to cell suspensions of brown trout kidney infected with T. bryosalmonae (PKX organism). After their initial attachment to the zooid, the sacs became free in the coelom, grew to 0.9–1.2 mm long and gave rise to masses of typical tetracapsulid spores within the sac wall, which was made up of two membranes. The spores were slightly flattened, with small projections from the base. There were two sporoplasms measuring 10 8.0 mm within four cytoplasmic valve cells, and there were four spherical polar capsules measuring 1.0 mm diameter at one pole. Spores were released into the bryozoan coelomic fluid, then disseminated throughout the colony before their release to the exterior between the tentacles of the retracted lophophore. The similarity between the elongate sacs full of tetracapsulid spores in P. repens and the vermiform B. plumatellae described by Schro¨der (1910) was noted by Morris et al. (2002). While recognizing that both parasites were malacosporean myxozoans, they considered that the species in P. repens differed from B. plumatellae. (b) Electron microscopy. The ultrastructure of the vermiform stage has been studied in the bryozoans H. punctata from Ohio, USA and P. repens from France (Okamura et al., 2002a). The same material of P. repens was studied more extensively, together with ‘worms’ from P. fungosa from England and Fredericella sp. (now identified as F. sultana) from Germany, by Canning et al. (2002). In all cases the ‘worms’ exhibited an outer and inner cell layer separated by a basal lamina and four muscle blocks but no gut or apparent nervous or excretory systems or special reproductive organs (Figure 14). The outer cells, termed mural cells (equivalent to

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m m

i

i m m

mc

14

15


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Schro¨der’s Aussenzellen), are flattened and linked by junctional complexes, consisting of an outer dense adherens junction continuous with an inner, sinuous gap junction. Similar bipartite junctions, which function to prevent ingress of surrounding fluids, have been found in other myxozoans and are typical of the phylum. The outer surface of the mural cells is raised into small papillae, projecting into the bryozoan coelom. In the worms collected from P. repens, the papillae were more numerous on the mural cells than they were in the ‘worms’ from the other hosts. All cells exhibit the usual eukaryotic organelles and vesicles, of which lipid globules with a peripheral pale ring and mitochondria with tubular cristae are notable. Immediately within the mural cells are four separate blocks of longitudinal muscle. In each muscle block, the individual myocytes meet at gap junctions. The sarcoplasm with nuclei, ribosomes and abundant mitochondria occupies the centre of each myocyte, while the contractile fibres are restricted to the surfaces facing the basal lamina and body cavity. The contractile fibres often appear to run obliquely to the longitudinal axis because the worms are twisted but may actually be longitudinal. As several of these cells abut to form the muscle bundle, the contractile elements are seen to enclose the entire bundle, so that the arrangement is circomyarian. No nerve connections are visible so contraction is likely to be myogenic. In very small ‘worms’ there are small spaces between the muscle blocks occupied by four sets of cells which run in presumed dorsal, ventral and lateral rows. These were termed primary type A cells by Okamura et al. (2002a). They are provided with abundant flat cisternae and expanded vesicles of endoplasmic reticulum and clearly correspond to Schro¨der’s (1912) alveolar Zell-Linean Z. Whereas the muscle blocks are separated from the mural cells by a fibrous basal Figure 14 Buddenbrockia plumatellae vermiform stage in Hyalinella punctata: transverse section of a young ‘worm’ showing mural cells (mc), muscle blocks (m) and inner cell layer (i) with primary type A cells (arrowheads). Bar ¼ 5 mm. Reproduced by permission of Cambridge University Press from Okamura et al. (2002), Parasitology 124, 215–223. Figure 15 B. plumatellae vermiform stage from Fredericella sultana. Section of part of a ‘worm’ with several nearly mature spores, within a wall of mural cells and muscle. Bar ¼ 5 mm. Reproduced by permission of the Society of Protozoologists from Canning et al. (2002), Journal of Eukaryotic Microbiology 49, 280–295.

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lamina, the primary type A cells are connected to the mural cells by prominent striated tubules. Schro¨der (1912) speculated that these four cell lines running from anterior to posterior between the muscles might be nerve or excretory cells. Rather than having a nervous function, Canning et al. (2002) proposed that the tubular connections of the primary type A cells with the mural cells might provide the mechanism for transport of nutrients absorbed from the bryozoan coelom across the basal lamina. Although an excretory role to remove metabolites from the body cavity of the ‘worm’ is conceivable, the role of secondary type A cells in secretion (see below) favours transport inwards across the tubules. The primary type A cells make contact with both the muscle block and the inner layer of cells. Outer arms extending from the type A cells connect with the muscle blocks by simple membrane apposition, while inner arms form complexes of adherens and gap junctions with adjacent cells (type B cells) of the inner layer. This time the adherens region is on the inner (body cavity) side of the ‘worm’, in an arrangement that would prevent fluids in the body cavity seeping outwards between cells. In the youngest ‘worms’, all cells of the inner layer, apart from the primary type A cells, form a complete sheet of type B cells internal to the muscle blocks. Type B cells lack the expanded endoplasmic reticulum and appear denser and more compact than the type A cells. During growth the number of muscle cells making up each block increases to at least 25 and secondary type A cells, formed by division of the primaries, become interpolated between the type B cells in the inner sheet of cells. After multiplication, the type A cells become grossly hypertrophied and the cytoplasmic vesicles become distended with fibro-granular secretions. Breakdown in organization of the inner cell layer releases the secretory cells into the body cavity, where they discharge the secretory products as spheres, which disperse to form a matrix filling the body cavity. There may be two types of secretory cells, as observed in parasites of H. punctata (see Canning et al., 2002; Okamura et al., 2002a). Some enlarged cells contain numerous vacuoles filled with concentric lamellae, which are discharged directly into the body cavity of the ‘worm’. Other secretory cells are


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distinguished by the release of their fibro-granular products first into the cell cytoplasm by breakdown of the endoplasmic recticulum cisternae. These cells are also characterized by the presence of a single large vacuole with a double wall. The function of this vacuole in processing the secretions for release into the body cavity is suggested by the presence of concentrations of the secretions retained by a fine membrane between the two walls of the large vacuoles and simultaneous depletion of the cytoplasmic areas of the fibro-granular products. How the membrane-bound products reach the surface of the cell for discharge as spheres has not been determined. Type B cells proliferate by binary fission and endogeny, piling up as several layers before separating and dispersing in the body cavity. One or more areas within the cytoplasm of these cells are isolated by a membrane from the rest of the cytoplasm and contain degenerate mitochondria and small vesicles. As these cells are destined to become spores, this is likely to be a mechanism for getting rid of spent organelles by exocytosis. Amongst groups of B-type cells were numerous anucleate, membrane-bound spheres measuring about 2 mm diameter. The cytoplasm of these anucleate spheres resembled that of B-type cells but contained only lipid and degenerate mitochondria, some exhibiting only ribosomes. These spheres probably represented the regions of cytoplasm expelled from the B-type cells and had disappeared by the time of spore maturation. The size of these spheres corresponds to the 2–3 mm eosinophilic structures seen by Schro¨der (1910) between the ‘ova’ in B. plumatellae, which he suggested might be degeneration products. Schro¨der similarly noted that these spheres had disappeared when the ‘embryos’ were formed. Some, if not all, B-type cells undergo meiosis, of which leptotene and zygotene within an intact nuclear envelope were observed (Canning et al., 2002). Division of these cells gives rise to compact groups, which mature into typical malacosporean spores with four valve cells overlapping four capsulogenic cells and enclosing two sporoplasms (Figures 3 and 15). These can be identified with the ‘embryos’ of Schro¨der (1910, 1912). The valve cells retain their cytoplasmic integrity and do not secrete hard walls. They become flattened, so much so that in places there

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is only a thin cytoplasmic layer with ribosomes between the two membranes, while elsewhere they are enlarged to accommodate nuclei, mitochondria and lipid globules. The valve cells form the spore wall by enclosing the sporoplasms and extending over the capsulogenic cells except at the four points of exit for the polar filaments. The polar capsules are spherical and their apertures are superficial (Figure 10). The capsular wall is composed of three layers. The outer dense membrane covers the whole capsule. The middle, lucent layer 130 nm thick, with radially arranged striae and the inner, homogeneous, moderately-dense layer, 20 nm thick, both stop short of the orifice for polar filament emergence. The precise arrangement of the polar filament has not been determined. Sections show it as a tubular structure, variously arranged as four to eight (rarely ten) triplets or S-shapes embedded in a dense matrix. At the point of exit the tubule expands to form a cup, plugged with dense material, which spills over the surface of the capsule as an umbrella-shaped cap. This is topped by a pad, containing parallel tubular or fibrous structures, overlain by the capsule membrane. This in turn is covered by the membrane of the capsulogenic cell. Microtubules surrounding capsular primordia and their external tubes were observed in Buddenbrockia in F. sultana. No external tubes were seen during capsulogenesis in the ‘worm’ from P. fungosa, either because an external tube is not formed or because the phase is transient. Even when the polar capsules are mature, the capsulogenic cells retain their nuclei and are well supplied with mitochondria. Almost always two sporoplasms, at least one with a visible endogenous cell, are present in spore sections. On one occasion three nucleated sections were visible. The presence of three sporoplasms in sections might indicate that division of sporoplasms occurred by binary fission rather than endogeny or that one of the endogenous cells had been released. However, release would necessitate division of the primary cell, then release of the secondary cell from its half of the primary cell and degeneration of the remains of the primary cell that enclosed it. More likely explanations are that there was an exceptional enclosure of three sporoplasms or that one section represents an amoeboid extension of a binucleate sporoplasm before


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enclosure of one of its nuclei to form a secondary cell. Schro¨der’s (1910) drawings of ‘embryos’ show enclosure of two large uninucleate cells, or two large binucleate cells or two large and two small uninucleate cells, suggestive of a developmental sequence within the ‘embryos’. A possible four-cell stage has not been confirmed by electron microscopy. The sporoplasms are abundantly supplied with lipid reserves, mitochondria and rough endoplasmic reticulum. The primary cells of mature sporoplasms are distinguished by peripherally-distributed, spherical sporoplasmosomes 150 nm in diameter, typical of malacosporeans. These dense structures are bounded by two membranes and contain a lucent bar-like invagination. The genesis of identical structures in T. bryosalmonae from Golgi cisternae and their possible functions are discussed in Section 5.3.4. Valve cells and capsulogenic cells are linked by typical junctional complexes, of which the adherens part is outermost. Points of contact between the two sporoplasms and between sporoplasms and valve or capsulogenic cells are made by simple membrane apposition. In fixed preparations spores are misshapen by the inward collapse of the valve cells and a variety of membrane condensations are evident in the spore cavity. Fresh spores are almost spherical, suggesting that in life their shape is determined by turgor pressure of fluid within the spore cavity. Small uninucleate dividing stages, seen in the surface epithelium of a zooid of P. fungosa infected with B. plumatellae, were considered as possible early stages derived from sporoplasms entering via the body wall (Canning et al., 2002). They were abundantly supplied with lipid globules and the nuclei were more indicative of myxozoans than bryozoans. Morris et al. (2002) reported an increase in the number of ‘elongate sacs’ of Buddenbrockia with time in laboratory-cultured P. repens, which had been collected from the field and then exposed to disrupted brown trout kidney infected with Tetracapsuloides bryosalmonae ( ¼ Tetracapsula bryosalmonae ¼ PKX organism). Subsequent analysis of 18S rDNA has demonstrated that B. plumatellae from the bryozoans P. fungosa, P. repens and H. punctata and T. bryozoides from the bryozoan Cristatella mucedo

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are synonyms (Monteiro et al., 2002) and thus that the Buddenbrockia could not have developed from T. bryosalmonae. More likely, the Buddenbrockia infection had already been established in the field. Natural infections of B. plumatellae, latent in the form of uninucleate stages in the body wall, as seen in P. fungosa, could have accounted for the increase in number of B. plumatellae sacs in the coelom of P. repens during host culture. (c) Timing of developmental sequence. The sequence of development from cryptic stages in the body wall through to mature ‘worms’ occurs rapidly. Canning et al. (2002) observed that P. fungosa exposed for a period in the field when an infection would have been acquired, and then subsequently maintained in laboratory culture for 17 days, developed at least 22 ‘worms’ in 19 zooids. These ‘worms’ had all developed over a period of six days. As colonies were not examined within the six-day period, the timing of development to ‘worms’ may be even more rapid than the data suggest. 5.1.3.

Morphology and Development of the Sac Stage

In contrast to the numerous bryozoan species reported as hosts for the vermiform stage of B. plumatellae, the rounded sacs originally described as Tetracapsula bryozoides by Canning et al. (1996) have been recognized only in Cristatella mucedo and, conversely, in spite of extensive research on the population biology of this host, no vermiform stage has ever been found in C. mucedo (summarized by Canning et al., 2002; also B. Okamura and S. Tops, personal observations). The sac-like stages are generally ellipsoid and measure 50–700 mm in their longest dimension (Okamura, 1996) (Figure 12). Some show constrictions indicative of budding (Okamura, 1996) and there are also club-shaped and branching forms (S. Tops and B. Okamura, personal observations). Similar odd shapes were observed in the Buddenbrockia-like parasites by Morris et al. (2002). Sacs are a simplified form of the vermiform stages, lacking the blocks of longitudinal muscle and, in consequence, also lacking the organization of type A cells into identifiable rows. The mural cells divide to provide for growth of the sac. Any dying cells in this layer


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are removed to the exterior by undergrowth of adjacent cells, thus preventing breaches in the wall. The mural cells are linked to one another by the usual adherens/gap junctional complexes and exhibit nuclei with large nucleoli, abundant mitochondria with tubular cristae, and lipid reserves. The inner layer of cells was not complete in any of the sacs examined by Canning et al. (1996). Early in the growth of the sacs, some of the cells attached to the wall begin their proliferation, forming one or more inward bulges stretching across and dividing the cavity into regions (Figure 16). These are type B cells, destined to become spores. Adjacent to the basal lamina, filose projections of these cells are linked by typical junctional complexes, with the adherens portion innermost. Further away from the wall the filose connections are lost and eventually the cells form a population of free, rounded cells measuring about 2.5–15.0 mm, with large nuclei and prominent nucleoli. The smaller cells almost certainly arise by budding, as there was little evidence of proliferation by endogeny and there were occasional cytoplasmic bridges between large and small cells. As in the vermiform stage, the large type B cells were able to sequester spent organelles in endogenously-formed vacuoles for disposal to the exterior in cytoplasmic fragments measuring about 1.0 mm in diameter, a large number of which were interspersed between true cells in the cavity of the sac. These later disappeared. A few isolated cells with expanded endoplasmic reticulum containing homogeneous secretions remained attached to the basal lamina. These could have been type A cells but, in the absence of muscle blocks and tubules linking them to mural cells, their orientation could not be determined. Spore formation is by division of the large type B cells (Figure 17) into ten cells: four valve cells enclosing two larger cells, which are destined to become sporoplasms, and almost completely overlapping four capsulogenic cells. In the original material meiosis was not observed but it has been seen in recently studied material (S. Tops and B. Okamura, unpublished observations). External tubes have not been seen during capsulogenesis. The mature spores were exactly as in the vermiform stages (Figure 20), as were the polar capsules (Figure 21).

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mc

17 16

19 18

c v

s

ss

s

v v

20

21


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5.1.4. Ecology, Pathogenicity and Behaviour Most of the studies on B. plumatellae simply record the presence of the vermiform stage in a variety of phylactolaemate bryozoans (Section 5.1.1) and discuss its phylogenetic position. There are practically no data on prevalence but it is clear that the vermiform stage can be very difficult to find. This is due in part to very localized distribution within a larger population of hosts. For example, B. plumatellae was restricted to H. punctata in two very specific areas in Cowan Lake, Ohio and was not found in other bryozoan species in the same lake (see Canning et al., 2002). If its distribution is restricted within a larger host population, this implies that an alternate host, if one exists, is also restricted. Canning et al. (2002) found that at least 22 ‘worms’ infecting 19 zooids had developed in P. fungosa within six days. Such rapid growth, followed by breakdown and expulsion of mature ‘worms’ from bryozoans (Canning et al., 2002; Morris et al., 2002) and slow maturation of possible cryptic stages would also account for its apparent rarity, and its real prevalence may be much higher. The sac stage has been found only in C. mucedo. Populations of this host have been studied extensively (see Okamura et al., 2002b for a review) and the free-floating sacs have been found many times (Okamura, 1996). The prevalence of sac stages in bryozoan populations varies both temporally and spatially. In one site, 10– 49% of colonies of C. mucedo were observed to be infected over the summer period, while prevalence levels ranged from 0–36% at other

Figures 16–21 Buddenbrockia plumatellae sac stage in Cristatella mucedo. Figure 16 Young sac showing mural cells (mc) and compact growth of inner cells as bulges dividing the central cavity. A smaller stage (arrow) is an almost compact ball of cells before expansion of the cavity. Bar ¼ 10 mm. Electron micrograph provided by Dr A. Curry. Figures 17, 18, 19 Sporogonic cells from a semi-mature sac, showing successive divisions to form the cells of the incipient spore. Bar (on Figure 16) ¼ 1.7 mm, 1.9 mm and 3.4 mm, respectively. Electron micrographs provided by Dr A. Curry. Figure 20 Mature spore showing two sporoplasms (s), one with a secondary cell (ss) within the attenuated valve cells (v), and a capsulogenic cell (c). Note abundance of dense sporoplasmosomes. Bar (on Figure 16) ¼ 2.5 mm. Figure 21 Polar capsule in a capsulogenic cell. Note the lucent capsule wall with radial striae and the tubule-containing pad overlying the apex of the capsule (arrow). Bar (on Figure 16) ¼ 0.5 mm. Figures 20 and 21 are reproduced with permission of the Institute of Parasitology, Academy of Sciences of the Czech Republic, from Canning et al. (1996), Folia Parasitologica 43, 249–261.

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sites (Okamura, 1996). Attempts to find an alternate host should be guided by data on the prevalence and seasonality in bryozoan hosts. Schro¨der (1910, 1912) and Braem (1911) studied B. plumatellae in fixed material and did not observe living ‘worms’. In the recently reported cases of infection with the vermiform stage, the ‘worms’ were undergoing sinuous, writhing movements in the coelom of their hosts (Canning et al., 2002; Morris et al., 2002; Okamura et al., 2002a). Such writhing does not result in noticeable translocation within the colonies, but ‘worms’ have been observed to work their way slowly out of their degenerating bryozoan hosts, presumably through the vestibular pore at the base of the tentacles (Canning et al., 2002). Once released from the bryozoan, the ‘worms’ were seen to alternate between curling into a tight corkscrew and straightening, for as much as 80 min before death. ‘Worms’ artificially expelled from colonies through external pressure showed similar behaviour. Some effects of Buddenbrockia infection on the bryozoan hosts have been documented (Canning et al., 2002). Hosts respond to infections of worm-like stages of B. plumatellae by inward pinching of the peritoneum from the outer body wall between zooids and isolating the infection. Even so, the overall effect on colony health may be detrimental. In an experiment where infected and uninfected colonies of H. punctata were monitored in their natural environment after attachment to plastic surfaces, the infected colonies grew at a slower rate than uninfected ones and did not produce statoblasts. Statoblasts were also not produced in laboratory-cultured infected P. fungosa, although they were produced during the same period in uninfected P. fungosa. In addition, infected H. punctata and P. fungosa maintained in laboratory culture conditions quickly regressed and died. Exit of mature ‘worms’ from the base of the lophophore was observed to damage the retraction mechanism of the lophophores (Canning et al., 2002). Sac stages of B. plumatellae have been shown to compromise statoblast but not larval production in a population of C. mucedo (Okamura, 1996). Infected colonies are malformed through swelling and show generally slower lophophoral retraction in response to


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prodding. Colonies with heavy infections are filled with parasitic sacs with most zooids being degenerate (Okamura, 1996). Sacs can apparently be expelled from colonies since free sacs are often found near colonies held overnight in containers (B. Okamura, personal observation). Presumably this occurs by means of the same mechanism as release of statoblasts from the vestibular pore. The detrimental effect of B. plumatellae is believed to have contributed to a notable population ‘crash’ of C. mucedo at one site (Vernon et al., 1996). If exit of mature ‘worms’, or of spores released from them, occurs through the statoblast pore at the base of the lophophore it may be advantageous for the ‘worms’ to maintain a position close to the lophophore. In bryozoans which have narrow branches and can isolate uninfected regions by inward pinching of the peritoneum, ‘worms’ may avoid retention in degenerating regions by movement towards newly formed zooids. Thus, the muscular system to facilitate this movement may be essential. In C. mucedo, which has a voluminous coelomic cavity connecting with all zooids, a more rounded form of the parasite, capable of budding and increasing in numbers, may be advantageous. These non-motile stages would have more unrestricted access to the zooids, many becoming trapped near the base of the lophophore.

5.2. Malacosporean Stages in Carp, Cyprinus carpio Previously unobserved intracellular parasites very similar to the PKX organism (see Tetracapsuloides bryosalmonae, Section 5.3) were found in the pillar cells of secondary gill lamellae (Voronin and Chernysheva, 1993) and in endothelial cells of blood vessels of gills, kidney and brain (Voronin, 1993) of common carp. In both cases the parasites were detected in young carp under two years of age that had shown signs of swim bladder inflammation due to Sphaerospora renicola but had also exhibited pale gills and suffered anaemia, which are not typical of S. renicola infections. The new parasites were single cells undergoing endogeny in vacuoles in the host cells. The primary cells showed a peripheral

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distribution of typical malacosporean sporoplasmosomes (referred to as haplosporosomes, as designated by Seagrave et al., 1980). Wellknown stages of S. renicola were also present in the fish, associated with haemorrhagic inflammation of the swim bladder but these stages do not develop sporoplasmosomes. At the time that the parasites were observed in carp, the PKX organism had not been identified or described as a malacosporean species (see Anderson et al., 1999b; Canning et al., 1999b, 2000) and Voronin (1993) suggested that the new parasite might be a PKX-like stage of S. renicola, responsible for the acute anaemia and mortality. S. renicola is now known to be a myxosporean myxozoan, unrelated to the PKX organism or to B. plumatellae. The discovery of these organisms in carp has particular importance in the light of the demonstration that B. plumatellae infects a range of freshwater bryozoans and is a malacosporean myxozoan for which a vertebrate host is as yet unknown.

5.3. Tetracapsuloides bryosalmonae (Canning, Curry, Feist, Longshaw and Okamura, 1999), the PKX organism 5.3.1.

Taxonomic History

Proliferative kidney disease (PKD), characterized by abdominal swelling and pale, anaemic gills, was recognized as a serious disease of salmonid fish from the early part of the 19th century. The abdominal swelling is caused by an intense immune response to the presence of unicellular organisms and involves macrophages, neutrophils, plasma cells and lymphocytes infiltrating into the interstitial tissues of viscera, particularly kidney and spleen. The enigmatic organisms were first named PKX, the unknown organism ‘X’ responsible for PKD (Seagrave et al., 1980). The myxozoan nature of these organisms was determined by Kent and Hedrick (1985a), when they observed spores with polar capsules in infected kidneys. The incrimination of annelids in the life cycle of M. cerebralis by Markiw and Wolf (1983) led to searches for alternate hosts for PKX but these were fruitless


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until molecular data indicated the involvement of bryozoans (Anderson et al., 1999a,b). Anderson et al. (1999a,b) obtained 18S rDNA sequences from saclike organisms resembling T. bryozoides (now B. plumatellae, see Section 5.1.3), parasitizing the bryozoans Cristatella mucedo, Pectinatella magnifica and Plumatella rugosa, collected in Ohio and Michigan, USA. They next determined that three PKX sequences from trout in England and two sequences from trout in Washington State, USA, varied by 0.5–1.1% in comparison with a standard sequence U70623 from France (Saulnier and de Kinkelin, 1997). Phylogenetic analyses of sequences from trout and bryozoans showed that one sequence from C. mucedo and two from P. magnifica fell within this range (0.5% different from U70623) and that a clade of six other sequences from the bryozoans differed by only 1.9–2.4%. When a phylogenetic tree was constructed using these PKX-like sequences, together with sequences of T. bryozoides and other myxozoan genera, there were two major branches, with T. bryozoides and all PKX-like sequences diverging from the others. This work, by Anderson et al. (1999a,b), was a major advance, not only demonstrating that bryozoans were the source of infection of PKX to salmonid fish but also leading to the description of the bryozoan phase of PKX and its naming as Tetracapsula bryosalmonae by Canning et al. (1999b). It further led to the establishment of the new myxozoan class Malacosporea embracing T. bryozoides and T. bryosalmonae (see Canning et al., 2000) and heralded a new era of research on PKD, especially on its transmission (Feist et al., 2001; Gay et al., 2001). When the species name Tetracapsula bryozoides was found to be a synonym of Buddenbrockia plumatellae (see Monteiro et al., 2002) a new generic name, Tetracapsuloides, was proposed for Tetracapsula bryosalmonae. It was deemed that there were sufficient differences between it and B. plumatellae, including 19.4–21% difference in the 18S rDNA sequence (Anderson et al., 1999b), to warrant the establishment of a new genus. The previously given names Tetracapsula bryosalmonae Canning, Curry, Feist, Longshaw and Okamura, 1999 and Tetracapsula renicola Kent, Khattra, Hedrick

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and Devlin, 2000 became synonyms of Tetracapsuloides bryosalmonae (see Canning et al., 2002). A comprehensive review of PKD and of the PKX organism responsible for the disease was presented by Hedrick et al. (1993). Pertinent data are summarized in the present review and the more recent investigations on the malacosporean connection are dealt with more fully. 5.3.2.

Hosts

Pike, Esox lucius (Family Esocidae) (Seagrave et al., 1981) and salmonid fish are known as vertebrate hosts of T. bryosalmonae and are susceptible to a greater or lesser extent. Although there has been some taxonomic disagreement, it is widely believed that esociforms, salmonids and osmerids are closely related groups (see Berra, 2001 for discussion). Nelson (1994) considered pike (Esocidae) to be the primitive sister group to the osmerids and salmonids. T. bryosalmonae has also been reported from phylactolaemate Bryozoa, including Pectinatella magnifica and Plumatella rugosa (see Anderson et al., 1999a,b), Fredericella sultana (Figure 13) (Longshaw et al., 1999; Gay et al., 2001), Plumatella emarginata (see Okamura and Wood, 2002), and probably also Cristatello mucedo (see Anderson et al., 1999a,b). The most common bryozoans associated with sites of PKD outbreaks are F. sultana, F. indica and P. emarginata (see Okamura and Wood, 2002). 5.3.3.

Geographical Distribution

At present PKD appears to be restricted to the northern hemisphere, where it has been reported from many European countries, the USA and Canada. Most reports of PKD from North America concern the western states (Hedrick and Kent, 1986) but cases have been found in Newfoundland (Brown et al., 1991). The most serious effects are apparent when fish are overcrowded in fish farms and hatcheries fed by river water and not spring water (Hedrick and Kent, 1986). Wild populations of brown trout, Atlantic salmon, arctic char, pike and


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grayling have been shown to be infected in Europe (Seagrave et al., 1981; Wootten and McVicar, 1982; Bucke et al., 1991). Infected feral populations of rainbow and cutthroat trout have been found in Utah and Montana in the intermontane region of the USA (Smith et al., 1984; MacConnell and Peterson, 1992). Infected colonies of the bryozoans P. magnifica, P. rugosa and C. mucedo were found in Ohio and Michigan, midwestern regions of the USA, where PKD has not been reported. The lack of salmonids in midwestern sites where infected bryozoans were encountered suggests that the distribution of T. bryosalmonae is not influenced by the distribution of salmonids. The broad distribution of bryozoans and the apparent lack of host specificity of T. bryosalmonae suggest that T. bryosalmonae is likely to infect bryozoans across a broad geographical range. The lack of detection of infected bryozoans in many regions probably reflects the generally low prevelance of T. bryosalmonae in bryozoans and the small amount of research done on freshwater bryozoans. The apparent lack of PKD in salmonid farms in the southern hemisphere suggests that the parasite may be restricted to the northern hemisphere. 5.3.4. Pathogenicity and Development in Fish Proliferative kidney disease (PKD) in salmonid fish is a seasonal disease, occurring from early to late summer when water temperatures rise. It is basically a disease condition caused by a severe inflammatory response to the presence of T. bryosalmonae trophozoites in many body tissues, especially kidney and spleen. In the early stages of infection trophozoites are widespread but notably are present as free parasites in small blood vessels and as extracellular stages in kidney interstitium, where they evoke an immune response (Feist, 1997). Later they enter renal tubule epithelia, also as proliferating trophozoites, and finally emerge into the tubule lumina where presporogonic pseudoplasmodia and spores are formed (Kent and Hedrick, 1985a, 1986; Kent et al., 2000a). Fish surviving the primary attack are able to eliminate the parasites and are resistant to further infections. Kent and Hedrick (1986)

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traced the development of infections in kidney of rainbow trout, Salmo gairdneri, after exposure to infection in a river hatchery followed by laboratory maintenance at 15–18 C. Trophozoites were first detected at three weeks and reached peak prevalence at seven weeks, when intraluminal stages in renal tubules were first seen. Nephritis was observed at five weeks, peaked at the 11th week and had regressed by the 21st week. Intraluminal spores persisted after disease remission. The trophozoites (extrasporogonic stages) in the interstitium range from single cells measuring about 5 mm diameter to primary cells containing several secondary and tertiary cells (Figure 22). The primary cells measure up to 25 mm in diameter, depending on the number of secondary cells within them. The primary cells are characterized by typical malacosporean Golgi-synthesized spherical sporoplasmosomes (Figure 23) measuring about 150 nm in diameter, which migrate from a general cytoplasmic distribution to a position beneath the plasma membrane which is overlain by an electron dense coat (Feist, 1997). Of the two membranes surrounding the sporoplasmosomes, the inner is reflected inwards as the lining of a lucent, bar-like invagination, which at its base may divide into four lobes (Morris et al., 2000a). The sporoplasmosomes adopt a peripheral position, aligned so that the opening of the bar-like invagination faces the plasma membrane, with which they fuse. The electron dense material on the plasma membrane may be sporoplasmosome-derived. Lipid globules, mitochondria and vesicles containing membranous debris (possibly spent organelles) are also present in these cells. Secondary and tertiary cells, which lie in closefitting vacuoles in the mother cell, show increasing density of cytoplasm and nuclei but lack sporoplasmosomes. Breakdown of primary cells releases secondary cells, which then become primary cells with sporoplasmosomes. The electron dense covering of the trophozoites may initially protect against the host’s primary immune responses, which may be antibody- and phagocyte-based (MacConnell et al., 1989), but dense secretions released from the sporoplasmosomes may be involved in primary cell membrane dissolution. Release of multiple antigens at the time


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1 3

2 n

n

22

23

Figure 22 Endogeny of Tetracapsuloides bryosalmonae (PKX organism) from kidney interstitium of rainbow trout Oncorhynchus mykiss showing the primary (1), secondary (2) and tertiary (3) cells and nuclei (n). Only the primary cell has sporoplasmosomes (arrows). Bar ¼ 1.0 mm. Electron micrograph provided by Dr S.W. Feist. Figure 23 Sporoplasmosomes generated by Golgi cisternae in a primary cell of Tetracapsuloides bryosalmonae (PKX organism). Bar ¼ 0.2 mm. Reproduced by permission of the European Association of Fish Pathologists from Feist (1997), Bulletin of the European Association of Fish Pathologists 17, 209–214.

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of primary cell breakdown and secondary cell release may induce the cell-mediated response, which eventually neutralizes the infection. Lymphocytes and plasma cells become prominent in the inflammatory tissue, while attachment of neutrophils to parasites is followed by engulfment of trophozoites by macrophages. The lymphokine mediators of the T cell response have not been identified. The inflammatory response ends with granuloma formation. The kidney interstitium becomes massively infiltrated leading to necrosis and disorganization of the renal tubules. External signs shown by heavily infected fish are abdominal distention due to enlargement of spleen and kidney and accumulation of ascitic fluid, pale gills indicative of anaemia due to destruction of haemopoietic tissue in the kidney, bilateral exophthalmia and occasional darkening of the skin (melanosis). Mortality may approach 100% in severe outbreaks but is usually caused by secondary infections. As infection advances the same kinds of trophic stages are found in the renal tubule epithelia, where proliferation by endogeny continues (Kent and Hedrick, 1986). Secondary cells escape to the tubule lumina and again undergo endogeny. Finally, cells of the intraluminal series become sporogonic pseudoplasmodia, each giving rise to a single spore (monosporous pseudoplasmodia). The spores measure 12.0 7.0 mm and are composed of two capsulogenic cells forming spherical polar capsules, 2.0 mm in diameter, and a sporoplasm partially surrounded by two unstrengthened valve cells (Kent and Hedrick, 1986). Further data were provided by Kent et al. (2000a), who reported that there are four coils of the polar filament and that the sporoplasm is surrounded by refractile granules and enveloped by the pericyte without evidence of intervening valve cells. Sporoplasmosomes, typical of the class Malacosporea, were not present in the sporoplasms in the spores in fish kidney. The absence of sporoplasmosomes and lack of valve cells suggest that the observed spores were not mature. This is in accordance with previous thinking that salmonids are not satisfactory hosts for T. bryosalmonae.


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5.3.5. Morphology and Development in Bryozoa Free-floating sacs in the coelom of C. mucedo, P. magnifica and P. rugosa collected from Ohio and Michigan, USA, were identified by their rDNA sequences (Anderson et al., 1999a,b) in comparison with a sequence obtained by Saulnier and de Kinkelin (1997) from PKX organisms from rainbow trout. Development of sacs and spore production have been studied in Fredericella sultana and a plumatellid that was later confirmed to be Plumatella emarginata (Canning et al., 1999b, 2000; Okamura and Wood, 2002), after similar rDNA identification of the sac stages by Longshaw et al. (1999). Sacs have subsequently been seen in F. sultana in France and Washington (Gay et al., 2001; B. Okamura, unpublished observations). Sacs are spherical (Figure 13), measuring up to 350 mm in diameter. The wall is formed of a single layer of flattened mural cells linked by complexes of adherens and gap junctions. The earliest stages observed were in F. sultana and already had a population of cells filling the central cavity without a defined inner layer of cells. The cells filling the cavity did not form as a compact bulge as in B. plumatellae (Section 5.1.3). The free cells, which were all linked by filose processes, at first appeared of similar density, although some were slightly less dense and larger than others (Figure 24). They quickly become differentiated as large pale cells, surrounded by smaller, stellate cells (Figure 25). These clusters become the units from which the spores are formed but a few cells flattened against the wall probably serve to increase the population in the central cavity. The large pale cells are destined to become sporoplasms. They undergo meiosis and, although cytoplasmic fission was not seen, there are ultimately two sporoplasms, each with an endogenously-formed secondary cell, accounting for the four haploid nuclei derived from meiosis. The stellate cells differentiate into four attenuated valve cells enclosing the sporoplasms and four capsulogenic cells at one pole (Figure 26). Immature and mature spores were seen in P. emarginata. The valve cells overlap the capsulogenic cells except at the points of exit of the polar filaments from the polar capsules. The polar capsules are similar to those of B. plumatellae with the orifice for the filament

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sp

m

sp

24

25

c

c v

v s

s

c

pf

ss 26

27

l

28


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overlain by an umbrella-shaped cap and a pad enclosing tubular structures. External tubes were not observed during capsulogenesis. The capsule wall differs from that of B. plumatellae in lacking both an inner layer distinct from the lucent layer and lacking the radial striae in the lucent layer which are present in B. plumatellae (Section 5.1.2.b) (Figures 27 and 28). A few peripherally-distributed sporoplasmosomes, typical of malacosporeans, were present in some sporoplasms but far fewer than in B. plumatellae. It is possible that none of the spores in the available material was completely mature and that the final structure of the polar capsule may change and the numbers of sporoplasmosomes may become as great as in the proliferative cells. Other features of the spores, such as mitochondria, lipids and cell junctions, were indistinguishable from those of B. plumatellae. The rapid development of ‘worm’ stages of B. plumatellae suggested that T. bryosalmonae may also develop rapidly from cryptic stages, and this has recently been demonstrated by the proliferation of sacs within three days in colonies collected from the field and maintained in laboratory culture (S. Tops and B. Okamura, unpublished observations). The probability of rapid development is supported by the generally low prevalence of T. bryosalmonae in bryozoan populations, and the brevity of periods in which prevalences are high (Section 5.3.6).

Figures 24–28 Tetracapsuloides bryosalmonae in Plumatella emarginata (Figures 24, 26, 27, 28) and Fredericella sultana (Figure 25). Figure 24 Young sac with sporoplasmogenic cells (sp) and stellate cells (arrows) dispersed within a wall of narrow mural cells (m). Bar ¼ 10 mm. Electron micrograph provided by Dr A. Curry. Figure 25 Sporoplasmogenic cell (sp) with three attached stellate cells (arrows). Bar ¼ 5 mm. Figure 26 Spore showing three of the four capsulogenic cells (c) (capsules not visible) and overlapping valve cells (v) forming a narrow wall enclosing two sporoplasms (s), one with a secondary cell (ss). Bar ¼ 2 mm. Figure 27 Polar capsule in a capsulogenic cell (c). The wall, consisting of lucent and moderately dense layers enclosed by a membrane, surrounds the dense matrix with the polar filament (pf). Note that the lucent layer of the wall has no radial striae. Bar ¼ 0.5 mm. Figure 28 The extrusion point for the polar filament, which is continuous with an umbrella shaped lid (l) overlain by a pad containing tubular structures (arrow). Bar ¼ 0.9 mm. Figures 26 and 28 reproduced by permission of the Society of Protozoologists from Canning et al. (2000), Journal of Eukaryotic Microbiology 47, 456–468.

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5.3.6.


Prevalence of T. bryosalmonae in Bryozoa

Based on visual inspection of bryozoans for sacs, prevalence of T. bryosalmonae in populations is generally very low and shows spatial and temporal variation. For instance, in 1998 six of 30 colonies of P. magnifica (20%) were infected at one site and two of 30 colonies of C. mucedo (6.7%) were infected at another site (Anderson et al., 1999b). In the following year, no infections were found in colonies at either site or at six other sites in the region, despite searching through many thousands of colonies (Okamura et al., 2001). Subsequent screening of thousands of bryozoans collected from sites associated with PKD outbreaks in California, Oregon and Washington State, USA, France, Italy, Germany, Switzerland and the UK have similarly failed to reveal infections (Okamura and Wood, 2002; Tops and Okamura, in press) apart from one colony from a site in France (Gay et al., 2001) and one colony from a site in Washington State, USA (B. Okamura, unpublished observations). Infections therefore appear to be covert for most of the year. June and July were the only months in southern England when abundant sacs of spores were found with up to 15 sacs per zooid, although, even at this time, the infections were highly patchy (S. Tops and B. Okamura, unpublished observations). After this, infections apparently disappear and sacs are not discernible, although transmission studies indicate that infective spores are present throughout the year (Section 5.3.7). 5.3.7.

Transmission of T. bryosalmonae

Before the discovery that bryozoans release spores, which are infective to fish in the life cycle of T. bryosalmonae, the only known procedures for infecting fish were intraperitoneal inoculation of homogenates of infected salmonid kidney, spleen or blood (CliftonHadley et al., 1984; Kent and Hedrick, 1985b), and exposure of naive salmonids to water bodies endemic for PKD, provided that the water temperature for maintenance of the fish was not below 12–13 C (see Gay et al., 2001). The size of the infectious agent was


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determined as 20–25 mm by passing water, from a site endemic for PKD, through a series of filters (P. de Kinkelin, unpublished data quoted by Hedrick et al., 1993). These data correlate with the seasonal recurrence of PKD, which occurs only when water temperatures rise to about 15 C. This happens in early summer, coinciding with the growth of new bryozoan colonies from statoblasts or, in the case of F. sultana, new growth from overwintering colonies. The first experimental transmissions to rainbow trout, using T. bryosalmonae sacs dissected from F. sultana colonies, were reported by Feist et al. (2001). In their experiments rainbow trout were exposed, for 90 min in still water, to disrupted F. sultana colony branches and to T. bryosalmonae sacs dissected from F. sultana before maintenance of the trout in flowing water. Separate groups of fish in this experiment were exposed to (i) F. sultana colonies on willow roots from the infected site in flow through water throughout the 12 week experiment, (ii) disrupted T. bryosalmonae sacs administered by intraperitoneal injection, and (iii) supernatant from kidney homogenates from clinically infected rainbow trout used as positive controls. Negative controls received inoculations of sterile phosphate buffered saline only. T. bryosalmonae was detected by light microscopy and/or the polymerase chain rection (PCR) in the groups of fish exposed to spore-containing water. Intraperitoneal inoculation was successful only with kidney homogenates, indicating that sporoplasms require a natural route of infection, whereas trophozoites are partially adapted to fish tissue responses and can survive inoculation into the peritoneal cavity. Further insight into the epidemiology was provided by the experimental transmission studies of Gay et al. (2001), in which uninfected rainbow trout were introduced into known PKD infection sites at monthly intervals throughout the year and maintained there for several days on each occasion. Some introductions at times when water temperatures were nonpermissive for development of PKD were included. Fish in all groups developed infections on transfer to 16 C in the laboratory, although initial exposure temperatures had ranged from 9 C to 18.9 C, demonstrating that spores were present throughout the year

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even at the lowest temperatures. One site was a recycling water system (Chilmonczyk et al., 1989) in which P. repens was present (Gay et al., 2001; Okamura and Wood, 2002). The other was a river in which F. sultana was abundant. Confirmation that F. sultana colonies from the river site could carry infections throughout the colder months was obtained by PCR amplification of T. bryosalmonae sequences from the colonies collected regularly from October to May. Finally, rainbow trout co-habited with these F. sultana colonies in an aquarium developed PKD when kept at 16 C. The presence of T. bryosalmonae was detected in the bryozoans by PCR and by infectivity to fish. Morris et al. (2000b) demonstrated, by means of hybridization in situ, that T. bryosalmonae was present in the gill arch of rainbow trout three days after exposure of the fish in a site enzootic for PKD. In this first step towards elucidating the early development of T. bryosalmonae in fish, they suggested that the gill acts as a portal of entry. As proliferative stages abound in blood, it is reasonable to propose that the sporoplasms or their progeny enter blood vessels in the gills to be transported to other organs in the body. More recently, Longshaw et al. (2002) extended these results by using the hybridization in situ technique on rainbow trout exposed in the laboratory to F. sultana infected with T. bryosalmonae. Sporoplasms were detected entering the skin through mucous cells within the first minute of exposure and stages were found in skeletal muscle up to 72 h after exposure. These fish later developed PKD, demonstrating that the parasites had completed their migration to the viscera. So far transmission from fish to Bryozoa has not been achieved with certainty, although possible success has been reported by Morris et al. (2002). In their experiments, colonies of P. repens were maintained with brown trout infected with T. bryosalmonae or with disrupted infected kidneys of brown trout or rainbow trout. The bryozoans were examined by PCR after exposure periods of four and eight weeks. All gave negative results for T. bryosalmonae except two colonies exposed to disrupted brown trout kidney, which gave positive PCR results. Morphological evidence was


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not obtained and the PCR results could have resulted from uptake of water containing T. bryosalmonae stages released from the disrupted kidney. Other attempts to infect bryozoans from infections in fish have been unsuccessful (B. Okamura and S. Tops, unpublished observations). Two obstacles to fish-to-bryozoan transmission experiments are that the spores, which develop in the lumen of kidney tubules, are present in small numbers only, except in arctic char where they are reported to be numerous (Kent et al., 2000a), and that bryozoans may be difficult to maintain in laboratory culture for long enough for infections to become overt. If the putative stages seen in the epithelium of P. fungosa by Canning et al. (2002) were indeed stages of B. plumatellae, similar stages may exist in bryozoans infected with T. bryosalmonae and a stimulus, as yet unknown, may be required to trigger entry to the coelom and to promote their development into sacs. Once triggered, they may develop very quickly. Hybridization in situ, as used by Morris et al. (2000b), would be a convenient way to track the development and migration of such latent stages after exposure to spores from fish. A further consideration is whether T. bryosalmonae infections can pass between bryozoans without requiring a fish host. T. bryosalmonae was first discovered in bryozoans from three lakes in the USA in which salmonid fish were not known to be present (Anderson et al., 1999b; Okamura et al., 2001). Thus, either salmonids were present but not reported or some non-salmonid fish was present in the lakes that could act as the alternate host. Otherwise, fish could not be obligatory in the life cycle. Pike, Esox lucius, have been reported as hosts of T. bryosalmonae by Seagrave et al. (1981), but pike were also absent from at least one of the North American lakes (Cowan Lake) where bryozoans were infected (Okamura et al., 2001). The PKX-like stages in pike detected by Seagrave et al. (1981) were found at a time when molecular identification was not practised and the parasites might have belonged to another species of malacosporean. PKX-like organisms have been found in carp (Voronin and Chernysheva, 1993) and others may yet be discovered.

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5.3.8.


Differentiation of the Sac-like Stages of B. plumatellae and T. bryosalmonae

Apart from a sequence difference of about 20% in the 18S rDNA (Anderson et al., 1999a, b) the two malacosporean genera can be differentiated morphologically as follows: (i) Sacs of T. bryosalmonae are spherical, measuring up to about 250 mm (Figure 13). Division has not been observed and may not occur. Those of B. plumatellae are ellipsoid (Figure 12), elongate, irregular or constricted and divide by fission or budding. Sacs may attain 700 mm in length. (ii) Early proliferation of T. bryosalmonae gives rise to a population of cells filling the sac with only a few cells adherent to the mural cells (Figure 24). In early sacs of B. plumatellae, proliferation gives rise to one or more compact inward bulges separating the cavity into sections (Figure 16). (iii) Inner cells of T. bryosalmonae are differentiated as pale sporoplasmogenic cells and dense stellate cells (Figure 25). When free in the lumen of the sac, all cells of B. plumatellae are rounded and different types are not apparent. (iv) Spore formation of T. bryosalmonae is by aggregation of stellate cells around sporoplasmogenic cells (Figure 25). In B. plumatellae, the cells comprising the spore are formed by division of a single sporogonic cell (Figure 17). (v) The polar capsule wall of B. plumatellae includes a lucent layer with radial striations (Figure 21). In T. bryosalmonae the lucent layer has no striations (Figures 27 and 28). 6. PHYLOGENETIC RELATIONSHIPS OF MYXOZOA 6.1. Inferred Higher Level Phylogeny 6.1.1.

Early Classifications

In one of the early attempts at classification, the Myxozoa were placed with Microsporidia and Actinosporea in the class


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Cnidosporidia Doflein, 1901. Microsporidia have since been recognized to be a distinct group, now accorded phylum rank, and, far from being primitive protozoans, they belong to the fungal lineage (Edlind et al., 1994, 1996). The simple unicellular or syncytial organization of trophic stages of myxozoans had led early workers to classify them with Protozoa but, with the recognition of their spore multicellularity (Sˇtolc, 1899), it was generally accepted that they were metazoans. Their status as a phylum, Myxozoa, was proposed by Grasse´ (1970). With the revelation of actinosporeans and myxosporeans as stages of the same life cycle (Markiw and Wolf, 1983), it was proposed that the class Actinosporea and the order Actinomyxidia should be suppressed (Kent et al., 1994), so that the phylum consisted of a single class Myxosporea Bu¨tschli, 1881. In recent years, myxozoan affinity within the Metazoa has been controversial. Weill (1938) was the first to draw attention to the remarkable similarity between myxozoan polar capsules and coelenterate (cnidarian) nematocysts (cnidocysts). Typically, in myxozoans, a capsular primordium in the capsulogenic cell has an external tube and both capsule and tube are surrounded by a corset of longitudinally orientated microtubules. As the external tube is resorbed, the central thread is wound into the capsule to form the polar filament and the microtubules are de-polymerized. The aperture of the polar capsule is sealed by a plug, which is of variable structure in spores of myxosporean and actinosporean stages and in malacosporean spores from bryozoans (Figures 8–10). The structure of the capsules of T. bryosalmonae in fish is not known. Morphogenesis of cnidarian nematocysts exactly follows this pattern. The hypothesis that myxozoans represent highly degenerate cnidarians has repeatedly gathered support (summarized by Siddall et al., 1995). 6.1.2. 18S rDNA Phylogeny The first voice of dissent was that of Smothers et al. (1994), who analysed 18S rDNA sequences from, among others, protozoans, myxozoans, cnidarians and a range of bilateral animals. They found that myxozoans were included in the bilateral animal clade, possibly

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as a sister group to the nematodes. This position was established whether the data were analysed by parsimony, distance or maximum likelihood methods. They found no support for a close relationship with the cnidarians. Similar results were obtained by Schlegel et al. (1996), who demonstrated grouping of the myxozoans with bilateral animals by parsimony and maximum likelihood analyses, while neighbour-joining analysis provided weak support for nematodes as a sister group. Almost concurrently with the study of Schlegel et al. (1996), Siddall et al. (1995) used a combination of morphological and 18S rDNA sequence data, including those for the parasitic narcomedusan Polypodium hydriforme. In all their analyses myxozoans and P. hydriforme grouped together. In each instance P. hydriforme was sister to the Myxozoa. In their analysis of partial 18S rDNA sequence data available for a wide range of cnidarians, this clade (i.e., Myxozoa þ P. hydriforme) nested within a monophyletic Cnidaria. However, in their analyses using complete 18S rDNA sequence data (including a smaller number of cnidarians) the Myxozoa þ P. hydriforme clade was a sister taxon to the Bilateria, leaving the Cnidaria paraphyletic. In consequence, Siddall et al. (1995) reiterated that myxozoans should no longer be regarded as protists and proposed that they should be assigned to the phylum Cnidaria as a sister taxon to P. hydriforme. Morphological characters used in their analyses to support the status of myxozoans as metazoans were multicellularity, cellular junctions with desmosome-like fibres, and terminal differentiation of some cells. They also discussed collagen, but it is now clear that this is not relevant since collagen is not apomorphic for metazoans (Celerin et al., 1996). Morphological characters used to support the status of myxozoans as cnidarians were the presence of nematocysts and radial symmetry. However, bilateral symmetry is exhibited by anthozoans, which are widely regarded as the most primitive class of cnidarians (Bridge et al., 1995; Odorico and Miller, 1997; Nielsen, 2001), and Buddenbrockia almost certainly exhibits bilateral symmetry. Thus, the only feature employed in this analysis of relevance to the affinity of Myxozoa with the Cnidaria was the presence of nematocysts/polar capsules.


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At about this time several 18S rDNA phylogenetic studies of lower metazoan taxa emphasized that long branch attraction could have dramatic effects on phylogenetic resolution. Hanelt et al. (1996) were interested in the phylogeny of Rhombozoa (dicyemids) and Orthonectida and included protists, diploblasts, myxozoans and a range of lower and higher triploblasts in their parsimony and minimum evolution analyses of full length 18S rDNA sequences. They found that myxozoans branched with triploblast phyla except when P. hydriforme sequences were included and reasoned that this was due to long branch attraction between the myxozoans and P. hydriforme. In addition, even when P. hydriforme sequences were included, myxozoans and P. hydriforme formed a clade intermediate between triploblasts and diploblasts and this clade was never within the diploblasts. (The aberrant nature of P. hydriforme is discussed in Section 6.1.3.) Other phylogenetic analyses based on full or nearly full length 18S rDNA sequences have similarly resulted in myxozoans grouping as the sister taxon to the Bilateria (Cavalier-Smith et al., 1996; Pawlowski et al., 1996; Winnipenninckx et al., 1998; Kim et al., 1999) or grouping variously with nematodes, flatworms or rhombozoans as sister to the rest of the Bilateria (Smothers et al., 1994; Katayama et al., 1995; Hanelt et al., 1996; Pawlowski et al., 1996; Schlegel et al., 1996; Winnipenninckx et al., 1998). None of these other analyses included P. hydriforme. Zrzavy´ et al. (1998) included P. hydriforme in their parsimony analyses and found that myxozoans grouped with cnidarians in analyses of morphological data alone and in analyses combining both morphological and partial 18S rDNA sequence information. However, in their analysis of 18S rDNA sequences alone, myxozoans and P. hydriforme formed the sister taxon to the Bilateria. By removing taxa to determine their effects on phylogenetic trees, Zrzavy´ et al. (1998) found that the inclusion of Myxozoa in the Cnidaria was strongly dependent on the presence of P. hydriforme in the analysis. Siddall and Whiting (1999) countered claims by Hanelt et al. (1996) that the clustering of myxozoans with P. hydriforme (revealed in the earlier study by Siddall et al., 1995) was due to long branch attraction.

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They showed that the placement of both Myxozoa and P. hydriforme remained the same whether both taxa or just one of them was included, demonstrating that long branch attraction was not a factor. However, it is noteworthy that in each of their their analyses of full length sequences of 18S rDNA, as opposed to partial sequences, the Myxozoa þ P. hydriforme group, or each of these taxa alone, occupied a position as sister to the Bilateria (Siddall and Whiting, 1999). In their parsimony analysis of the partial 18S rDNA sequences available for a broad range of cnidarians, Siddall and Whiting (1999) found that rhombozoans grouped with orthonectids, and they therefore criticized the earlier conclusion of Hanelt et al. (1996) (based on full length 18S rDNA sequences) that rhombozoans and orthonectids are not sister taxa. They argued that Hanelt et al. had insufficient grounds for excluding P. hydriforme from their phylogenetic analysis on the basis of sequence dissimilarity, when other taxa that were included showed similar or greater levels of dissimilarity. Curiously, the parsimony analyses by Siddall and Whiting (1999) of full length data revealed that rhombozoans and orthonectids were never sister taxa, whether Polypodium or myxozoans were included or excluded. While these issues are peripheral to the specific issue of long branch attraction between P. hydriforme and myxozoans, they further exemplify that analyses based on full length versus partial 18S rDNA sequences can lead to different conclusions regarding the placement of, for example Myxozoa, Rhombozoa and Orthonectida. These contradictory results suggest that phylogenetic resolution based on 18S rDNA sequences of some metazoans would require full length sequences for a broad range of taxa and would hence be computationally intense, or they should be restricted to inclusion of taxa with the shortest possible branches (see the contribution of Kim et al., 1999, cited below). Alternatively, other genes may be better suited for phylogenetic resolution. Indeed, Siddall et al. (1995) suggested that definitive genomic signatures retained in the highly simplified myxozoans, such as linear mitochondrial genomes or Hox genes, could prove to be useful in establishing the link with cnidarians. Unfortunately, Siddall and Whiting’s (1999) arguments regarding the phylogeny of Myxozoa


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did not take account of the work already conducted on Hox genes in myxozoans (Anderson et al., 1998). In an attempt to overcome the problem of long branch attraction in analyses of lower metazoan relationships based on 18S rDNA sequences, Kim et al. (1999) selected or generated full or nearly full length sequences of the species with the shortest branch to the ancestral node in each monophyletic lineage. They chose a species of Myxidium as representative of the Myxozoa and analysed sequences of representatives of the Choanozoa, Porifera, all classes of Cnidaria, including P. hydriforme, and representatives of deuterostome and protostome triploblasts. The distance analyses weakly supported a sister taxon relationship between Myxidium and P. hydriforme but neither of these fell within the Cnidaria, both being a sister to the triploblast phyla. Furthermore, in the maximum likelihood analyses, the long branches of Myxidium and P. hydriforme were not attracted to one another, Myxidium was a sister taxon to the triploblasts, and P. hydriforme occupied an unresolved position within the diploblasts.

6.1.3. Polypodium hydriforme Larval stages of Polypodium occur as intracellular parasites in the oocytes of acipenseriform fishes (Raikova, 1994). All parasitic stages are encircled by a polypoid unicellular trophamnion (formed from the second polar body). The two epithelia of the parasitic stage are inverted, with the inferred gastrodermal cells facing outwards towards the trophamnion and involved in the uptake of yolk, and the epidermal epithelium forming the lining to a fluid-filled internal cavity. Upon spawning, an adult stolon stage everts, reversing the epithelia, and a chain of ‘polyps’ is released which subsequently fragments into individual, free-living polyps. Although traditionally assigned to the Hydrozoa, it is not clear whether the adult stage of Polypodium is homologous with the polyp or medusoid stages of cnidarians (Hyman, 1940; Raikova, 1994). Furthermore, characters such as the presence of gonoducts and the complete separation of epidermal and muscle cells in Polypodium (see Raikova, 1994; Zrzavy´,

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2001) suggest the possibility of a non-cnidarian, triploblast nature. As we have seen earlier, inclusion of the only available full length 18S rDNA sequence for P. hydriforme in phylogenetic analyses has produced variable results, including paraphyly for cnidarians and a sister taxon relationship to the Bilateria along with the Myxozoa. Features of Polypodium which are shared with myxozoans are exclusive armament by atrichous isorhizae, fish parasitism, and reduction/loss of cilia. The presence of tubular cristae in mitochondria of both Polypodium and myxozoans is not phylogenetically relevant (Zrzavy´, 2001) since the configuration of cristae can reflect their metabolic activities (Vickerman et al., 1991). Without further data, the phylogenetic status of Polypodium and its relationship to the Myxozoa and even to the Cnidaria remains obscure. The disparate opinions on the diploblast versus triploblast origins of myxozoans, based on the phylogenetic analyses of the 18S rDNA sequences, might have reached stalemate had it not been for two other lines of evidence, namely the Hox gene study and the fortuitous encounters with B. plumatellae. 6.1.4.

Hox Gene Study

Anderson et al. (1998) obtained four central class Hox genes from ‘T. bryozoides’ (now B. plumatellae) and Myxidium lieberkuehni, which could be assigned to paralogy groups 4, 5, 6 or 7. Their analyses showed that myxozoans have Hox genes that can be clearly aligned to genes in bilaterians but not to the Hox-like genes (Cnox) of cnidarians, reflecting great divergence from common gene precursors (Ferrier and Holland, 2001; Finnerty, 2001). Furthermore, three of the myxozoan Hox sequences belong to the central class of Hox genes, a class which has so far lacked firm demonstration in cnidarians (Ferrier and Holland, 2001; Finnerty, 2001). In a similar way another central class Hox gene, designated DoxC, isolated from a dicyemid ‘mesozoan’ (or rhombozoan), indicated the dicyemid origins from the lophotrochozoan bilaterian animals (Kobayashi et al., 1999). The dicyemids and the myxozoans have similarly both undergone extreme character loss due to parasitism.


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6.1.5. Ancestral Body Form The bilaterian nature of myxozoans has recently been independently confirmed by ultrastructural studies of B. plumatellae. Canning et al. (2002) and Okamura et al. (2002a) described the vermiform stage of B. plumatellae from several bryozoan species (Section 5.1.2.b) as having outer and inner cell layers separated by four longitudinal muscle blocks but otherwise lacking body organs, including a gut. Polar capsules were present in spores that arose through differentiation of cells originally derived from the inner cell layer. In the case of B. plumatellae in H. punctata, polar capsules were present in cells of the outer layer. The derivation of polar capsules from both layers demonstrates that the spores could not be attributed to a myxozoan infection in a non-myxozoan ‘worm’, but that the ‘worm’ itself was a myxozoan. Confirmation of the myxozoan nature of Buddenbrockia and further evidence that the polar capsules were indigenous was provided by analysis of 18S rDNA sequences. The use of universal primers resulted in amplification of sequences identified as myxozoan, bryozoan (host), and those of the protist Stentor (an associated contaminant), suggesting that all eukaryotic 18S rDNA genes present in the sample were amplified. There are, thus, three lines of evidence that support the theory that Myxozoa are highly degenerate bilaterian animals. (i) Most of the phylogenetic analyses using 18S rDNA sequences placed myxozoans within the Bilateria (Smothers et al., 1994; Katayama et al., 1995; Cavalier-Smith et al., 1996; Hanelt et al., 1996; Pawlowski et al., 1996; Schlegel et al., 1996; Winnipenninckx et al., 1998; Kim et al., 1999). Only the 18S rDNA analyses of Siddall et al. (1995), Zrzavy´ et al. (1998) and Siddall and Whiting (1999) placed them within the Cnidaria and then only when P. hydriforme was included. (ii) Some of the structural characters of an ancestral bilaterian animal have been retained in the vermiform stage of B. plumatellae. (iii) The large divergence between myxozoan Hox and cnidarian Cnox genes.

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6.2. Myxozoan Polar Capsules and Cnidarian Nematocysts The balance of evidence is now heavily weighted in favour of Myxozoa as a phylum within the Bilateria. The major stumbling block to acceptance of this phylogenetic position is the presence in Myxozoa of polar capsules, which are so similar to nematocysts in their ontogeny and mature structure that they would appear to have the same origin. One explanation for the similarity of polar capsules and nematocysts would be common or independent acquisition via endosymbiosis. Alternatively, these structures could either have a non-symbiotic origin, explained by convergent evolution or gene transfer, or be homologous and derived from a common ancestor. Other issues relevant to the evolution of the Myxozoa are the identity of the original hosts, how the digenetic cycles evolved, and what steps towards morphological simplicity due to parasitism can be postulated based on presentday genera. The remainder of our review deals with these questions. 6.2.1.

Endosymbiont Acquisition of Polar Capsules and Nematocysts

Shostak (1993), quoting a personal communication from Pierre Tardent, developed the argument that, rather than ‘re-inventing the wheel’, nematocysts in cnidarians could have resulted from a symbiotic event involving a cnidocyst-bearing protist. Okamura et al. (2002a) suggested that the notable resemblance of polar capsules in myxozoans to nematocysts in cnidarians might reflect independent incorporation of eukaryotic symbionts in these two taxa. Below we weigh the evidence for such postulates by considering the multiple origins of endosymbiotic association, the presence of similar extrusible organelles in the lower Metazoa, and candidate endosymbionts for polar capsules and nematocysts. (a) Multiple acquisition and incorporation of endosymbionts. There is increasing evidence that obligate endosymbiotic associations have arisen on multiple occasions in evolutionary time. They have resulted from direct acquisition of prokaryotes (primary endosymbiosis) and


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by retention of plastids from eukaryotic symbionts after loss of most traces of the original eukaryote (secondary endosymbiosis) (see, e.g., Gray, 1992; van Hoek et al., 2000; Selosse et al., 2001). In both types, gene transfer between symbiont and host demonstrates their increasing interdependence. Gene transfer has occurred extensively in obligate primary endosymbionts such as mitochondria and various plastids (Selosse et al., 2001) and has also occurred in secondary endosymbioses. It has been estimated that about eight percent of the genes of the malaria parasite, Plasmodium falciparum, are of primary (mitochondrial) or secondary (plastid) endosymbiotic origin (Wilson, 2002). Gene transfer may have resulted from selection for rapid organelle replication, selection against higher organelle mutation rates, or selection for small genome size in organelles (Selosse et al., 2001). DNA still remaining in many of these organelles is one legacy of their separate evolutionary origins but genetic erosion is a general trend and, in some cases, such as in hydrogenosomes (Embley et al., 1997), DNA transfer has been complete. Relictual mitochondria in microsporidia (Williams et al., 2002) further exemplify such great genetic erosion and it appears likely that such organelles are redeployed following loss of original function (e.g., aerobic respiration) and gain a new biochemical role. In B. plumatellae, parasitic in H. punctata, the capsulogenic cells and their polar capsules are present in the outer cell layer, an obvious original site for the incorporation of symbionts, and the site where extrusible organelles occur in other triploblasts (Section 6.2.1.b). Spores with polar capsules in capsulogenic cells are formed in the body cavity of Buddenbrockia but only the sporoplasm within the spore carries the genetic information for synthesis of polar capsules in the next generation. Thus, if polar capsules (and nematocysts) are derived from endosymbionts, all the genetic material directing their synthesis must have been transferred from the symbiont to the myxozoan (or cnidarian) and, thus, the degree of integration of the endosymbionts in myxozoans and cnidarians may be unparalleled in evolutionary history. Other organelles deriving from symbioses are transmitted vertically through gametes, but polar capsules and nematocysts are derived de novo in each generation.

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(b) Occurrence of extrusible organelles in the Metazoa. Nematocystlike organelles have been described in several lower invertebrates. These include paracnids in the epidermis of some turbellarians (Karling, 1966; Sopott-Ehlers, 1981) and pseudocnids in the proboscis epithelium of some nemerteans (Turbeville, 1991; Montalvo et al., 1998). This range suggests the possibility that precursor nematocyst/polar capsule organelles evolved in the common ancester of Cnidaria and Bilateria, and have been retained in at least some of the triploblast phyla. Multiple endosymbiotic origins are, however, theoretically possible. Gschwentner et al. (1999) have also pointed out that extrusible organelles are common to lower eumetazoans, including nematocysts in cnidarians, polar capsules in myxozoans, colloblasts in ctenophores, sagittocysts, rhabdites, and paracnids in turbellarians, and rhabdoids in nemertines and gastrotrichs. Gschwentner et al. (1999) referred to such organelles collectively as extrusomes, and pointed out the similarity of sagittocyst discharge to tube-eversion discharge of nematocysts and the fact that some cnidarian nematocysts (ptychocysts) are discharged whole, as occurs in many other invertebrate extrusomes. The relationships of this diverse array of extrusomes are unknown. An extreme hypothesis, suggested by the arguments of Gschwentner et al. (1999), is a common origin for all invertebrate extrusomes followed by adaptive divergence. A more conservative hypothesis would be a common origin for the explicitly nematocyst-like organelles. Gschwentner et al. (1999) further suggested that the origin of extrusomes in lower invertebrates correlates with the origin of the eumetazoan gut, with extrusomes aiding in food capture within a water-filled gastrovascular cavity. They proposed that this arrangement set the stage for the development of true predatory life styles. This hypothesis would provide an explanation for the widespread loss of nematocyst-like organelles in taxa from mobile benthic groups, in which other mechanisms of food capture evolved, and retention in cnidarians and myxozoans for food capture and host attachment.


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(c) Candidate symbionts for nematocysts and polar capsules. Buss and Seilacher (1994) proposed that nematocysts and polar capsules may be derived through an endosymbiotic association with microsporidia. The polar tube of microsporidia is an intracytoplasmic organelle that functions by eversion to form a hollow tube, through which the nucleus and cytoplasm are passed to enter a new host cell, leaving behind the original plasma membrane. This mechanism differs fundamentally from that of nematocysts and polar capsules, where the filament is isolated in its capsule within a capsulogenic cell and is alone everted as a capture or attachment organelle, leaving the other cell components behind. Furthermore, microsporidia are no longer considered to be ancient eukaryotes as was first indicated by 16S rDNA sequences (Vossbrinck et al., 1987) but, according to tubulin gene sequences, appear to have been derived from fungi (Edlind et al., 1994, 1996; reviewed by Weiss and Vossbrinck, 1999). Even if microsporidia are unlikely candidates for myxozoan endosymbiosis, their possession of an eversible tube demonstrates that such structures have evolved more than once. Other candidate endosymbionts whose incorporation may have led to nematocyst-like organelles are ancestral dinoflagellates (Dinozoa ¼ Dinophyta) (Shostak, 1993). These are mostly free-living alveolate protists, of which about half of the species are photosynthetic. Some, lacking chloroplasts, are parasitic in marine animals such as molluscs; other parasitic forms, with chloroplasts (zooxanthellae), are endosymbionts in tissues of cnidarians such as corals and sea anemones. Sometimes cnidarians acquire their zooxanthellae de novo from the environment but in other cnidarians they are passed to the next generation in egg cytoplasm. Shostak (1993) discussed the resemblance of cnidarian nematocysts to a complex organelle in the extant dinoflagellate Polykrikos (Polykrikidae). Part of the complex, a nematocyst-like structure with a coiled tube, is connected to an anterior chamber containing a stylet and sealed by an operculum (Westfall et al., 1983). A trigger mechanism associated with the other part of the complex, the taeniocyst, is indicative of a discharge mechanism and the extruded tube, still attached to the everted anterior chamber, has been observed (Chatton, 1914).

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The idea that nematocysts were incorporated as symbionts into corals and other cnidarians by association with dinoflagellates has some attraction. The fossil record based on cysts dates mainly from the Triassic period and this is supported by isotopic analysis confirming the presence of zooxanthellae symbionts in fossil corals from the late Triassic (Stanley and Swart, 1995). There are two fossil records from the Palaeozoic era which are possible dinoflagellates but fossils of Polykrikidae are not found until the Oligocene period. Thus, if the dinoflagellates emerged only in the Mesozoic era they could not have provided nematocysts as symbionts of cnidarians. Recently, dinoflagellate specific biological markers (dinosteranes and 4-methyl-24-ethylcholestane) have been used to identify certain acid-resistant microfossils resembling dinoflagellate cysts (acritarchs) in Early Cambrian sediments as dinoflagellate ancestors (Moldowan and Talyzina, 1998). This supports the widespread belief that prokaryotic features of dinoflagellates, such as absence of histones and nucleosomes, are indicative of evolutionary divergence of dinoflagellates as early as the Precambrian era. If so, dinoflagellates could have been incorporated into ancestral cnidarians and myxozoans. Although nematocysts are regarded as diagnostic of cnidarians, they are not preserved in the fossil record and so it cannot be determined whether cnidarians have always possessed them. Buss and Seilacher (1994) have suggested that the Precambrian Ediacaran vendobiont fossils were gutless cnidarians that lacked nematocysts and derived nutrition through photosynthetic or chemosynthetic symbionts. This hypothesis indicates a later endosymbiotic origin of nematocysts. The first known cnidarian fossils date from the Cambrian. Unlike the Ediacaran fauna, these were benthic rugose corals and, in view of the reliance on nematocysts by all living cnidarians, it is difficult to imagine how these fossil corals, which are presumed to have had a mouth and gut, would have achieved prey capture without them. It thus seems likely that, if nematocysts were attained through endosymbiosis, this must have been an ancient event. Furthermore, the complete erosion of the putative endosymbiont genome implies that this was a very ancient event indeed.


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Cryptomonad flagellates possess organelles (ejectosomes) consisting of a tightly coiled ribbon housed within a membrane (Dodge, 1969). The ribbons are not everted but roll up to form a tube when ejected. Cryptomonads have themselves incorporated photosynthetic symbionts, of which the nucleomorph and chloroplast are remnants (see Douglas et al., 1991). The evident resemblance of cryptomonad ejectosomes to cnidarian nematocysts has already been noted (Hovasse and Mignot, 1975). Another protistan group with nematocyst-like extrusive organelles are the karyorelictid ciliates. Their organelles are membrane-bound capsules containing a coiled, multilayered filament, and have an apical cap-like structure consisting of longitudinal fibrils and pierced by an apical channel (Raikov, 1992). Such organelles occur nowhere else among the protists (Raikov, 1992). Both morphological (Raikov, 1992) and molecular sequence data based on small subunit (ssu)rRNA (Hirt et al., 1995), histone H4 (Bernhard and Schlegel, 1998), phosphoglucose kinase (Tourancheau et al., 1998), and large subunit rRNA sequences (Fleury et al., 1992) indicate that karyorelictids represent an ancient branch of ciliates. The ancient nature of karyorelictids suggests that their ancestral forms may be good candidates for endosymbionts whose incorporation led to nematocyst-like organelles. Endosymbiosis involving myxozoans as symbionts in cnidarians is a highly unlikely explanation for the presence of nematocysts and polar capsules, as it would entail an unparalleled level of complexity of symbiotic partners, both being metazoans. If such an unparalleled event had occurred, it is more likely to have entailed derivation of myxozoan endosymbionts (polar capsules) from cnidarian nematocysts. Since differences in Hox genes (Section 6.1.4) and 18S rDNA phylogenetic analyses (Section 6.1.2) both indicate that myxozoans emerged after the cnidarians (Schlegel et al., 1996), derivation of nematocysts from myxozoan endosymbionts would require that cnidarians lacked nematocysts until after the evolution of the Myxozoa. In summary, the possibility that the variety of extrusible organelles in several metazoan phyla had their origin(s) in single or multiple endosymbiotic events involving eukaryotes as both host and symbiont

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is conceivable since (i) the occurrence of eukaryotic symbionts is well documented—e.g., cryptomonads (Douglas et al., 1991), apicomplexan parasites (reviewed by Wilson, 2002), and others (Selosse et al., 2001)—and (ii) strikingly similar extrusible filaments are commonly found in extant protists and may have evolved after incorporation to assist in food capture. However, the suggestion by Tyler et al. (1980) that adhesive cells of the lower invertebrates are modified cilia or sensory cells provides an alternative explanation for the origins of at least some extrusible organelles. Phylogenetic analyses of the genes coding for the array of extrusible organelles and their potential homologues may eventually provide answers to the question of the origins of these curious structures. 6.2.2.

Non-symbiotic Origins of Polar Capsules and Nematocysts

Other explanations for the derivation of polar capsules and nematocysts are convergent evolution, gene transfer, or the fact that they could be homologous structures. The last relates to evidence that cnidarians are actually triploblasts. (a) Convergent evolution. Unless myxozoans are cnidarians, which is unsupported by their Hox genes, an alternate explanation to endosymbiosis for the presence of nematocysts in cnidarians and polar capsules in myxozoans is convergent evolution. The abundance and diversity of structure of extrusible organelles in protists and lower metazoans suggests that there may have been strong selective pressure for their evolution. However, even the most famous apparent example of convergent evolution, namely the presence of complex eyes in cephalopods and vertebrates, is not as strong a case as was widely believed. The universality of Pax6 and rhodopsin in the Metazoa and the demonstrated role of Pax6 as a master control for eye morphogenesis suggest that the different types of eye found in the Metazoa are derived from a single prototypic eye (for review, see Gehring and Ikeo, 1999). The development of increasingly complex eyes may then be explained by subsequent divergent, parallel and convergent evolution achieved by the intercalation of genes into


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the cascade of genes associated with eye morphogenesis, via gene duplication and divergence, recruitment of novel genes, and recombination of different genes by evolutionary tinkering. Such evolutionary steps may also explain the array of nematocyst-like and extrusible organelles amongst the lower invertebrates and perhaps even the eversible tubes of microsporidia and nematocyst-like organelles of dinoflagellates. (b) Gene transfer. Although nearly all eukaryotic genes are inherited by common descent, lateral gene transfer which introduces genes from independent lineages occasionally occurs. This process occurs extensively between prokaryotes, and, as discussed in Section 6.2.1.a, is common in endosymbiotic associations. A recent example, apparently not involving endosymbiosis, is a cellulose synthase gene in an ascidian, which is involved in the manufacture of cellulose for the ascidian tunic (Dehal et al., 2002) and which may have been imported from nitrogen-fixing bacteria. This gene is present in plants, but its presence in ascidians is unique for animals. We suggest that lateral gene transfer that would result in the manufacture of organelles as complex as polar capsules is unlikely, since the assembly of such structures would necessitate the incorporation of multiple genes. (c) Homology of polar capsules and nematocysts. In contrast to traditional views, there is a plethora of evidence which indicates that Cnidaria fall within a monophyletic triploblast group, as sister to the remaining triploblasts. Other evidence suggests that they may be paraphyletic, with Anthozoa being sister to the other Cnidaria þ triploblasts, or that they are a monophyletic diploblast group (the classical view). In any of these scenarios, a common origin for nematocysts (in Cnidaria) and polar capsules (in Myxozoa) may be readily envisaged. Indeed, possession of extrusible organelles could be seen as a synapomorphy for the triploblasts (including Cnidaria) or for the triploblasts þ Cnidaria, if the latter are excluded from the triploblasts proper. Of course such scenarios entail early and/or widespread loss or modificiation of such extrusible organelles in the remainder of the triploblasts.

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Hyman (1940) rejected the distinction between diploblastic and triploblastic organization. She regarded the connective tissue layer between the outer and inner epithelia of sponges, cnidarians and ctenophores as fundamentally triploblastic and equivalent to the mesoderm of higher animals. She observed that this central layer is simply developed to different degrees and is invariably present, except in Hydrozoa (which we now know to be derived). Pantin (1960) similarly emphasized that the Anthozoa are triploblastic and that the mesogloea of Scyphozoa and Ctenophora frequently contains cells. Fautin and Mariscal (1991) argued that embryogenic processes in cnidarians and other animals provide further evidence for cnidarian triploblasty. It is now clear that the mesoderm (and, to a lesser extent, the endoderm) can develop in different ways, at different times, and may even have multiple origins (see discussions by Willmer, 1990 and Nielsen, 2001). Nielsen (2001) suggested that the different types of mesoderm formation may reflect an unspecified ancestral ability to proliferate extraepithelial cells, but argued that, because bilaterian mesodermal tissues are easily distinguished from the isolated cells in the mesogloea of cnidarians, there is a significant difference between ‘mesoderm’ of bilaterians and ‘mesogloea’ of cnidarians. Several recent studies of the developmental biology of cnidarians lend further credence to a triploblast status for the cnidarians and provide evidence that the origins of bilateral symmetry lay in the common ancestor of the Cnidaria and Bilateria. Boero et al. (1998) argued that, in hydroids, a localized triploblast structure develops in medusoid buds by proliferation into the blastocoel and cavitation to form a coelom lined by myoepithelial cells. They suggested that this represents the origin of a coelomate bilaterian, although Nielsen (2001) believed that the process merely reflected budding. Evidence that the proliferated cells were equivalent to bilaterian mesoderm was provided by the detection of a mesoderm specification factor Twist in particular stages during development of planulae and medusae in the scyphozoan cnidarian, Podocoryne carneae (Spring et al., 2000), although it is unclear whether the functional role is equivalent in cnidarians and bilaterians. Gro¨ger and Schmidt (2001) reported on the existence of a nervous system in


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P. carneae that develops gradually in an anterior to posterior direction, and Yanze et al. (2001) found expression of Hox-like genes in P. carneae that were involved in determination of polarity. Although not all cnidarian Hox-like genes have orthologues in bilaterians, at least part of the control of polarity is conserved between cnidarians and bilaterians and thus the genes must have a common ancestry. In addition, it is now evident from a variety of recent data that the origin of one, or possibly two, Hox clusters dates from a precnidarian–bilaterian divergence and that middle group Hox genes were apparently either lost in the cnidarians or arose only in the bilaterian lineage (see review by Ferrier and Holland, 2001). These findings demonstrate that some of the genes that play essential roles in bilaterian body patterning are present in cnidarians, although there has subsequently been great divergence between the two groups. On the basis of apparent mesoderm formation in scyphozoans, it has been suggested that cnidarians have been a centre for metazoan radiation, with triploblasts deriving from cnidarians (Boero et al., 1998). This view suggests that the cnidarians may be paraphyletic, with anthozoans forming a basal metazoan branch and the Bilateria deriving from the Medusozoa (scyphozoans, cubozoans and hydrozoans). This is consistent with some phylogenetic analyses (e.g., that by Winnipenninckx et al., 1998) but contrasts with others in which the cnidarians comprise a clearly monophyletic group that, in the case of fully resolved trees, displays a sister group relationship to the triploblasts (see, e.g., Cavalier-Smith et al., 1996; Littlewood et al., 1998; Zrzavy´ et al., 1998; Peterson and Eernisse, 2001). It is possible that a cnidarian-like organism with a planuloid body is the common ancestor for both ‘triploblasts’ and ‘diploblasts’ (for review, see Holland, 2000). If such an organism possessed nematocysts, these may have been retained in the parasitic myxozoans as a means of host attachment, but lost or modifed in other bilaterians which evolved predatory lifestyles (Section 6.2.1.b). Regardless of the specific relationship between the cnidarians and bilaterians, it is clear that they have both undergone subsequent divergence from a common ancestor, which is reflected in their contrasting body plans and Hox gene complements.

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7. MYXOZOAN LINEAGES AND EVOLUTION OF PARASITISM Sequence data from a range of myxozoan genera have indicated that there was an ancient split (Kent et al., 1998; Anderson et al., 1999b) that led to the class Malacosporea, which utilize bryozoans as invertebrate hosts, and the class Myxosporea, which utilize annelids. The original parasitic event would have led to a direct cycle with one host type and spores liberated into the environment for reinfection. If parasitism co-evolved with the host phyla, invertebrate hosts are likely to have been the first hosts of the ancestral myxozoans. Putative lophophorate relatives of phylactolaemate bryozoans include marine bryozoans and phoronids (Larwood and Taylor, 1979). Marine bryozoans date from the Ordovician. Possible trace fossils of the softbodied phoronids date from the late Precambrian (as a burrow), but borings more diagnostic of modern phoronids date from the Devonian (Taylor, 1993). The fossil record of phylactolaemate bryozoans dates to the Permian (Vinogradov, 1996), but a much older origin for the Phylactolaemata is entirely possible, particularly since the fossilized remains of their statoblasts are unlikely to be widely recognized by palaeontologists. Good fossil annelids date from the Cambrian but some fossils from the Precambrian Ediacaran assemblage may be annelids (Clarkson, 1986; Budd and Jensen, 2000). In adapting to bryozoans, the infective sporoplasms of malacosporeans would have become surrounded by protective cells to withstand their dispersal into the aquatic environment, dispensing with a free-swimming larval stage. The loss of ciliary basal bodies at this stage may have been accompanied by loss of centriolar involvement in nuclear division. Similarly, the myxosporeans would have become adapted to annelids and evolved the triradial actinospores. From the direct, one-host life cycle, selection for higher levels of transmission may have resulted in the incorporation of vertebrate hosts (Poulin, 1998). The alternative, that fish were the primary hosts and that invertebrates were added as alternate hosts, is unlikely. The Actinopterygii (ray-finned fishes) include the Chondrostei, the Holostei and the Teleostei. Although the chondrosteans were common in the Upper


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Paleozoic and holosteans in the Mesozoic, the teleosts did not become common until the Cretaceous (McKerrow, 1978). The availability of teleost hosts thus post-dates the availability of invertebrate hosts. Further evidence that fish were not the primary hosts is the occurrence of the myxozoan sexual cycle in invertebrate hosts. By analogy with digenean life cycles, the host in which the sexual cycle of the parasite takes place is the definitive host and the host in which asexual proliferation occurs is the intermediate host. As development of Myxozoa in fish is entirely asexual, except for the possible restoration of diploidy in malacosporean cycles, the incorporation of intermediate vertebrate hosts is likely to have evolved due to the selective advantage of asexual proliferation. In the myxosporeans another type of spore, the myxospore, evolved to protect the asexual sporoplasm when released into the environment. In most myxosporeans the alternation of hosts is probably obligate. However, another phase in the evolution of myxosporean life cycles may be represented by the direct transmission between sparid fish by trophozoites as recorded by Diamant (1997), Redondo et al. (2002) and Yasuda et al. (2002), in which the putative annelid hosts have been omitted making indirect transmission no longer obligatory, as occurs in some trematodes (Poulin and Cribb, 2002). Insufficient data are available on malacosporean cycles to substantiate whether the bicapsulid spores formed in salmonid fish are infective to bryozoans. An indirect life cycle involving salmonid fish cannot be obligate for T. bryosalmonae, since salmonids are absent from some sites supporting infected bryozoans. 7.1. Morphological Simplification Due to Parasitism If myxozoans are bilateral animals, then at one stage in their history they would have had a triploblast organization with gut, muscle, nervous and reproductive tissue. While losing the gut, B. plumatellae has retained a worm-like body form and four muscle blocks which undergo co-ordinated movements. A further step in simplification is exemplified by its sac stage (‘T. bryozoides’), in which the muscles have been lost and the inner cell layer is quickly disassembled. Loss of

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the gut is a feature of other lower metazoan bilaterians which have become parasitic, such as the Rhombozoa (dicyemids), Orthonectida and a few species of Nematoda. In the first two phyla, the body is organized as an outer ciliated epithelium surrounding a single cell or a solid multicellular or plasmodial core. In the case of orthonectids a cuticle develops between the cilia on the epithelial surface. There is no body cavity, no gut, and the inner cells are reproductive. However, the emplacement of both groups in the Bilateria has been indicated by 18S rDNA sequence analysis (Katayama et al., 1995; Hanelt et al., 1996) and that of the dicyemids also by Hox gene analysis (Kobayashi et al., 1999). In the body plan of nematodes there are four longitudinal muscle blocks situated in dorsal, ventral and lateral positions. Some nematodes, such as the mermithids, have a non-feeding stage in which the gut is absent. There are others, e.g. Rhabdonema, in which feeding continues in the absence of a gut, via microvillous extensions of the epidermis which project through the cuticle (Riding, 1970). Cross sections of Buddenbrockia also show four longitudinal muscle blocks and a body cavity without a gut, suggestive of a common ancestry with nematodes or their precursors. Monteiro et al. (2002) suggested that Buddenbrockia (and thus the Myxozoa) occupies a more basal bilaterian position. Although we hold the opinion that myxozoans are specialized bilateral animals, it is necessary to point out that the vermiform stage of Buddenbrockia is superficially similar in structure to the planula larva of cnidarians. Although adult cnidarians develop a functional gut by opening the central cavity to the exterior, the planula consists only of an outer layer of ciliated cells, some armed with nematocysts, and an inner mass of cells which eventually flattens to form an inner layer around a central cavity. The planula differs from Buddenbrockia in having ciliated outer cells and no muscle blocks, and it is a larval stage, developing from a fertilized egg and proceeding to the adult stage in which the gametes are differentiated to complete the cycle. In contrast, Buddenbrockia is a terminal stage undergoing sexual reproduction, producing spores for the next generation and then dying. Thus the planula and the ‘worm’ would not appear to be equivalent. Ciliated cells are not known in Myxozoa.


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The organisms in the class Malacosporea appear to have diverged from the main evolutionary line of the Myxozoa before the radiation that gave rise to the numerous better-known genera in the class Myxosporea (Anderson et al., 1999a, b). In the malacosporean stages in bryozoans the tendency has been towards retention of ancestral characters and almost identical spore structures in the two known genera. In contrast, myxosporeans show a considerable reduction of body complexity in stages in the vertebrate hosts, as well as a wide range of spore structures in both the invertebrate and vertebrate stages. One factor that may have been inhibitory to diversification in malacosporeans is the lack of strengthened valves around the sporoplasms. Infectivity of the spores is lost in less than 12 h (De Kinkelin et al., 2002). The protective valves of both phases of the myxosporean life cycle may have enabled myxosporeans to establish themselves in new environments, with the accompanying potential for adaptation to new hosts. In both classes there is such a degree of simplification of presporogonic stages, especially in the vertebrate hosts, that it is not surprising that myxozoans were taken to be protists. Nonetheless, as bilaterian animals, the ancestral free-living myxozoans would have had a triploblast organization with ectoderm and endoderm derived by gastrulation, and mesoderm from which the musculature, excretory and reproductive tissue would have differentiated. Gametes might have been anisogamous. Among the present day myxozoans, all of which are parasitic, only the malacosporeans in their bryozoan hosts have retained a semblance of triploblasty. The vermiform stage of Buddenbrockia has retained the musculature but the gut with its endodermal and mesodermal components has been lost and the reproductive cells are included in the inner (mesodermal) layer of cells, which probably also has a nutritional and secretory function. In the sac stages of both Buddenbrockia and Tetracapsuloides even the musculature and a stable inner cell layer have disappeared. No evidence of male and female gamete differentiation was obtained in the recent ultrastructural studies of these genera (Canning et al., 1996, 2002; Okamura et al., 2002a), although meiosis was observed during

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sporogony. Nevertheless, the possibility of gametogenesis with spermatozoa and ova, followed by fertilization as described by Schro¨der (1910), cannot yet be ruled out entirely. In the class Myxosporea, the actinosporean stages in annelids have retained a true sexual cycle, with anisogamous gametes and zygotes, before spore formation. The strange process in actinosporeans, by which sporoplasms develop separately from the capsulogenic and valvogenic cells in the pansporocyst and the valve cells open to allow entry of sporoplasms, must have evolved as a mechanism to protect the sporoplasms before attachment to, and entry into, a new host. This has no counterpart in malacosporeans, the sporoplasms being encased in the protective cells from their outset. In the vertebrate hosts none of the known species exhibits any triploblast morphological features. The known stages, except those of Myxobolus pendula, are either multinucleate syncytial plasmodia or single cells. The plasmodia, whether histozoic or coelozoic, have a simple plasma membrane, usually with an underlying cytoplasmic layer devoid of cell organelles, around the core of cytoplasm containing somatic and generative nuclei. Plasmodia could have arisen by proliferation of the inner cell layer to pack the central cavity followed by loss of cell walls, thus becoming syncytial. Sporogony involves the endogenous formation of cells around generative nuclei and enclosure of sporogonic cells by pericytes. Spore formation is continuous until the plasmodium becomes packed with spores. M. pendula (Figure 7) might represent an intermediate stage between the sac-like and plasmodial body forms, in which some of the cells of the outer and inner layers have been retained as columnar cells of putative secretory function surrounding the syncytial core (Martyn et al., 2002). Many of the coelozoic myxosporeans do not form multinucleate plasmodia but proliferate by endogeny within uninucleate pseudoplasmodia, which attach themselves to lining epithelia of the urinary bladder or gall bladder (Sitja`-Bobadilla and Alvarez-Pellitero, 1993; Canning et al., 1999a). After proliferation, the endogenous cell becomes sporogonic and forms two spores (disporic), while the primary cell becomes the pericyte. The pseudoplasmodium, then, appears to be the equivalent of one pericyte and sporogonic cell unit in


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a large plasmodium and may have arisen by division of cells derived from the sporoplasm in place of syncitium formation. In Sphaerospora renicola, the proliferative stage is represented by cells which reproduce repeatedly by endogeny up to a tertiary level in blood, swimbladder or renal tubules before becoming sporogonic pseudoplasmodia. An equivalent level of simplicity is exhibited by T. bryosalmonae stages in fish hosts (and probably other malacosporeans if they have become established in fish), where proliferative (PKX) cells can reproduce extracellularly up to a quaternary level of endogeny in host blood and solid tissues before producing spores in kidney tubule lumina. Details of sporogenesis in fish hosts are unknown for T. bryosalmonae. Reduction of body form from triploblast organization, through multinucleate plasmodia to uninucleate pseudoplasmodia, is an extreme example of simplification due to parasitism. ACKNOWLEDGEMENTS We thank the following persons for help in preparing this review: G. Azevedo, S.S. Desser, M. El-Matbouli, S.W. Feist, J. Lom and S. Tops for provision of micrographs; J. Dodge, D.E. Ferrier, P.M. Hammond, J. Lom, S. Tops, K. Vickerman, J. Webster and R.J.M. Wilson for information and discussion; and especially A. Curry for electron microscopy. The original work by the authors reported in this review was supported by NERC grants GR3/09956, GR3/11068, GR9/04271, NER/A/S/1999/00075, and NER/B/S/2000/00336. Preparation of the review was carried out during the tenure of a Research Fellowship awarded to B.O. by the Leverhulme Trust. REFERENCES Anderson, C.L., Canning, E.U. and Okamura, B. (1998). A triploblast origin for Myxozoa? Nature 392, 346. Anderson, C.L., Canning, E.U. and Okamura, B. (1999a). 18S rDNA sequences indicate that the PKX organism parasitizes Bryozoa. Bulletin of the European Association of Fish Pathologists 19, 94–97.

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The Mitochondrial Genomics of Parasitic Nematodes of Socio-Economic Importance: Recent Progress, and Implications for Population Genetics and Systematics Min Hu, Neil B. Chilton and Robin B. Gasser Department of Veterinary Science, The University of Melbourne, Werribee, Victoria 3030, Australia

Abstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Background on the Mitochondrial Genomics of Animals . . . . . . 2.1. General and Historical Perspective . . . . . . . . . . . . . . . 2.2. Structural and Functional Aspects . . . . . . . . . . . . . . . 2.3. Significance of Studying Mitochondrial Genomes . . . . . . 3. The Mitochondrial Genomes of Parasitic Helminths . . . . . . . . . 3.1. Brief Account of Mitochondrial Genomes of Flatworms . . . 3.2. Mitochondrial Genomes of Nematodes. . . . . . . . . . . . . 3.3. Approaches for the Sequencing of Mitochondrial Genomes from Nematodes and Recent Technological Improvements . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4. Recent Progress in Mitochondrial Genomics of Nematodes and Evolutionary Implications . . . . . . . . . . . 4. Mitochondrial Gene Markers for Studying the Molecular Systematics and Population Genetics of Parasitic Nematodes . . . 4.1. Species Identification and Differentiation . . . . . . . . . . . 4.2. Systematic Studies Using Individual Mitochondrial Gene Sequences . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3. Population Genetic Investigations . . . . . . . . . . . . . . . 4.4. Technological Considerations for Population Genetic Investigations . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5. Recent Population Genetic Investigations by Mutation Scanning Analysis of Selected Mitochondrial Gene Markers


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5. Conclusions, Perspectives and Prospects . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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ABSTRACT Mitochondria are subcellular organelles in which oxidative phosphorylation and other important biochemical functions take place within the cell. Within these organelles is a genome, called the mitochondrial (mt) genome, which is distinct from, but cooperates closely with the nuclear genome of the cell. Investigating mt genomes has significant implications for various fundamental research areas, including mt biochemistry and physiology, and, importantly, such genomes provide a rich source of markers for population genetic and systematic studies. While 250 complete mt genome sequences have been determined for a range of metazoan organisms from various phyla, few of these represent parasitic helminths. Until 1998, only two mt genome sequences had been determined for parasitic nematodes, in spite of their socio-economic importance and the need for investigations into their population genetics, taxonomy and evolution. However, since that time, there has been some progress. The main focus of the present chapter is to review the state of knowledge of the mt genomics for parasitic nematodes, to describe recent technological improvements to mt genome sequencing, to summarize applications of mt gene markers for studying the systematics and population genetics of parasitic nematodes, and to emphasize prospects and opportunities for future research in these areas.

1. INTRODUCTION The Nematoda represents a large phylum in the animal kingdom, containing a diverse range of species with different life histories (Hugot et al., 2001). Many species of parasitic nematodes are pathogens of plants and/or animals, including humans (Blaxter and

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Bird, 1997; Anderson, 2000), causing significant diseases (Eisenback and Triantaphyllou, 1991; Albonico et al., 1999; McCarthy and Moore, 2000) and major socio-economic losses. Central to the control of these parasites is knowledge of their population genetics, which can also have important implications for understanding ecology, transmission patterns and the development of drug resistance (Anderson et al., 1998; Blouin, 1998; Viney, 1998; Gasser and Newton, 2000). The basis for investigating population genetics is the accurate analysis of genetic variation, which is generally considered to be widespread in parasite populations (e.g. Grant, 1994; Anderson et al., 1998; Blouin, 1998; Viney, 1998). Molecular methods have proven useful for assessing genetic variation within and between parasite populations (Nadler, 1990; Grant, 1994; McManus and Bowles, 1996; Gasser and Newton, 2000). Mitochondrial (mt) DNA markers have been considered to be particularly applicable to population genetics and systematic investigations due to their high mutation rates and proposed maternal inheritance (Avise et al., 1987; Avise, 1991, 1994, 1998; Anderson et al., 1998; Blouin, 1998, 2002). While there is a wealth of information on the mt genomes for a range of animals other than helminths, there is a paucity of knowledge for parasitic nematodes of socio-economic importance. For example, the orders Strongylida, Ascaridida, Spirurida and Rhabditida contain many species infecting animals and/or humans (Anderson, 2000), but until recently, only a small number of complete mt genome sequences was available for species from these orders. The purpose of this chapter is: (1) to present background on mt genomics of animals and emphasize the significance of studying mt genomes, (2) to review the state of the knowledge of the mt genomes of parasitic helminths, (3) to provide an account of some of the approaches used for mt genome sequencing and for population genetic studies, (4) to summarize some applications of mt gene markers to study the molecular systematics and population genetics of parasitic nematodes, and based on the review of the literature, (5) to conclude by emphasizing the prospects and opportunities provided by recent studies of the mt genomes of parasitic nematodes.

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2. BACKGROUND ON THE MITOCHONDRIAL GENOMICS OF ANIMALS 2.1. General and Historical Perspective Mitochondria, the center of oxidative phosphorylation, are usually ovoid-shaped organelles, proposed to originate from free-living eubacteria undergoing endosymbiosis (Lang et al., 1999; Saccone et al., 1999; Kita and Takamiya, 2002). Over millions of years, there has been a progressive transfer of genes from primitive eubacteria to the nuclear genome. In most mt genomes of extant animals, there are 12–13 protein genes (cox1–cox3, nad1–nad6, nad4L, cob, atp6 and/or atp8) encoding enzymes of the oxidative phosphorylation, two ribosomal RNA genes (rrnS and rrnL) encoding the RNA components of mt ribosome, and 22 transfer RNA genes (trn) required for the translation of the different mt proteins (Wolstenholme, 1992). In the literature, different abbreviations have been employed for mt genes and their products. For instance, both abbreviations COI (Zhan et al., 2001) and cox1 (Blouin, 2002) have been used for the cytochrome c oxidase subunit I gene, while ND4 (Hoberg et al., 1999) and nad4 (Blouin, 2002) are used to represent the nicotinamide dehydrogenase subunit 4 gene. To maintain consistency throughout this chapter, the abbreviations of individual mt genes follow those recommended by Le et al. (2000a). In addition to the genes, there is also a relatively large non-coding (control, D-loop or AT-rich) region which, for some vertebrates, is known to be responsible for the regulation of replication and transcription (Figure 1). In most species, the genes are located on and transcribed from two strands, whereas in some groups, such as flatworms, the genes are transcribed from one strand (Le et al., 2002a). With the exception of some cnidarians (Bridge et al., 1992), the mt genome is usually circular and compact in structure, varying in size from 13 to 20 kb. Usually, non-coding regions are present between some genes, and there is sometimes limited overlap between genes (Wolstenholme, 1992; Boore, 1999).


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Figure 1 Schematic representation of the mitochondrial genome of Ascaris suum, as an example to show the organization of 12 protein-coding genes (atp6, cob, cox1–3, nad1–6 and 4L), 2 ribosomal RNA genes (rrnS and rrnL), 22 transfer RNA genes (one-letter code for transferred amino acid) and non-coding region (AT) (modified from Okimoto et al., 1992).

Since the discovery of the mt genome 41 years ago (Nass and Nass, 1962), research on its structure and function has been progressing steadily with the continuous advent of new technologies. In the 1960s, the metazoan mt genome was demonstrated by electron microscopy to be circular (Hollenberg et al., 1970). In the 1970s, mt genomes of 15–20 kb were reported for various metazoan species (Brown, 1983). In the early 1980s, with the advent of DNA sequencing techniques, the entire sequences of both the

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human and mouse mt genomes were completed (Anderson et al., 1981; Bibb et al., 1981). This work stimulated studies of the mt genomes of other animals, with the number of species being investigated increasing rapidly over the last decade. To date, >250 complete mt genome sequences from metazoan organisms have been determined and deposited in current gene databases (e.g. DDJB, EMBL and GenBank). Relevant information (including accession numbers, species names and related publications) is accessible via the website http://www.ncbi.nlm.nih.gov/PMGifs/ Genome/mztax-short.html. This information provides a valuable resource for studying many aspects, such as mt genome arrangements, genome replication and transcription, protein coding genes and genetic codes, transfer and ribosomal RNA genes, and noncoding regions. The following sections review relevant aspects relating to these areas. 2.2. Structural and Functional Aspects 2.2.1.

Gene Arrangement

Usually, mt gene arrangements are relatively conserved within major taxonomic groups, but variable among them (Boore, 1999). In vertebrates, the arrangement of all 37 genes is relatively conserved (Anderson et al., 1981; Roe et al., 1985; Miya et al., 2001), although minor rearrangements have been detected in some ray-finned fishes (Inoue et al., 2001; Miya et al., 2001), lamprey (Lee and Kocher, 1995), amphibians (Sumida et al., 2001), reptiles (Kumazawa and Nishida, 1999), birds (Desjardins and Morais, 1990; Mindell et al., 1999; Haddrath and Barker, 2001) and marsupials (Paä¨bo et al., 1991; Janke et al., 1994, 1997, 2002; Phillips et al., 2001). These rearrangements relate mainly to translocations of a small number of trn genes and/or a translocation of the protein gene nad6. Compared with vertebrates, gene arrangements in invertebrates are more variable, both within and among phyla, including the Arthropoda, Annelida, Mollusca, Platyhelminthes and Nematoda. For example, mt genome organisations for members of Arthropoda


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(e.g. insects, crustaceans and chelicerates) were originally reported to be very similar. Only the positions of a few trn genes in the vicinity of the control region and in the gene cluster trnA-trnR-trnN-trnS1-trnEtrnF varied (Boore, 1999). However, a recent study has shown that significant rearrangements of the protein and the rrn genes occur for three orders of hemipteroid insects (Shao et al., 2001). Novel gene arrangements have also been described for ticks (Black and Roehrdanz, 1998; Campbell and Barker, 1998, 1999), the hermit crab (Hickerson and Cunningham, 2000) and millipedes (Lavrov et al., 2002). In contrast to the large number of genome sequences representing members of Arthropoda, only two complete mt genome sequences have been determined for the Annelida. They include the common earthworm, Lumbricus terrestris (see Boore and Brown, 1995) and Dumeril’s clam worm, Platynereis dumerilii (see Boore, 2001). Comparison of the gene order between these two genomes revealed 7 trn gene translocations, but the annelid mt genome organisation differs significantly, in that the atp6 and atp8 are not adjacent to one another, whereas in chordates, echinoderms and most arthropods these 2 genes are usually adjacent (Boore, 1999). The first mollusc for which the complete mt genome arrangement was determined was the blue mussel, Mytilus edulis (see Hoffmann et al., 1992). The arrangement in this species is remarkably different from any other mt genome reported previously. Following this report, complete mollusc mt genomes were sequenced for 7 species: Albinaria coerulea (door snail) (Hatzoglou et al., 1995), Cepaea nemoralis (banded wood snail) (Terrett et al., 1996), Crassostrea gigas (pacific oyster), Katharina tunicata (black chiton) (Boore and Brown, 1994), Loligo bleekeri (bleeker’s squid) (Sasuga et al., 1999), Pupa strigosa (opisthobranch gastropod) (Kurabayashi and Ueshima, 2000) and Venerupis philippinarum (Japanese little-neck clam) (Okazaki and Ueshima, unpublished). Interestingly, the gene arrangements of these molluscan mt genomes are highly divergent from one another, with only a few shared gene boundaries. Also, their gene contents are variable. Of the 8 molluscan species investigated to date, three of them lack the atp8, a gene usually

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present in metazoans. Also, both K. tunicata and M. edulis have 1–2 additional trn genes in their mt genome. Recently, complete or near complete mt genome sequences have been determined for 11 species of flatworm (Platyhelminthes) (Le et al., 2000a, 2001a,b, 2002a,b; von Nickisch-Rosenegk et al., 2001). Like some molluscs, all platyhelminth mt genomes sequenced to date lack the atp8 gene. The mt genome organisation within this phylum is relatively conserved, with a few trn transpositions, except for Schistosoma mansoni, which differs significantly from other members within the phylum (Le et al., 2000c) in that there are 3 protein gene and gene block translocations, and multiple transpositions of trn genes or non-coding regions (Le et al., 2000c). To date, only a small number of mt genomes have been sequenced for the Nematoda, and these have some unique features compared with other organisms (an account of the mt genomes of nematodes is given in Section 3). The description of mt gene rearrangements has stimulated further study of the mechanisms for their occurrence. Gene rearrangements were usually found to occur among genes adjacent to trn genes or noncoding regions and, thus, were proposed to be caused by tandem duplication of genes due to replication slippage, followed by gene deletions (Moritz and Brown 1986, 1987; Paä¨bo et al., 1991). Other proposals suggest that gene rearrangements are initiated by errors in replication of the ‘light strand’ of mt DNA (Macey et al., 1997, 1998). While there has been almost a consensus view that animal mt DNA does not recombine (Moritz and Brown, 1987), some studies have provided evidence to the contrary (e.g. Holt et al., 1997; Lunt and Hyman, 1997; Kajander et al., 2000), which would better explain the translocation and inversion of large fragments compared with the ‘duplication and deletion theory’ (Dowton and Campbell, 2001; Dowton et al., 2002; Lavrov et al., 2002).

2.2.2.

Replication and Transcription

Although the mechanisms or processes of mt gene rearrangement are still unclear, the replication and transcriptional processes in mt


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genomes, which are usually asymmetrical and unidirectional, are relatively well understood for vertebrates (Clayton, 1982, 1984, 1991, 1992). The mt genomes of vertebrates usually have two separate and distinct origins of replication which are designated as OH (the origin of H-strand synthesis) and OL (the origin of L-strand synthesis) (Clayton, 1992). OH is located within the displacement loop (D-loop) region of the genome, while OL is distant from OH and is usually surrounded by a cluster of several trn genes (Clayton, 1982). Promoters for H-strand transcription (HSP) and L-strand transcription (LSP) are also positioned in the D-loop region (Clayton, 1984). Mitochondrial DNA replication and transcriptional processes are linked, because the same transcription event serves for both replication priming and transcription initiation (Clayton, 1991). RNA transcripts derived from LSP serve as primers for H-strand DNA replication. The transitions from RNA to DNA synthesis at OH occur in three conserved sequence elements (known as conserved sequence blocks, CSB1–CSB3) in the D-loop region (Clayton, 1991; Shadel and Clayton, 1997), with the aid of a (mt RNA processing) ribonucleoprotein, called RNase MRP (for mt RNA processing) (Lee and Clayton, 1998). After transition, synthesis of the daughter H-strand continues unidirectionally. When H-strand synthesis is twothirds complete, the OL is exposed as a single-strand which is then recognised by a mt DNA primase, and RNA priming and DNA synthesis of the L-strand are thus initiated (Clayton, 1991). In the final stage of replication, the two newly synthesised (daughter DNA) molecules are separated, RNA primers at both origins are removed, the remaining gaps in the molecule are filled and ligated, and the super-helical turns are formed into a covalently closed circular mt DNA molecules (Clayton, 1991). Each of the two promoters (HSP and LSP) in the D-loop region responsible for the transcription of one of the two strands of the circular mt DNA region (H-strand and L-strand, respectively) contain two functional elements, namely a short sequence at the transcription start sites necessary for transcription initiation, and an upstream region for the binding of the mt transcription

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factor (mtTFA) which is encoded by a nuclear gene (Chang and Clayton, 1984; Fisher and Clayton, 1985). Human mtTFA contains 204 amino acids, and has the capacity to unwind and bend DNA (Clayton, 1991). This mtTFA is required for the efficient initiation of mt DNA transcription in the presence of a mt RNA polymerase. Mitochondrial RNA synthesis has a unique feature in that single, large poly-cistronic primary transcripts are produced and subsequently cleaved into individual products (Gilham, 1994; Reichert et al., 1998). Cleavage occurs at the positions of the trn genes, since their secondary structure can serve as a signal for cleavage by processing enzymes (Ojala et al., 1981). However, in some circumstances, at processing sites where no trn sequences are present, a stem-loop structure occurring at the border between protein-coding and/or rrn can replace the trn as a processing signal (e.g. Bibb et al., 1981). Although detailed information on these molecular events has been determined for mammals (reviewed by Shadel and Clayton, 1997), the conserved structure of the D-loop region in vertebrates (Wolstenholme, 1992) and the organisation of invertebrate mt genomes (Boore, 1999) suggest that similar mechanisms are applicable to invertebrates. However, to date there is no precise information of these processes for any parasitic helminth.

2.2.3.

Protein Genes and Genetic Code

Of the genes replicated and transcribed in the mt genome, 13 are protein genes, including three subunits of cytochrome c oxidase complex (cox); cytochrome b (cob); two subunits of adenosine triphosphatase, atp6 and atp8; and the nicotinamide dehydrogenase (nad) complex consisting of nad1–nad6 and nad4L subunits. Interestingly, the atp8 gene is absent from the mt genomes of three species of mollusc (Crassostrea gigas, Mytilus edulis and Venerupis philippinarum), the ascidian Halocynthia roretzi, the secernentean nematodes, and all flatworms sequenced to date


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(Hoffmann et al., 1992; Okimoto et al., 1992; Yokobori et al., 1999; Le et al., 2002a). Since the publication of the human mt genome sequence (Anderson et al., 1981), the identification of the protein genes in mt genomes of other animal species has been based largely on amino acid sequence and hydropathic profile similarities to the corresponding genes of human mt DNA (e.g. Bibb et al., 1981; Desjardins and Morais, 1990). While gene content and size are relatively invariable for many animal mt genomes, the genetic code is distinctly different from the standard or nuclear genetic code (Osawa et al., 1992; Wolstenholme, 1992). While ATG is a universal initiation codon for methionine in the nuclear genome of animals, ATT, ATA, ATC, GTG, GTT and TTG are also used as initiation codons in the mt genome (Wolstenholme, 1992). Although GTT is sometimes employed in parasitic helminths (Blair et al., 1999), TTG is commonly used as an initiation codon in nematodes (Keddie et al., 1998). Also, ATAA has been suggested as the translation initiation codon for the Drosophila melanogaster cox1 gene (de Bruijn, 1983). Usually, TAA or TAG are used as translation termination codons in animal mt genomes, except for vertebrates where either AGA or AGG is employed. Some of the protein genes have an abbreviated termination codon, such as T and/or TA (Wolstenholme, 1992), and it has been demonstrated that post-transcriptional polyadenylation completes these codons by adding additional A’s to the abbreviated stop codons (Ojala et al., 1981). In addition to these unusual initiation and termination codons, a modified genetic code is employed for mt protein genes. For example, while TGA is used as a stop codon in the nuclear genome, it encodes tryptophan in the mt genome. Both AGA and AGG function as termination signals in the mt genome of vertebrates, but code for serine in invertebrates (Wolstenholme, 1992) and for glycine in the H. roretzi (see Yokobori et al., 1999). For most metazoans, ATA encodes methionine, but in echinoderms, hemichordates, trematodes and cestodes, it encodes isoleucine. AAA specifies lysine in the universal genetic code, but in many mt genomes of metazoans, it encodes asparagine. The possible

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mechanisms for such a ‘modification’ of the genetic code during metazoan evolution include codon capture, and have been described in detail in a previous review (Osawa et al., 1992). 2.2.4.

Transfer and Ribosomal RNA Genes

In the animal mt genome, 64 genetic codes relate to the anticodons of 22 transfer RNA (trn) genes. These trn genes are distributed throughout the mt genome and are sufficient to encode the 12 or 13 protein-coding genes for mt ribosomes using ‘relaxed wobble rules’ (Crick, 1966), in which only one nucleotide at the first position in the anticodon pairs with any of the four possible nucleotides at the third position of the codon. Currently, the exceptions to this are for the mt genomes of cnidarians, the ascidian, H. roretzi and the mollusc, M. edulis. In the cnidarian mt genome, there are only 2 trn genes which recognise tryptophan and methionin-specifying codons, and, thus, it has been suggested that the remaining trn genes are transcribed and imported from the nuclear genome (Wolstenholme, 1992). In contrast, in both the mt genomes of M. edulis and H. roretzi, there is an additional trn gene for glycine (Hoffmann et al., 1992; Yokobori et al., 1999). Most mt trn genes take on a ‘clover-leaf’ like secondary structure (Figure 2). However, compared with their nuclear ‘counterparts’, mt trn genes show some unusual features. For instance, the trnS (AGN) gene of all mt genomes lacks a DHU arm (Figure 2). Also, other mt trn genes of some species lack either the DHU or T C arm (Figure 2) (Okimoto et al., 1992; Yamazaki et al., 1997; Lavrov et al., 2000; Masta, 2000; Noguchi et al., 2000). For some species of invertebrate, both arms are shortened, suggesting less constraint on the tertiary interactions between DHU and T C loops (Campbell and Barker, 1999). In addition, the 30 CAA tail sequence, possessed by all trn genes, is not encoded, but post-transcriptionally added to the trn (Hsuchen et al., 1983). Moreover, there can be overlaps between trn genes and/or other genes, and sometimes mismatches are present in these ‘overlapped amino acid acceptor arms’. Since a well-matched acceptor stem is central to trn processing and


Figure 2 Secondary structure of mitochondrial (mt) transfer RNA (trn): a; mt trn with TV-replacement loop. b; standard mt trn genes. c; mt trn with DHU-replacement loop (taken from Lavrov and Brown, 2001).

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recognition by aminoacyl-tRNA synthetase (Dirheimer et al., 1995; Martin, 1995; McClain, 1995), the finding of unusual mismatches in amino acceptor stems stimulated studies which indicated that RNA editing corrects any mismatch and completes the transcript from the trn gene (e.g. Yokobori and Paä¨bo, 1995; Bo¨rner et al., 1997; Lavrov et al., 2000). In addition to the 22 trn genes, two rrn genes (rrnS and rrnL) are present in all metazoan mt genomes, based on sequence similarity to the 16S and 23S rrn genes of Escherichia coli (see Bibb et al., 1981; Clary and Wolstenholme, 1985; Noller et al., 1986). These two genes encode the RNA components of the small and large subunits of mt ribosomes (Noller et al., 1986; Noller, 1991). In vertebrates, and most arthropods (Wolstenholme, 1992), platyhelminths (Le et al., 2002a) and annelids (Boore and Brown, 1995; Boore, 2001), these two genes are usually in close vicinity to one another (often only separated by a single trn gene), while in most molluscs (Kurabayashi and Ueshima, 2000; Grande et al., 2002), echinoderms (Scouras and Smith, 2001) and secernentean nematodes (Okimoto et al., 1992; Keddie et al., 1998), they are separated by protein coding genes. Interestingly, although the mt rrnS and rrnL genes of metazoans are smaller than the corresponding genes of E. coli (see Table VI, Wolstenholme, 1992), both genes can be folded into secondary structures which resemble the universal core structures representing this bacterium (Gutell et al., 1985; Gutell and Fox, 1988; Wolstenholme, 1992). The primary sequences of these core structures contain nucleotides responsible for subunit interactions and also for key interactions with trn genes (Noller, 1991, 1993; Wolstenholme, 1992).

2.2.5.

Non-coding Regions

In addition to the protein and rrn genes described above, the mt genomes in most animal species studied thus far usually contain one main non-coding region which is rich in tandem-repetitive sequences (Wolstenholme, 1992). As this region is known to be


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responsible for the regulation of transcription and of DNA replication in some species (e.g. Clayton, 1982, 1991), it has been named the ‘control’ region. Functionally, within the control region of vertebrate mt genomes, a newly synthesised H-strand binds to the parental L-strand and displaces the parental H-strand to form the D-loop region. Although the DNA sequence of the control region is variable among species, there are three conserved sequence blocks (CSB1–CSB3) which are close to the H-strand origin of the D-loop. These conserved sequence blocks are recognised to represent the binding sites of the factors involved in the initiation of mt transcription (Clayton, 1991). The control region of invertebrate mt genomes is rich in adenine and thymine (and is thus called the ‘AT-rich region’) and varies in length among different taxa (e.g. Wolstenholme, 1992; Zhang and Hewitt, 1997). Usually, the primary DNA sequence of this region has little similarity among related species, but a relatively conserved stemand-loop structure can be identified (Zhang and Hewitt, 1997), which is considered to contain the regulatory element serving as binding sites for proteins involved in DNA-protein interaction(s) with the cell nucleus or the regulation of replication and transcription in mt DNA (Gilham, 1994; Zhang et al., 1995; Boore, 1999; Loguercio-Polosa et al., 1999).

2.3. Significance of Studying Mitochondrial Genomes Investigating mt genome structure has significant implications for areas, including mt genome evolution, diseases and syndromes linked to mutations in mt genes, and population genetics and systematics.

2.3.1. Genetic and Genomic Studies As indicated previously, the mt sequences can display a number of unique features, which have been utilised for studying fundamental

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aspects of mt genetics and genomics. For example, the animal mt genome is small and independent from, but has a close relationship with the nuclear genome (Boore, 1999). The organization and mode of gene expression relating to mt genomes of animals are distinct from those of fungi and plants (Lang et al., 1999), which provides opportunities for investigating the evolution of the mt genome and the origin of eukaryotes (Lang et al., 1999). In addition, animal mt genomes have other unique features, such as a relatively constant gene content, proposed maternal inheritance, rare recombination, rearrangement of gene order, variation in genetic codes, base composition and secondary structures of trn genes. All of these features have been employed for studying evolutionary genomics in some metazoan groups (Saccone et al., 1999, 2000, 2002), the mechanisms of gene rearrangements (Macey et al., 1997; Dowton et al., 2002), RNA editing (Yokobori and Paä¨bo, 1995; Bo¨rner et al., 1997; Lavrov et al., 2000), and the evolutionary history of extant species, including humans (Ingman et al., 2000). Also, the study of mutations in mt DNA can provide insights into some disease processes.

2.3.2.

Mutations – Disease, Apoptosis and Ageing

Mutations in mt DNA are an important cause of genetic diseases. For example, in humans, such mutations can relate to a variety of disorders with varying clinical expression, such as mt myopathy, sideroblastic anaemia, thrombocytopenia, neutropenia and endocrine pancreatic dysfunction (Larsson and Clayton, 1995; Chimery and Turnbull, 2001; Nardin and Johns, 2001). Mitochondrial dysfunction is known to mainly affect tissues with a high energy demand, such as skeletal muscle, heart, kidney and endocrine systems (Wallace et al., 1995; Nariaux, 2000; Nardin and Johns, 2001). High mt DNA turnover and a poor DNA repair system compared with the cell nucleus turnover have also been proposed to relate to cancer (Hochhauser, 2000; Penta et al., 2001). For some cancers, it has been demonstrated that when the rate of somatic mutations increases, the


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expression of mt DNA-encoded subunits of the mt electron respiratory chain is elevated (Hochhauser, 2000; Penta et al., 2001). In addition, mitochondria are found to play a key role in controlling life and death of the cell, and thus represent a central control point of apoptosis (Desagher and Martinou, 2000; Wang, 2001). While the effects of mutations have not yet been studied in any detail for invertebrates, such as parasitic helminths, it is likely that such mutations will also affect their health and longevity.

2.3.3. Sequences and Gene Order as Genetic Markers for Molecular Systematic Studies The high rates of mutations in mt genes and the differing rates among these genes in the same genome make them useful genetic markers for molecular systematic and population genetic studies, including those of parasitic helminths (Avise et al., 1987; Avise, 1991, 1994; Moritz, 1994; Simon et al., 1994; Hillis et al., 1996; Avise, 1998; Blouin, 1998; Rand and Kann, 1998; Avise and Walker, 1999; Le et al., 2000a). A detailed account of these aspects is given in Section 4. Interestingly, although mt DNA sequences evolve rapidly, the genome organization within a particular taxonomic group appears to be relatively stable over long periods of evolutionary time (Boore, 1999). As a large number of potential gene rearrangements in a mt genome is possible, it is unlikely that the same gene order arises independently (Boore and Brown, 1998; Boore, 1999). Therefore, a shared gene arrangement is likely to relate to a common ancestry (Boore and Brown, 1998; Boore, 1999). While gene order has been used for molecular phylogenetic studies of deep branch taxa, including echinoderms and arthropods, suggesting that gene arrangement is a useful marker for phylogenetic studies (Sankoff et al., 1992; Smith et al., 1993; Boore et al., 1995; Staton et al., 1997; Boore and Brown, 1998; Boore et al., 1998; Rokas and Holland, 2000), this approach had not been applied to study the phylogenetic relationships of nematodes.

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In conclusion, Section 2 presented some relevant information on structural and functional aspects of animal mt genomes. However, far less is known about the mt genomes of parasitic helminths. In the following section, current information of mt genomes of parasitic helminths is reviewed. 3. THE MITOCHONDRIAL GENOMES OF PARASITIC HELMINTHS Parasitic helminths belong to the Metazoa, and include the Platyhelminthes (i.e. flatworms, such as trematodes and cestodes) and the Nematoda. Within these groups are some very important animal and human pathogens. As indicated in Sections 1 and 2, studying their mt genomes has important implications for elucidating their systematics and population genetics. However, until 1999, little was known about the mt genomes of parasitic helminths.

3.1. Brief Account of Mitochondrial Genomes of Flatworms It was not until recently that mt genome sequencing of parasitic flatworms made substantial progress (reviewed by Le et al., 2000b, 2002a). From this effort, complete or near-complete mt genomes are now available for 11 species of flatworm (reviewed by Le et al., 2002a). Mitochondrial genomes of flatworms are usually small, and their coding regions are 13–14 kb in size. They contain 36 genes, like secernentean nematodes and some molluscs, and lack the atp8 gene. As distinct from the nematode mt genomes published, most platyhelminth mt genomes have an AT content of 12 Ts) which have not yet been described for any other animal mt genome and whose function is presently unclear. However, the most striking finding is the uniqueness of the mt gene arrangement of S. stercoralis compared with all other nematodes studied to date (Hu et al., 2003a); only a limited number of gene boundaries are shared among other secernentean nematodes. Evidence of a high degree of variation in gene arrangement opens up exciting prospects for discovering a diversity of gene arrangements for other parasitic nematodes. The diversity in the mt genome structure and organization among some species of the nematode examined (e.g. S. stercoralis and C. elegans) indicates that mt genes in this group undergo relatively


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frequent rearrangement compared with other organismal groups (such as vertebrates and many arthropods; see Boore, 1999), which should have implications for studying the mechanisms of mt gene rearrangements and mt genome evolution. While some nematode lineages (e.g. Rhabditida) reveal evidence of an increased rate of rearrangement, little is known about the mechanisms leading to these rearrangements, although there are different theories for other invertebrate and vertebrate groups (e.g. Macey et al., 1997, 1998; Kajander et al., 2000; Dowton et al., 2002; Lavrov et al., 2002). Two main models have been proposed to explain rearrangements in animal mt genomes, namely the ‘duplication/random loss’ model (Macey et al., 1997, 1998; Lavrov et al., 2002) (referred to here as Model 1), and the ‘recombination model’ (Poulton et al., 1993; Holt et al., 1997; Kajander et al., 2000; Dowton and Campbell, 2001) (designated as Model 2). Model 1 was originally employed to explain the rearrangements in some vertebrate mt genomes, the mechanisms of which were interpreted to relate to tandem duplication of genes, resulting from errors in light-strand replication (Macey et al., 1997) or strand slippage and mispairing (Moritz and Brown, 1986, 1987; Levison and Gutman, 1987; Madsen et al., 1993), followed by random deletion of genes (Moritz and Brown, 1986, 1987; Moritz et al., 1987; Boore and Brown, 1998; Boore, 2000). However, a more recent study of mt genomes of two species of millipede (Lavrov et al., 2002) suggests that deletions are non-random, which could have significant evolutionary implications. Model 2, originally proposed by Rand and Harrison (1989), was used to explain the presence of precise, tandemly-repeated mt sequences. This model appears to be supported by recent studies inferring that mt recombination does occur in some animals (Lunt and Hyman, 1997; Kajander et al., 2000; Ladoukakis and Zouros, 2001a,b; Hoarau et al., 2002), being in contrast to earlier reports (e.g. Clayton et al., 1974; Hayashi et al., 1985; Moritz and Brown, 1987). Recombination in mt DNA is also supported by recent investigations into humans (Kajander et al., 2000), other vertebrates such as the flat-fish Platichthys flesus (see Hoarau et al., 2002), and some invertebrates including the root-knot nematode Meloidogyne javanica (see Lunt and Hyman, 1997;

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Ladoukakis and Zouros, 2001a). Although Model 1 may explain the translocation of small fragments, it does not necessarily explain the translocation of large fragments (gene blocks) and/or inversions, which may be explained by Model 2. Given these controversies, the diversity in mt genome organization among some nematodes may provide an opportunity for testing these theories. Elucidating the mechanisms of mt gene rearrangements may also have important implications for investigating the evolutionary relationships of nematodes. Although animal mt genes mutate at a higher rate than nuclear genes, rearrangements seem to occur less frequently, whereby the mt genome remains relatively stable for long periods of evolutionary time, and thus contains the ‘message of common ancestry’ (Boore and Brown, 1998; Boore, 1999). Also, mt gene order is considered to be ‘selectively neutral’, and the large number of potential arrangement makes convergence less likely (Boore and Brown, 1998; Dowton et al., 2002). Furthermore, in spite of being exposed to long-term evolutionary processes (800 million years), the gene content of animal mt genomes has undergone relatively little change (Saccone et al., 2002), making (from a phylogenetic perspective) the determination of homologous characters easier. These aspects suggest that gene order data may be more useful for higher-level phylogenetic studies (among orders, classes or phyla) than morphology and/or nucleotide sequence data for individual genes, since morphological similarities may relate to convergent evolutionary change (Hillis et al., 1996) whereas nucleotide sequences may be saturated with change (reviewed by Boore and Brown, 1998; Dowton et al., 2002). Recently, mt gene order data sets have been employed for addressing phylogenetic questions in echinoderms (Smith et al., 1993; Arndt and Smith, 1998; Scouras and Smith, 2001), arthropods (Boore et al., 1995, 1998; Morrison et al., 2002; Roehrdanz et al., 2002), platyhelminths (von Nickisch-Rosenegk et al., 2001) and vertebrates (Kumazawa and Nishida, 1995; Macey et al., 2000). However, there is controversy as to the reliability of this approach when applied to a wide range of animal species (e.g. Curole and Kocher, 1999; Le et al., 2000c), because of the presence of parallel evolution (Mindell et al.,


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1998) and the occasional rapid genome rearrangement within a particular genus (e.g. Schistosoma; see Le et al., 2000c). This information suggests that the limitation(s) of using this approach may relate to convergence and gene rearrangement mechanism bias (Dowton et al., 2002), which may be overcome by broadening taxonomic sampling (Mindell et al., 1998; von Nickisch-Rosenegk et al., 2001; Dowton et al., 2002). The controversy surrounding the reliability of mt gene order for phylogenetic analysis is a stimulus to assess the applicability of complete mt genome data sets for nematodes. Clearly, this area deserves future study. While gene rearrangement data have not yet been used for investigating the phylogeny of nematodes, informative concatenated nucleotide and amino acid sequence data sets (derived from full mt genome sequences) have been demonstrated to be useful for studying their higher-order phylogeny (Hu et al., 2003a), which is of significance because it provides the prospect of testing current evolutionary theories relating to parasitic nematodes and their mt genomes. Nematodes are considered to belong to an ancient, diverse group of metazoans, ranging from free-living species in soil or water to those parasitic to plants, invertebrates and/or vertebrates. Traditionally, the phylogenetic analysis of the phylum Nematoda was based on using morphological character sets and/or ecological traits (cf. Dorris et al., 1999). However, such studies have been hampered by the paucity of phylogenetically-informative morphological characters, limited fossil record and an unstable classification at ordinal and higher levels (Inglis, 1983; Maggenti, 1983; Bird, 1986). It is thus not surprising that controversies have arisen from various studies (e.g. Maggenti, 1983; Adamson, 1987; Lorenzen, 1994; Malakhov, 1994). Clearly, molecular approaches provide useful complementary tools to address such controversies. For instance, nuclear ribosomal RNA sequence data have been used to propose an evolutionary framework for the Nematoda (Blaxter et al., 1998), which enabled a comparison to be made among species with differing morphology and ecology, across different orders and classes. However, some findings disagreed with those from analyses based on morphological data sets (e.g. Inglis, 1983; Adamson, 1987;

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Malakhov, 1994). For example, the division of Secernentea and Adenophorea was not supported by analyses of the molecular data (Blaxter et al., 1998; Dorris et al., 1999), which divided nematodes into five clades and grouped some free-living nematodes with parasitic ones within each of the clades. Independent molecular data sets, such as those derived from complete mt genome sequences, could be employed to address this controversy and to test current hypotheses regarding nematode phylogeny. In addition to their applicability to systematic studies, complete mt genome sequences also provide a rich source of genetic markers for investigating the population genetics of parasitic nematodes, because of their high mutation rates and (assumed) maternal inheritance. Indeed, a range of studies (using PCR-based RFLP, mutation scanning and/or sequencing approaches) have proven their applicability to population studies (Anderson and Jaenike, 1997; Blouin, 1998; Gasser and Newton, 2000). To date, a number of gene regions (including cox1, nad4, rrnL and rrnS) have been used for such studies (Sections 4.4–4.5). The approach of mutation scanning proved particularly useful for detecting population substructuring and (combined with appropriate analyses) for estimating gene flow. For instance, SSCP analysis of the cox1 gene identified population substructuring within each of three hookworm species An. caninum, An. duodenale and N. americanus. While the substructuring within each An. caninum and An. duodenale suggests the existence of different ‘strains’ with differing biological characteristics (such as infectivity to humans, in the case of the canine hookworm), particularly interesting finding was the substantial genetic variation detected between N. americanus from China and from Togo, raising questions as to species status of the parasite (Hu et al., 2003c). Future work should focus on addressing this and similar questions by studying relatively large sample sizes of N. americanus from different continents, including South and Central America, Africa and Asia. In another recent study (Hu et al., 2002d), the population genetic structure of the bovine lungworm, D. viviparus, was examined in Sweden. The analyses revealed that both the mt DNA diversity (in the cox1) within populations and the gene flow among populations of


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D. viviparus were low, which was more similar to findings for some parasitic nematodes of plants and of insects (Hugall et al., 1994; Blouin et al., 1999) than to species from gastrointestinal trichostrongylid nematodes of domesticated ruminants, which usually have relatively high levels of diversity and gene flow (Blouin et al., 1992, 1995; Blouin, 2002). While this difference between parasites within the same order (Strongylida) was interpreted to relate mainly to differences in host movement and parasite biology, population sizes and transmission patterns, thus being of epidemiological relevance, the precise reasons are still unknown. In spite of some progress over the years, the population genetic structures for the vast majority of nematodes are still unknown, which provides excellent scope for future investigations using mt markers. Another important application of mt gene markers relates to the detection of cryptic species (e.g. Stevenson et al., 1995; Blouin et al., 1997; Anderson et al., 1998). Identifying such ‘hidden’ species is not only important for fundamental studies (population biology, ecology and evolution) but also has practical implications for epidemiology and disease control. As explained recently by Blouin (2002), mt DNA gene markers can be more useful for differentiating cryptic species than nuclear ribosomal DNA markers. With more complete mt genome sequences becoming available, mt markers could be widely applied to species–specific differentiation in nematodes. For example, Hypodontus macropi is a hookworm-like nematode which inhabits the ileum, caecum or colon of a number of species of kangaroo and wallaby (Beveridge, 1979). However, it was proposed to represent a species complex based on a review of the host range and distribution of the parasite (Beveridge, 1979). Allozyme electrophoresis was therefore employed to assess the level of genetic difference among samples from different host species (Chilton et al., 1992). Fixed genetic differences of 15–50% were found among samples studied, including those from hosts collected in sympatry, which provided evidence to support the hypothesis. Subsequent studies using nuclear ITS-2 ribosomal DNA sequence data also provided additional molecular evidence to support the cryptic species hypothesis (Chilton et al., 1995; Gasser et al., 2001), mt gene markers have not yet been used for studying H. macropi

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populations. For another example, there is allozyme electrophoretic evidence for hybridisation between Paramacropostrongylus iugalis and P.typicus(Nematoda:Strongyloidea)inareasofeasternAustraliawhere their hosts occur in sympatry (Chilton et al., 1997). These two nematode species occur in the stomachs of eastern (Macropus giganteus) and western (M. fuliginosus) grey kangaroos, respectively (Chilton et al., 1993). These hybrid individuals did not represent the F1 generation, suggesting there was some gene flow between the two species of nematode (Chilton et al., 1997). These examples indicate the exciting prospects of using mt gene markers to elucidate the genetic make-up of these parasites and to address questions regarding their speciation. While perhaps not immediately obvious, comparative mt genome analysis also has implications for functional genomic studies of parasitic nematodes. For example, the similarities in mt genome organization and the phylogenetic relationships proposed for some species of nematode based on mt genome data sets indicate that some strongylid nematodes, such as the human hookworms, An. duodenale and N. americanus, are relatively closely related to C. elegans (see Hu et al., 2002b, 2003a), which is accordance with previous analyses of nuclear ribosomal DNA sequence and expressed sequence tag (EST) data sets for a range of nematodes from different orders (Blaxter, 1998; Blaxter et al., 1998). Importantly, the free-living nematode, C. elegans, represents one of the best characterised multicellular organisms, and its nuclear genome sequence is known (The C. elegans Sequencing Consortium, 1998). Of its 20,000 genes, 58% appear to be nematode-specific, and key biological processes appear to be conserved among nematodes (Blaxter, 1998), which suggests that C. elegans is a useful functional model for parasitic nematodes (cf. Bu¨rglin et al., 1998; Geary et al., 1999; Gasser and Newton, 2000), in spite of not being a dioecious nematode. Other features, including the nematode’s short life cycle (three days at 25 C), ease of propagation of well-defined lines using a simple bacterial food source (Escherichia coli), relatively small genome size, ability to produce clonal progeny from hermaphrodites and to cross hermaphrodites with males, make C. elegans an attractive system (cf. Bu¨rglin et al., 1998). Also, the advent of techniques, such as gene transformation, RNAi and global


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profiling of gene expression by micro-chip gene array (e.g. Hanazawa et al., 2001; Newton et al., 2002) represent powerful tools for functional genomics studies. Therefore, given that strongylid nematodes are currently considered to be relatively closely related to C. elegans (see Blaxter et al., 1998; Hu et al., 2002b, 2003a), the chance that a gene from a member from this group has a homologue in C. elegans is high (with the exception of genes involved specifically in host-parasite interactions). This statement is supported by unpublished results showing that 60% of genes from subtractive gene libraries (Nisbet et al., personal communication) have a homologue (with a P-score of 0.001) in C. elegans. Thus, until culture systems become available for the effective in vitro propagation and maintenance of life cycles of (at least some) strongylid nematodes, C. elegans represents a useful system for testing the function of genes homologous to theirs (cf. Boag et al., 2003). In conclusion, recent studies have provided new insights into mt genome structures, and the population genetics, taxonomy and evolution of parasitic nematodes of socio-economic importance. Importantly, molecular tools available and advances made provide unique prospects and opportunities for future work on a number of fundamental areas, such as the mechanisms and processes of gene rearrangements, mutation rates, the inheritance of mt genomes and genes, and the systematics and population genetics of nematodes. This also provides a foundation for tackling questions regarding the ecology and epidemiology of parasitic nematodes, thus contributing, in the broader sense, to the diagnosis and control of parasitic diseases. While the main focus of this chapter has been on parasitic nematodes, approaches established and recent progress made could have implications for studying other groups of pathogens, thus contributing more widely to the field of infectious diseases.

ACKNOWLEDGEMENTS MH was the recipient of postgraduate scholarships and the Rhowden White Prize (2002) from the Faculty of Veterinary Science of The

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University of Melbourne, and was granted a Travel Scholarship from the Australian Society for Parasitology (2002). The authors are grateful to a number of colleagues, in particular Xingquan Zhu, Ian Beveridge, Youssef Abs EL-Osta, Johan Ho¨glund and Anton Polderman, for their contributions to some of the primary author’s studies referred to in this review. Funding support to RBG was from various sources, including the Australian Research Council, the Canine Research Foundation and the Australian Animal Health Foundation. REFERENCES Adamson, M.L. (1987). Phylogenetic analysis of the higher classification of the Nematoda. Canadian Journal of Zoology 65, 1478–1482. Albonico, M., Crompton, D.W. and Savioli, L. (1999). Control strategies for human intestinal nematode infections. Advances in Parasitology 42, 277–341. Allan, D. and Johnson, A.W. (1960). A short history of husk. Veterinary Record 72, 42–44. Anderson, R.C. (2000). Nematode Parasites of Vertebrates. Their Development and Transmission. Wallingford: CAB International. Anderson, R.C., Chabaud, A.G. and Willmott, S. (1974). CIH Keys to the Nematode Parasites of Vertebrates. Farnham Royal: Commonwealth Agricultural Bureaux. Anderson, S., Bankier, A.T., Barrell, B.G., de Bruijn, M.H.L., Coulson, A.R., Drouin, J., Eperon, I.C., Nierlich, D.P., Roe, B.A., Sanger, F., Schreier, P.H., Smith, A.J.H., Staden, R. and Young, I.G. (1981). Sequence and organization of the human mitochondrial genome. Nature 290, 457–465. Anderson, T.J.C. (1995). Ascaris infection in humans from North America: molecular evidence for cross-infection. Parasitology 110, 215–219. Anderson, T.J.C. (2001). The dangers of using single locus markers in parasite epidemiology: Ascaris as a case study. Trends in Parasitology 17, 183–188. Anderson, T.J.C., Blouin, M.S. and Beech, R.N. (1998). Population biology of parasitic nematodes: applications of genetic markers. Advances in Parasitology 41, 219–283. Anderson, T.J.C., Romero-Abal, M.E. and Jaenike, J. (1993). Genetic structure and epidemiology of Ascaris populations: patterns of host affiliation in Guatemala. Parasitology 107, 319–334.


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The Cytoskeleton and Motility in Apicomplexan Invasion Ruth E. Fowler1, Gabriele Margos and Graham H. Mitchell* Malaria Laboratory, Peter Gorer Department of Immunobiology, Guy’s, King’s College and St Thomas’s Hospitals’ School of Medicine, New Guy’s House, Guy’s Hospital, London, SE1 9RT, UK Abstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Basic Description of the Apicomplexan Zoite and Organisation of the Cytoskeleton. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. External Proteins Implicated in Motility . . . . . . . . . . . . . . . . . 4. Cytoskeletal and Motor Proteins . . . . . . . . . . . . . . . . . . . . . 4.1. Apicomplexan Actins . . . . . . . . . . . . . . . . . . . . . . . 4.2. Actin-binding Proteins . . . . . . . . . . . . . . . . . . . . . . 4.3. Apicomplexan Myosins . . . . . . . . . . . . . . . . . . . . . . 4.4. Myosin Binding Proteins/Regulatory Proteins . . . . . . . . . 4.5. Inner Membrane Complex Protein 1 (IMC1) and 2 (IMC2). 4.6. Tubulin and Microtubules (mt) . . . . . . . . . . . . . . . . . 4.7. Apicomplexan Tubulin Genes . . . . . . . . . . . . . . . . . . 4.8. -Tubulins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.9. Post-translational Modifications of -tubulin . . . . . . . . . 4.10. Microtubule Associated Motor Proteins . . . . . . . . . . . . 5. Models of Motility . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

214 214 215 219 221 221 224 226 233 235 235 237 238 239 241 242 249 249

1 Present address: Membrane Biology Group, Division of Biomedical Sciences, University of Edinburgh, George Square, Edinburgh EH8 9XD, UK.

*Author for correspondence.



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RUTH E. FOWLER ET AL.

ABSTRACT We consider the cytoskeletal structure, function, and motility of the invasive zoites of the Apicomplexa. This monophyletic group possess a prominent microtubular cytoskeleton, with a very distinct polarity. It is associated with a non-actin based filamentous system, and with a cisternal double membrane assembly beneath the plasma membrane. The origin of the microtubular cytoskeleton is a set of apical rings. Its role in motility is still unclear, but the present knowledge of apicomplexan tubulins’ molecular biology and chemistry is outlined. Actin and accessory proteins are present, and it is apparent that actin polymerisation is tightly controlled in zoites. It does not contribute to the cytoskeleton ordinarily, but is crucial in the acto-myosin linear motor which drives gliding, capping, and invasion, the best understood aspects of zoite motility. Several myosins distinct from the primary linear motor myosin are also found, but not yet well understood functionally. Many of the myosins fall into a class of the superfamily so far seen only in this phylum. The possible relationships of the actin, myosin, cytoskeletal linkage proteins, and external force-transducing adherent proteins are discussed. 1. INTRODUCTION Most studies on the phylum Apicomplexa have been limited to parasites of medical or veterinary importance which fall in the coccidian and piroplasm classes. As a result there are many relatively neglected genera and our review cannot claim to be comprehensive. The most detailed work on motility and invasion has been on the coccidians Toxoplasma, Plasmodium, and Eimeria. From these it is apparent that in generating motility during the host cell invasion process there are features and mechanisms in common, and which are peculiar to the phylum. It is clearly also the case that there is great variation in host-cell invasion events within the phylum: e.g. Theileria does not actively invade, but subverts host cell physiology to gain entry (Shaw et al., 1991; Shaw and Tilney, 1995), Cryptosporidium

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creates and inhabits an extra-cytoplasmic vacuole (Lumb et al., 1988; Beier and Sidorenko, 1991) and some members of the class Gregarinia, parasites of arthropods and annelids, never fully internalise. Here we shall concentrate on what we have learnt from the coccidian parasites, but draw on the other classes for parallels.

2. BASIC DESCRIPTION OF THE APICOMPLEXAN ZOITE AND ORGANISATION OF THE CYTOSKELETON Once an apicomplexan differentiates into a motile/invasive zoite form, the ‘apical complex’ (whence the name of the phylum) of polar rings and secretory organelles gives the cell polarity (Figures 1a and b). The structurally prominent zoite cytoskeleton no doubt functions in maintenance of cell shape and provision of mechanical strength to the plasma membrane, but it also plays an important part in motility and host cell invasion. Consequently in this review we put especial emphasis on structures that contribute to the composition of the cytoskeleton, e.g. the pellicle and its underlying microtubules (mt), polar rings and, in some genera, the conoid. Although in apicomplexan parasites actin does not seem to participate in the formation of the cytoskeleton as such, together with myosin it forms the motor that drives cell motility. Cell shapes of apicomplexan zoites vary from rounded merozoites to elongated sporozoites, related to the number, length and angle of the subpellicular mt which run along the cell. Where there are several subpellicular mt, they form rib-like structures. In some coccidians, e.g. T. gondii, and Besnoita jellisoni, these mt follow a gentle anticlockwise helix. P. falciparum merozoites are unusual in that 2 or occasionally 3 parallel subpellicular mt form a single band along two thirds of the cell (Bannister and Mitchell, 1995). In most coccidia the subpellicular mt terminate in the region of the nucleus in the posterior half of the cell (D’Haese et al., 1977; Nichols and Chiappino, 1987). Basal rings are found in some apicomplexans, and occasionally, as in the piroplasm Theileria for example, the subpellicular mt terminate at this ring (Shaw and Tilney, 1992).

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Figure 1 (a) A generalised body plan of an invasive apicomplexan zoite, to show the organisation of the cellular machinery involved in motility and invasion. The encircled area contains the structures shown more schematically and enlarged in Figure 1b; (b) Two possibilities are shown for the organisation of the main components of the linear motor believed to be responsible for zoite motility in invasion. The myosin may gain purchase on the IMC (left-hand side of diagram), or by linkage to the tails of substrate binding proteins at the plasma membrane (right-hand side of diagram). Abbreviations: IMC inner membrane complex (cisternal membranes, or alveolae); IMPs intra-membranous particles; MTIP myosin A tail interacting protein.


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The subpellicular mt are laterally anchored into an unusual mt organising centre (MTOC) (Russell and Burns, 1984), a polar ring at the zoite apex. Its biochemical make-up is unknown, although there is some evidence that proteins common to other eukaryotic MTOC may be present (Fowler et al., 2001). This ring often surrounds a cylindrical structure composed of delicate tubulin spirals, called a conoid, which is preceded by two pre-conoid rings (McLaren and Paget, 1968; Varghese, 1975; Morzaria et al., 1976; Nichols and Chiappino, 1987; Hu et al., 2002). However in Plasmodium zoites, there are simply three apical polar rings, the subpellicular mt adjoined to the most distal ring (e.g. Bannister and Mitchell, 1995). Together these structures create the prominence through which the apical organelles secrete their contents. In many coccidians there are also two short mt, anchored into the first pre-conoid ring, which run asymmetrically through the centre of the conoid (Porchet-Hennere and Nicolas, 1983). Closely bound together, they are surrounded by a dense matrix material and appear to be connected by bridges to the conoid, the pellicle and the secretory organelles (Nichols and Chiappino, 1987). The subpellicular mt underlie an unusual double membrane structure, which consists of either numerous flattened cisternal vesicles fused together in longitudinal plates, e.g. the inner membrane complex (IMC) of Toxoplasma, Eimeria and Sarcocystis (Dubremetz and Torpier, 1978; Chbouki and Dubremetz, 1985), or a single flattened cisterna, as seen, for example, in Plasmodium merozoites (Bannister and Mitchell, 1995). This double membrane lies under the plasma membrane (together they form the parasite pellicle) lining the entirety of the cell body, with interruptions at the apical prominence, the basal pole and micropores (Chobotar et al., 1975). In the coccidians Toxoplasma, Eimeria and Sarcocystis, there are uniformly sized intramembranous particles (IMPs), arranged in longitudinal rows, within the double membrane, following the plane of the IMC plates, which in turn follow the angle of the subpellicular mt (Dubremetz and Torpier, 1978; Cintra and deSouza, 1985b). In freeze fracture studies these are apparent on the plasma faces of the double membrane, forming double rows which appear to lie over the

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subpellicular mt, and are likely to mark the positions of ‘bridges’ observed between the mt and the membranes. These double rows are interspersed by parallel, equally spaced single IMP rows. Morrissette et al. (1997) demonstrated that these IMPs had a longitudinal repeating pattern which extended the full length of the parasite. They proposed that this implied an additional set(s) of filaments. Recently, this subpellicular network has been observed in deoxycholate-extracted T. gondii tachyzoites (Mann and Beckers, 2001). Composed of 8–10 nm diameter, interwoven filaments, it extends from the polar ring to a smooth cup-like structure at the posterior end of the parasite. It appears to be tightly associated with the subpellicular mt and the cytoplasmic face of the IMC, and Mann and Beckers concluded that the network imparted mechanical stability to the parasite. They also characterised two novel component proteins, TgIMC1 and 2. A homologue of TgIMC1 was found in the P. falciparum database, suggesting that other apicomplexans may have similar structures (Mann and Beckers, 2001). Some previous studies support this proposal: D’Haese et al. (1977) reported a lattice in coccidian pellicles, Yasuda et al. (1988) saw a ‘meshwork’ in T. gondii trophozoites, in gregarines Preston and others reported ill-defined structures lying between the flattened cisternal membranes (Preston et al., 1993a), and 12 nm filaments had been observed lying under the double membrane in gregarine surface folds (Schrevel et al., 1983). Since the IMP lattice extends the full length of the parasite, the reports of apical/anterior concentration of actin in T. gondii tachyzoites (Cintra and De Souza, 1985a; Endo et al., 1988; Yasuda et al., 1988) indicated that filamentous (F-) actin was not part of the lattice structure (Morrissette et al., 1997). The vast majority of actin in T. gondii (Dobrowolski et al., 1997b) was found to be sequestered as monomeric G-actin whereas in Plasmodium merozoites 1/3 of the actin was monomeric (Field et al., 1993). Despite careful conventional ultrastructural studies, actin filaments remain notoriously elusive in coccidians (Bannister and Mitchell, 1995;


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Shaw and Tilney, 1999), although biochemical and fluorescent detection of F-actin has been successful in P. falciparum merozoites (Field et al., 1993; Webb et al., 1996). Recently, actin filaments were detected in Toxoplasma by high resolution low voltage field emission scanning electron microscopy (Schatten and Ris, 2002). Dobrowolski et al. (1997b) suggest the earlier findings reflect the dynamic status of actin in the highly motile T. gondii tachyzoite, with polymerisation occurring only when required for motility/invasion; this is a view supported by studies using actin polymerising drugs (Shaw and Tilney, 1999; Wetzel et al., 2003). Chemically induced polymerisation of T. gondii actin indicates that polymerisation regulates motility, is tightly controlled and occurs at specific sites, primarily the parasite apex, but also directly under the IMC/subpellicular mt complex and at the basal-ring area of the parasite (Shaw and Tilney, 1999; Wetzel et al., 2003). In both Toxoplasma and malaria parasites, myosin too localised to the apex and the pellicle (Webb et al., 1996; Pinder et al., 1998; Hettmann et al., 2000; Margos et al., 2000). P. falciparum Pfmyo-A, P. berghei Pbmyo-A and T. gondii TgM-A represent the motor molecules mainly responsible for invasion/motility (Meissner et al., 2002b), and all three localised to the pellicle, initially concentrated towards the anterior. Subcellular fractionation of Pfmyo-A and Tgmyo-A indicates a tight membrane association, although it needs yet to be resolved whether these motors are associated with the plasma membrane or the cisternal membrane (see Figure 1, and below).

3. EXTERNAL PROTEINS IMPLICATED IN MOTILITY Apicomplexan plasma membranes carry numerous glycolipid-linked surface molecules with intramembranous proteins interspersed, and some of these have been intensively studied. Crucially, some, such as Toxoplasma microneme protein 2 (MIC2), appear to be distributed in the plasma membrane only when required for motility. Initially appearing within the apical organelles, MIC2 is then found at

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the anterior portion of the plasma membrane, subsequently spreading backwards (Carruthers and Sibley, 1997) to be finally deposited in a membranous trail behind the parasite (Hakansson et al., 1999). MIC2 is one of a family of proteins found on apicomplexans related to the vertebrate Thrombospondin (TSP) family, which are adhesive molecules (reviewed by Naitza et al., 1998; Menard, 2001). Curiously, in Plasmodium, proteins with TSP domains have been found only in invasive insect stages (ookinetes/ sporozoites) and not in merozoites. Perhaps this reflects the differences in cell types these forms invade. Disruption of these TSP-domain molecules prevents motility (Sultan et al., 1997; Sibley et al., 1998; Dessens et al., 1999; Kappe et al., 1999; Yuda et al., 1999a,b; Templeton et al., 2000). They are transmembrane proteins, with short cytoplasmic tails and as such are thought to provide purchase on the substrate/host cell so forming at least part of the link between the exterior and the acto-myosin motor. The function of these TSP-domain proteins may be restricted to adhesion to epithelial cells; indeed, the molecule/s that link the malaria merozoite to the surface of the red blood cell remain incompletely identified, and probably require specialised binding properties (Kappe et al., 1999). Other transmembrane proteins may serve as escorters for soluble adhesins which are secreted onto the surface prior to or during invasion (Reiss et al., 2001; Meissner et al., 2002a). The binding of malaria merozoites to the red cell surface is polyphasic, and ‘motility’ of a kind is present in at least two forms prior to the true motor-driven invasion step (Bannister and Mitchell, 2003). Initial, random, and reversible attachment appears to be mediated by the merozoite’s major coat proteins (although this is unproven) and is accompanied by a vigorous shimmying and flexing of the red cell as the parasite folds it about itself (Dvorak et al., 1975). We refer to this shimmying phase as sisterkation (Piron, 1927). Sisterkation is not interrupted (at least, not in P. knowlesi) by antibody directed against the microneme-originating (Bannister et al., 2003), secreted, membrane protein AMA1. Anti-AMA1 antibody, however, blocks invasion before the next discernible invasion step, apical re-orientation,


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is achieved (G. H. Mitchell and colleagues; submitted to Inf. and Imm.). This suggests that making-and-breaking interactions up a concentration gradient formed by AMA1 as it flows out into the membrane (Howell et al., 2001) may be responsible for re-orientating the merozoite. There is some ultrastructural evidence for such a gradient (Thomas et al., 1990). Were the invasion motor to start before re-orientation, then the merozoite would presumably be propelled so that it fruitlessly brought its basal pole into red cell contact. It is also clear that re-orientation is independent of the actin– myosin motor, since it (and the consequent junction formation between merozoite and red cell) occurs in the presence of cytochalasin B (Miller et al., 1979). Plasmodium AMA1 has a homologue described from Toxoplasma (Donahue et al., 2000); the structure, shared molecular motifs, and function of a number of apicomplexan microneme proteins have been discussed recently by Tomley and Soldati (2001).

4. CYTOSKELETAL AND MOTOR PROTEINS Most structures that constitute the cytoskeleton are well documented, but by no means all of the contributing proteins have been identified and biochemically characterised. The requirement of an acto-myosin motor for motility, first suggested by chemical inhibitor studies, was recently confirmed by genetic knock-out experiments (Meissner et al., 2002b). Although the elements composing this motor are now known, not many proteins involved in its regulation have been identified so far. Here, we shall describe known proteins of the cytoskeleton and the motor machinery.

4.1. Apicomplexan Actins The first apicomplexan actin genes to be described were pf-actin I and pf-actin II of P. falciparum (Wesseling et al., 1988a,b). Since then single copy actin genes have been described for C. parvum

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(Kim et al., 1992) and T. gondii (Dobrowolski et al., 1997b). All four proteins are predicted to be 42–44 kDa in size. The C. parvum gene has been mapped to a 1200 kb chromosome and the T. gondii gene to either chromosome 1A or 1B. Apicomplexan actin genes, like those of the ciliated protozoa, are amongst the most divergent known. Sequence homology scores for most eukaryotic actin genes are>87%, whereas the apicomplexan actin genes tend to score in the region of 76–85% (Wesseling et al., 1988a). Even the P. falciparum genes only have a sequence similarity of 79%, the lowest value reported for actins within a single species. pf-actin I appears to be transcribed throughout the parasite lifecycle, whereas the pf-actin II gene only appears to be active in the sexual stages (Wesseling et al., 1988b). This may reflect the differential rRNA expression and processing which is thought to occur in asexual and sexual stages of Plasmodium (Gunderson et al., 1987). Wesseling et al. (1988b) point out that the apicomplexan and the Tetrahymena actin genes appear to be more related to each other than amino acid (aa) sequence homology scores might indicate. For example, the P. falciparum genes have certain aa in nine key positions unique to them, and in a further 8 positions, aa profiles uniquely amongst them and Tetrahymena actin. Three regions of these actins, the N-termini in particular, were shown to have marked differences in their hydrophobicity profiles compared to the high conservation noted for ubiquitous actins (Wesseling et al., 1988b). This suggests that actin evolution is not as well conserved in the apicomplexa and the ciliated protozoans as it is in other eukaryotic lineages. It may also have implications for the mechanisms of F-actin assembly, since the N-terminus is involved in regulating F-actin assembly through interactions with actin binding proteins (ABPs). Apicomplexan actins also have an extra N-terminal aa compared to sequences from vertebrates (Dobrowolski et al., 1997b), although this may be removed if the actin undergoes post-translational modification. Six different isoforms of actin have been ascribed to vertebrate actins: 4 muscle ( actin) and 2 cytoplasmic ( and actins). Amino


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acids found at 20 key positions throughout actin sequences, and the first 18 aa of the N-terminal are used to assign the isotypes (Kim et al., 1992). Invertebrate muscle actins tend to be more like actins. In general, the actins of unicellular organisms do not conform to any of the isotypes, but possess characteristics of them all. This is true of the apicomplexan actins. C. parvum actin is described as being more like the cytoplasmic isoform (Kim et al., 1992). The P. falciparum genes appear to be more actin-like than those of most lower eukaryotes (Wesseling et al., 1988a,b). Wesseling et al., (1989a,b) describe P. falciparum actin II as one of the most muscle-like actins found in single cell organisms. They suggested that it might be required to drive ookinete motility, consistent with E.M. studies showing synthesis of actin filaments during the zygote stage of ookinete development (Sinden, 1983). Actin is also abundant in the P. falciparum merozoite, forming an estimated 0.3% and 0.2% of the total soluble and insoluble protein respectively (Field et al., 1993). Fluorescent labelling showed apical, diffuse cytoplasmic staining and pellicular localisation (Webb et al., 1996; Pinder et al., 1998). Field et al. (1993) found that most of the soluble actin and approximately 20% of the pelleted actin was (mono)ubiquitinated. This modification is usually associated with signalling for various protein sorting, trafficking and gene expression pathways (Pickart, 2001), but it is also a feature of the insect flight muscle actin, arthrin. The authors determined a ratio of approximately 1:2 G- : F-actin in the merozoite. In the insoluble fraction the actin was filamentous. They also concluded that the bulk of the actin in the cytoplasm was either monomeric or in the form of short capped filaments—states which would require ABP sequestration or capping. Dobrowolski et al. (1997b) noted apical concentration of actin only when they stained for globular (G-)actin with DNAse1. Using an antibody against recombinant Tgact-1, immunogold labelled the entirety of the pellicle, as well as the apical and basal poles. However, Western blots of parasite proteins from the detergent soluble and insoluble fractions showed that the vast majority of actin in Toxoplasma tachyzoites was globular. This also suggests sequestration by ABPs.

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4.2. Actin-binding Proteins As already discussed, apicomplexan actin is probably highly dynamic and polymerised when and where it is required. The assembly, disassembly and spatial organisation of actin molecules is regulated by ABPs. Different ABPs initiate/catalyse or inhibit nucleation, cap or uncap filament ends, and link actin filaments to each other or other cell components. Others sever actin filaments and encourage depolymerisation. A few ABPs have been discovered in malaria and Toxoplasma parasites. Tardieux et al. (1998a) describe a P. knowelsi 32/34 kDa actincapping doublet associated with a heat shock protein, HSC70. This complex inhibited the extent of actin polymerisation, and may also have had some nucleation activity—and may, therefore, encourage rapid polymerisation of short filaments when activated. The complex’s capping activity was calcium independent, but was inhibited by phosphatidylinositol 4,5-biphosphate, a second messenger lipid, suggesting that cell signalling cascades are involved in its regulation. A P. falciparum protein has also been identified with strong homology and similar properties to a Dictyostelium discoidum ABP, called coronin (Tardieux et al., 1998b). Coronin is thought to play a key role in re-organising the D. discoidum actin network, speeding up its disassembly and regulating nucleation. Tardieux et al. (1998b) suggest that the coronin homologue might couple regulatory proteins to the apicomplexan acto-myosin motor. Several other actin-binding proteins have been found in P. falciparum parasites, including -actinin-, tropomyosin- and spectrin-like proteins; -actinin has been reported to cross link actin fibres and tropomyosin is a regulator of acto-myosin motors (Forero and Wasserman, 2000). Ghazali et al. (1995) showed that antispectrin antibodies label the apical complex in T. gondii. The authors suggest that because spectrin has a broad range of interactions with other proteins, it is likely to act as a stabilising matrix for the apical complex. In T. gondii, Allen et al. (1997) identified an actin depolymerising factor (ADF), which appeared from its primary structure to be


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a member of a family of small G-actin-binding/F-actin-severing proteins, promoting the rapid turnover of F-actin. In both intra- and extracellular tachyzoites, immunofluorescence imaging of T. gondii ADF showed diffuse cytoplasmic staining, and a more intense rim of staining around the periphery, excluding apical and basal poles. Immunogold electron microscopy also showed labelling of the cytoplasm and the pellicle. ADF, therefore has a similar distribution to actin in tachyzoites. Poupel and colleagues (Poupel et al., 2000) discovered a novel T. gondii ABP, which they named toxofilin. Like T. gondii ADF, this protein bound to G-actin, sequestering monomers, but conversely, also capped F-actin, slowing filament disassembly. Immunofluorescence imaging of toxofilin showed that its distribution varied depending on whether the parasite was intracellular, extracellular and actively motile, or in the process of invading a host cell. Before exit from the host cell, intracellular tachyzoites showed strong, uniform staining at their anterior; however extracellular gliding parasites showed concentration at the anterior end, either as a solid mass, in patches, or in arrow shapes. Basal staining was seen in some of the mobile extracellular parasites, and in all invading tachyzoites. The presence of these ABPs in these parasites is consistent with a highly regulated and dynamic system playing a central role in apicomplexan motility and invasion, in response to appropriate cell signals. The result of overiding these signals and some of the ABP regulation in T. gondii was graphically illustrated in tachyzoites treated with jasplakinolide, a membrane-permeable actin polymerising and F-actin stabilising compound (Shaw and Tilney, 1999; Wetzel et al., 2003). The most dramatic effect was the formation of single, large, membrane-enclosed projections from the apices containing filamentous material. F-actin was observed between the plasma membrane and the outer IMC membrane, forming randomly orientated mats of F-actin as opposed to parallel filaments in nontreated parasites (Shaw and Tilney, 1999; Wetzel et al., 2003). In contrast to host cell actin, all the parasite filaments were approximately parallel with the apical–basal axis, which may be due to the action of ABPs (Shaw and Tilney, 1999). Tachyzoites


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incubated with jasplakinolide became more active, twisting and flexing more rapidly than control parasites (Shaw and Tilney, 1999); low doses of drug increased the speed of motility and reversed its normal direction (Wetzel et al., 2003) confirming F-actin’s role in generating several modes of tachyzoite motility. However these parasites were incapable of invading host cells until the compound was washed away, showing that regulation of actin dynamics was essential for parasite internalisation. Putting all of these results together, it seems likely that actin is rapidly polymerised at specific nucleating centres, i.e. the apex and under the plasma membrane. Possible interpretations are that an apical concentration of F-actin is required to drive the conoid, and/or that short actin filaments are directed between the membranes from the apex, and then depolymerised, either en-route, or when they reach the basal pole.

4.3. Apicomplexan Myosins The importance of myosin for gliding motility and host cell invasion was foreshadowed by the first description of actomyosin in Eimeria by Preston and King (1992), and confirmed by studies showing that the drug butanedione monoxime (BDM), a relatively specific inhibitor of myosin ATPases, reduced host cell invasion by P. falciparum (Pinder et al., 1998), Toxoplasma (Dobrowolski et al., 1997a) and Babesia (Lew et al., 2002).

4.3.1.

Structure

Myosins are molecular motors that move towards the barbed end of actin filaments (with one exception, class VI myosins, which move in the opposite direction). Myosin molecules typically consist of three structural regions, a head or motor domain, a neck domain and a function-specific tail domain. The force is generated by hydrolysis of myosin-bound ATP. Conformational changes in the head domain


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are transferred to the neck region which is thought to act as an amplifier for the power stroke. Phylogenetic tree analysis of the core motor domain divides members of the myosin superfamily into distinct classes. To date 17 classes of myosins have been recognised (reviewed by Mooseker and Cheney, 1995; Mermall et al., 1998; Bahler, 2000; Barylko et al., 2000; Reck-Peterson et al., 2000; Sellers, 2000) (see ‘myosin’ home page http://www.mrc-lmb.cam.ac.uk/ myosin). Apicomplexan myosins have been grouped into a new class XIV because some of them possess quite unusual properties (see below). Many apicomplexan myosins cluster well into this class while for others this assignment is uncertain (Lew et al., 2002). Structurally, the well conserved domains for actin- and ATP-binding within the head region identifies them as members of the myosin superfamily (Heintzelman and Schwartzman, 1997). This classification was supported by biochemical analyses of some of these molecules (Pinder et al., 1998; Heintzelman and Schwartzman, 1999; Hettmann et al., 2000; Herm-Gotz et al., 2002). Myosins have been identified in a number of apicomplexan parasites but for just a few of them (in Toxoplasma, Plasmodium and Babesia, see Table 1) detailed information is available which will be described below. For other apicomplexan myosins short gene sequences are in the public domain but the molecules have yet to be characterised. The first apicomplexan myosins to be described were TgM-A and TgM-B/C (later renamed Tgmyo-A and TgmyoB/C) of T. gondii (Heintzelman and Schwartzman, 1997). Homologues of Tgmyo-A (>50% identity) are myo-A molecules described in Plasmodium species (Pinder et al., 1998; Hettmann et al., 2000; Margos et al., 2000; Matuschewski et al., 2001), Babesia bovis (Lew et al., 2002), as well as two additional Toxoplasma myosins, Tgmyo-D (Hettmann et al., 2000) and Tgmyo-E. They are the smallest myosins found so far within the myosin superfamily with molecular masses between 82 and 93 kDa. These myosin molecules lack an obvious neck region and possess short tails. A truncated IQ motif was described in Tgmyo-A (Herm-Gotz et al., 2002) while in others no IQ motifs have been identified as yet, a novelty so far only found in these apicomplexan myosins. In other myosins the helical neck region

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Table 1

Descriptive data for the best described apicomplexan myosin molecule, and their associated accession numbers. References are given in the text

Species

Myosin name

Gene database accession no.

MW [kDa]

Plasmodium falciparum

Pfmyo-A Pfmyo-B Pfmyo-C Pfmyo-D Pbmyo-A Pymyo-A

Y09693/AF105117/ AF222716/ AF222717/ AF376800/ AF286048/AF255909/ AF286049

Tgmyo-A Tgmyo-B Tgmyo-C

berghei yoelii Toxoplasma gondii

AA at G699a

IQ motif

90 88 part seq 245 90 90

T Q E T N N

S S G G S S

n.i. n.i. 5 1 n.i. n.i.

Peripheral unknown unknown unknown periphery ook/spz peripheral/diffuse spz

AF006626 AF006627/ AF006628/

93 114 125

Q Q Q

S S S

1b 1 1

Tgmyo-D Tgmyo-E

AF105118/ AF221131/

91 93

N Q

S S

n.i. n.i.

Peripheral cytoplasmic? perinuclear/posterior pole peripheral bradyzoites

Bbmyo-A Bbmyo-B Bbmyo-C

AF403045 AF403046 AF403047

82 ?(part seq) ?(part seq)

N D ?

S S ?

n.i. 1 ?

perinuclear unknown unknown

n.i., not identified; part seq, partial sequence; ook, ookinete; spz, sporozoite.

Amino acid corresponding to position of G699 in chicken muscle myosin II. Truncated.

a b

Localisation


Babesia bovis

AA TEDS rule site


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may act as lever arm and contain IQ motifs which function as light chain or calmodulin binding sites, the core consensus being IQXXXRGXXXR (Mercer et al., 1991). The number of IQ motifs can vary, ranging from 1 to 6 (reviewed by Cheney and Mooseker, 1992). Myosin light chains are assumed to stabilise the neck and may be important for regulation of myosin activity. Tgmyo-B and TgmyoC are splice variants encoded by the same gene and varying in the length of their tails (Heintzelman and Schwartzman, 1997; Delbac et al., 2001). Tgmyo-B/C posses a single divergent IQ motif in their assumed neck region (Heintzelman and Schwartzman, 1997). The tail region of myosins varies widely and confers functional specificity to the molecules. For example, the tails of filamentforming and other two-headed myosins possess alpha-helical sequences that form a coiled-coil and allow these molecules to dimerise. Myosin tails may also contain functional motifs associated with signal transduction, membrane or receptor protein binding (such as Scr homology 3 (SH3) domain, GTPase activating protein (GAP) domain, pleckstrin homology (PH) domain, myosin tail homology 4 (MyH4), talin homology domain and zinc-binding domain; for a review see Oliver et al. (1999). The tails of Toxoplasma myosins have a high basic charge. This feature has been attributed to membrane binding properties of myosin I (Adams and Pollard, 1989). They do not show homology to functional domains previously described in other myosins (Heintzelman and Schwartzman, 1997; Hettmann et al., 2000). Apart from actin and nucleotide binding sites, myosin heads may contain insertions or extensions with supplementary functions as well as single amino acids of putative regulatory or functional importance. A putative regulatory site is the TEDS rule site (reviewed by Bement and Mooseker, 1995) consisting of a single amino acid 16 residues upstream of a DALAK motif which is not conserved in all myosins. Amino acids present at this site were found to be the phosphorylatable amino acids threonine (T) or serine (S) in myosin I of Acanthamoeba, Dictyostelium, Aspergillus, yeast and class VI myosins, or the negatively charged amino acids, aspartic acid (D) or glutamic acid (E) in most other myosins. Very few exceptions to

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this rule had been found within the myosin superfamily (see review by Mooseker and Cheney, 1995), but none of the Toxoplasma myosins follows the TEDS rule, either glutamine (Q) or asparagine (N) is found in the relevant position (Heintzelman and Schwartzman, 1997; Hettmann et al., 2000, see Table 1). The divergence from the TEDS rule has raised the question of how the activity of these myosins may be regulated (Heintzelman and Schwartzman, 1997). Another exceptional feature of the Toxoplasma myosins is that a serine residue is found at the position that corresponds to glycine699 of chicken muscle myosin II (Hettmann et al., 2000, see Table 1). A glycine residue at this position is conserved in many myosins and has been proposed to be of functional interest by acting as pivot for the lever arm (Kinose et al., 1996). Apart from pfmyo-A, three other myosin gene sequences have been found in P. falciparum, namely pfmyo-B, pfmyo-C and pfmyo-D (see Table 1). Pfmyo-B has been assigned to class XIV. Although more divergent (Pfmyo-B shows only 35% identity to Pfmyo-A and even lower scores to all other class XIV myosins), Pfmyo-B shares certain features with other members of class XIV myosins. It is a small molecule (predicted 188 kDa), with no apparent neck region or IQ motifs, and does not follow the TEDS rule; a glutamine residue is found at that position. Like the Toxoplasma myosins (Hettmann et al., 2000) and Pfmyo-A, it carries a serine residue at the position corresponding to glycine699 of chicken muscle myosin. In marked contrast to Toxoplasma myosins and Pfmyo-B, PfmyoC and -D as well as Pfmyo-A (Pinder et al., 1998) follow the TEDS rule (see Table 1). Interestingly, Pfmyo-C and Pfmyo-D show additional characteristics of ‘typical’ myosins: both carry a glycine at the position corresponding to the conserved glycine699, and have IQ motifs (Lew et al., 2002). Pfmyo-C and Pfmyo-D are relatively large molecules of predicted molecular masses of >200 kDa. When compared to other class XIV myosins, Pfmyo-C and -D have large inserts in the head region (Lew et al., 2002). In phylogenetic tree analysis Pfmyo-C and -D do not align with members of class XIV myosins and show only 20% identitiy to each other.


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B. bovis Bbmyo-A shows typical features of apicomplexan myosinA. The partial sequence known from Bbmyo-C shows homology to Pfmyo-C but Bbmyo-B has no homology to any other apicomplexan myosin. Unusually, Bbmyo-B has an IQ motif within its head region. Considering the deviation of Pfmyo-C/Bbmyo-C, and Pfmyo-D from the other class XIV myosins, it is possible that they constitute a new class(es) of myosins (Lew et al., 2002). In a PCR based approach, nucleotide sequences of approximately 150 bp were amplified from a number of apicomplexan species between the conserved GESGAGKT and EAFGNAKT domains (Heintzelman and Schwartzman, 2001). This screen resulted in the identification of 4–7 myosin sequences in each of T. gondii, P. falciparum, N. caninum, E. tenella, Sarcocystis muris, B. bovis and C. parvum. In an alignment of this short nucleotide stretch most of the myosins grouped as Tgmyo-A or Pfmyo-A like myosins. Only three (Ncmyo-b, Etmyo-f, Bbmyo-g) grouped with Tgmyo-B/C, and two (Ncmyo-c, Bbmyo-f) showed close similarity to Pfmyo-C. It is noteworthy that in this screen Pfmyo-D, for example, was not picked up, and it is therefore possible that additional myosins may still be found in Toxoplasma or other apicomplexan parasites.

4.3.2. Cellular localisation and function Myosins have been implicated in a variety of cellular processes including membrane and vesicle transport, cell locomotion, phagocytosis, cytokinesis and muscle contraction (reviewed by Mermall et al., 1998). The cellular localisation of myosins may provide clues for the function they perform. Myo-A had been implicated in motility and host cell invasion based on its intracellular peripheral localisation in close association with the plasma membrane (Pinder et al., 1998; Heintzelman and Schwartzman, 1999; Hettmann et al., 2000; Margos et al., 2000; Matuschewski et al., 2001). This function has been confirmed in Toxoplasma by use of an inducible gene knock-out system (Meissner et al., 2002b). Although parasites lacking the myosinA gene were non-motile, host cell

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invasion and egression was not completely prevented. Therefore, it remains to be seen whether any of the other myosins is involved in these processes (Meissner et al., 2002b). Tgmyo-C and Tgmyo-B seem to be involved in the proper distribution of the inner membrane complex and the surrounding plasma membrane between daughter cells during endodyogeny (Delbac et al., 2001), the form of cell division found in Toxoplasma. Tgmyo-D was also found near the cell periphery but its distribution was more diffuse and patchy (Hettmann et al., 2000) and no particular function has been assigned to it. Tgmyo-E mRNA was found in bradyzoites rather than tachyzoites but its function is unclear (Delbac et al., 2001). Nothing is known about the expression, localisation or function of Pfmyo-B, -C or -D, Bbmyo-B and -C. The membrane binding properties of myosin I have been attributed to a high basic charge in the tail domain (Adams and Pollard, 1989). Characterisation of Tgmyo-A, Tgmyo-D and Tgmyo-C provided evidence that the tail domain determines the final destination of the molecules (Hettmann et al., 2000; Delbac et al., 2001). It was further shown that in particular two arginine residues within the last 22 amino acids of the tail domain of Tgmyo-A seem responsible for directing the molecules to the peripheral cell membrane (Hettmann et al., 2000). This suggested that these amino acids are part of a signal patch which directs the molecule to the membrane or that they may be part of a motif involved in receptor binding of the myosin tail at a peripheral membrane, or both. The observation that a chimeric tail domain of Pfmyo-A and GFP was found in the cytoplasm rather than peripherally in Toxoplasma may support the idea of specific receptor protein interaction (Hettmann et al., 2000). Recently, a 50 kDa myristolated protein was identified in T. gondii which co-purifies with the tail region of Tgmyo-A. This protein has been termed Tgmyo-A docking protein (TgMADP) (gene bank accession no. AF453384) and may fulfil such a role (Herm-Gotz et al., 2002). An important but unresolved question with major implications regarding the motor mechanism is whether the myo-A tail is bound


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to the plasma membrane or the cisternal membrane (Pinder et al., 2000). A myoA tail domain interacting protein (MTIP) was found in P. yoelii sporozoites localising to the IMC membrane of sporozoites (Bergman et al., 2003) supporting the left-hand model in Figure 1. Conversely, and favouring the model on the right in Figure 1, immuno-EM studies on T. gondii parasites (after hypotonic swelling to separate IMC from plasma membrane) demonstrated that actin was associated with the IMC whilst an antibody to the LEAF region of myosin reacted with the plasma membrane (Dobrowolski et al., 1997a). Some myosin tail domains seem to be involved in regulation of protein expression. Most interestingly, recombinant myc-Tgmyo-A downregulated the expression of endogenous Tgmyo-A protein (Hettmann et al., 2000). It was shown that this downregulation takes place at the post-translational level since the amount of Tgmyo-A mRNA was not influenced by expression of recombinant myc-myoA (Meissner et al., 2001). How this regulation works and why expression of endogenous protein is downregulated is unknown.

4.4. Myosin Binding Proteins/Regulatory Proteins The activity of myosins is regulated by light chains bound to their neck region and may be modified by binding of molecules to functional domains in the tail region. In several myosins the regulatory light chains have been identified as calmodulin (reviewed by Mooseker and Cheney, 1995). In Toxoplasma, calmodulin was found to be concentrated near the apical end of tachyzoites and to co-localise with an undefined myosin molecule (Pezzella-D’Alessandro et al., 2001). However, whether calmodulin binds to any Toxoplasma myosin and does indeed function as a myosin light chain has still to be determined. A calmodulin-like protein of 31 kDa which co-purified with Tgmyo-A was shown to bind to the tail of Tgmyo-A (Herm-Gotz et al., 2002). The molecule has been named myosin light chain 1 (TgMLC1) and appears to constitute a novel MLC of Apicomplexa.

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TgMLC1 shows homology to MTIP in P. yoelii sporozoites (43% aa identity) (Bergman et al., 2003). The protein contains a single motif with similarity to an EF-hand domain. Immuno-EM and TritonX100 extraction indicate association with the IMC membrane. MTIP was shown to be expressed in sporozoites and in asexual blood stages of P. yoelii. Mutagenesis of the dibasic motif directing myo-A to the cell periphery (see above. Hettmann et al., 2000) abolished the interaction between MTIP and myo-A (Bergman et al., 2003). Although the aa sequence of MTIP does not predict membrane attachment, in Western blot analysis MTIP remained in the pellet fraction suggesting its binding to an unidentified anchor protein. Whether this anchor protein might be the Plasmodium homologue of TgMADP (shown to co-purify in Toxoplasma with the TgmyoA-TgMLC1 complex) (Herm-Gotz et al., 2002) needs to be established. The chemical inhibitor KT5926, which in animal cells acts on myosin light chain kinases, inhibited host cell invasion of Toxoplasma by inhibiting host cell attachment of parasites (Dobrowolski et al., 1997a). Detailed experiments on the effect of KT5926 and immunofluorescence data indicated that release of the micronemal protein MIC2, a Toxoplasma adhesin (reviewed by Naitza et al., 1998), was prevented while secretion of molecules from dense granules (dense granule protein GRA1) seemed unaffected. Interestingly, BDM did not have the same effect (Dobrowolski et al., 1997a). Although these results might suggest involvement of a myosin in discharge of the micronemes (and therewith in host cell invasion), one must ask why the myosin ATPase inhibitor BDM did not inhibit the same myosin. In a more recent study, Kieschnick et al. (2001) found that KT5926 blocks the phosphorylation of only three Toxoplasma proteins. A KT5926 sensitive calmodulin-like domain protein kinase (CDPK1) was identified in tachyzoites whose activity was calcium-dependent but not calmodulin sensitive. However, the nature of the proteins phosphorylated by this kinase is as yet unknown. While much progress has been made through the identification and localisation of myosin molecules especially in Toxoplasma parasites, many questions still remain to be solved on the regulation of


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apicomplexan myosins and binding partners. The development of an inducible expression system, as recently described in Toxoplasma (Meissner et al., 2002b), is an important step towards the generation of conditional knockouts and functional analyses of essential molecules such as myosins.

4.5. Inner Membrane Complex Protein 1 (IMC1) and 2 (IMC2) Two proteins of the subpellicular network have been identified in T. gondii with electrophoretic mobilities of 80 and 150 kD, respectively (Mann and Beckers, 2001). Extended coiled coil domains in their sequences imparted a resemblance to mammalian intermediate filament proteins. TgIMC1 showed homology to articulins, the major components of the membrane skeleton of algae and free-living protozoans. In Toxoplasma, this subpellicular network is formed as a detergent-soluble flexible structure in developing daughter tachyzoites. In mature parasites the network is detergent insoluble and provides rigidity to the cell. This difference in stability is likely to be mediated by proteolytical processing of TgIMC1 (Mann et al., 2002).

4.6. Tubulin and Microtubules (mt) As one of the major components of the cytoskeleton, mt play key roles in regulating cell shape, polarity and the distribution of organelles. They also generate motility, notably in stable structures such as cilia and flagella, and in the mitotic and meiotic spindles. Like actin, cytoplasmic mt are highly dynamic, allowing a plastic structural framework which can be regulated by several cellular effectors, including mt associated proteins (MAPs) and motor proteins (Drewes et al., 1998). The organisation of mt in many of the Apicomplexa is well documented: mitotic and meiotic spindles, centrioles, subpellicular mt and flagella in male gametocytes. In the invasive zoites of the

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Apicomplexa, the predominant mt structures are the subpellicular mt and the conoid. The unusual tubulin polymer nature of the conoid has only recently been described however (Hu et al., 2002). The conoid structure incorporates yellow-fluorescent-protein -tubulin, and was successfully labelled by antibodies to - and -tubulin. Surprisingly, however, the tubulin monomers do not form the classical microtubule structure, but ribbons of nine protofilaments forming a comma shape in cross section. The authors hypothesise that this unique structure may facilitate the extreme curvature required of the conoid tubulin polymers. Mt consist of - and -tubulin dimers. Both - and -tubulin bind GTP, but only the GTP on the -tubulin is hydrolysed to GDP during mt polymerisation; this in turn can be exchanged for GTP. -tubulin is found in mt organising centres (MTOC’s), and may act as a template for mt polymerisation. The more recently described , ", and tubulins, are thought to be predominantly associated with basal bodies and centriolar regions of eukaryotic cells (McKean et al., 2001). In vertebrates and plants, multigene families produce multiple isoforms of both - and -tubulin. In unicellular organisms, there is no general rule; they often possess several and genes scattered throughout the genome, although in Leishmania and trypanosome parasites the genes are clustered (Landfear et al., 1983; Seebeck et al., 1983). In general, the apicomplexa appear to have single genes expressed at any one stage of the lifecycle and they appear to be unlinked (Caccio et al., 1997; Bonafonte et al., 1999). However, differential splicing and post-translational modifications are mechanisms available for generating more than one isoform from single copy genes. Several post-translational modifications (ptms) of both tubulin gene translation products have been characterised: tyrosination and acetylation of -tubulin, and phosphorylation, polyglutamylation and polyglycylation of both - and -tubulin subunits, all of which are enzymatically mediated and are reversible (Gull et al., 1986; Sasse et al., 1987; Idriss, 2000; McKean et al., 2001). Of these, tyrosination, glutamylation and glycylation are unique to tubulin (Idriss, 2000) making it one of the most modified proteins known.


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4.7. Apicomplexan Tubulin Genes Tubulin sequences are known for Plasmodium spp. (Akella et al., 1988; Delves et al., 1989; Wesseling et al., 1989a; Holloway et al., 1990; Sen and Godson, 1990), T. gondii (Nagel and Boothroyd, 1988), C. parvum (Nelson et al., 1991; Caccio et al., 1997; Bonafonte et al., 1999), Babesia bovis (Casu, 1993) and E. tenella (Zhu and Keithly, 1996). However, remarkably little is known of post-translational modifications, mt based motor proteins and MAPs. The first of the apicomplexan - and -tubulin genes to be characterised were those of T. gondii (Nagel and Boothroyd, 1988). They were found to be single copy, unlinked and, unusually for the parasitic protozoa, to contain several introns. One of the -tubulin introns is conserved in all the other apicomplexa studied. As with other tubulin genes, the region of greatest variability was at the carboxy terminus. P. falciparum has four single copy genes: -I, - and -tubulin, transcribed throughout the lifecycle (Delves et al., 1989, 1990; Holloway et al., 1989, 1990; Wesseling et al., 1989a; Sen and Godson, 1990; Maessen et al., 1993) and -II tubulin, transcribed predominantly in gametocytes, when unique transcripts are also upregulated (Delves et al., 1990; Rawlings et al., 1992). P. falciparum -tubulin gene has been located on chromosome 10 (Holloway et al., 1990). The protein is 445 amino acids long; the gene has two introns and no unusual features. More than one transcript has been noted in northern blots for both - and -tubulin genes (Sen and Godson, 1990). The P. falciparum - and -tubulins show high identity with other sequenced tubulins (e.g. P. falciparum -tubulin has 88% homology with chicken brain tubulin) (Sen and Godson, 1990; van Belkum et al., 1991) but for P. falciparum -tubulin, as with other -tubulins, the identity is comparitively low (Maessen et al., 1993). -tubulin has also been noted in Plasmodium (McKean et al., 2001). The C. parvum -tubulin gene is 1365 bp long, predicting a typical -tubulin gene of 50.5 kD. It is most like T. gondii and P. falciparum genes, with 88.7% and 88.2% similarities respectively, compared to, for instance, 79.4% similarity to human -tubulin. The -tubulin

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gene is 1618 bp long, it has one intron, which has been conserved in most apicomplexan -tubulin sequences described to date. The open reading frame predicts a typical -tubulin of 49.74 kD. Its highest identity is with the T. gondii gene: 86.9%, but this is lower than the identities reported among the other apicomplexans (Caccio et al., 1997); for example, the -tubulin gene of E. tenella, which encodes a typical -tubulin of 499 aa, has 92–96% identity with other apicomplexa (Zhu and Keithly, 1996). This similarity extends to its three introns, which are found in the same positions as those found in T. gondii. The -tubulin gene of B. bovis, which encodes a protein of 441 aa, has 92.5% homology to that of P. falciparum but no introns (Casu, 1993).

4.8. b-Tubulins -Tubulins are of particular interest because they have been successfully used as the target for anti-parasitic chemotherapy. Studies of the apicomplexan -tubulin sequences indicated mixed susceptibilities to benzimidazole compounds (van Belkum et al., 1991; Edlind et al., 1994). Many of the parasitic protozoa are susceptible to the dinitroaniline herbicides, which have low toxicity to animal tubulins (Morrissette et al., 1994; Kaidoh et al., 1995; Arrowood et al., 1996; Benbow et al., 1998; Fowler et al., 1998). It has been suggested that this may be due to the phylogenetic similarity of the parasitic protozoan -tubulins to plant and algal tubulins (Fong and Lee, 1988; Caccio et al., 1997). The apicomplexan -tubulin sequences are closest to the single copy genes of the ciliates and also resemble those of the slime moulds, filamentous fungi and green algae (Zhu and Keithly, 1996). However, studies of the C. parvum genome (Edlind et al., 1994; Bonafonte et al., 1999) suggest that it is -tubulin that is susceptible to the dinitroanilines. All the apicomplexan -tubulins described to date have the tetrapeptide MREI, which is required for expression autoregulation (Caccio et al., 1997). The mechanism for -tubulin autoregulation is not known (Bachurski et al., 1994). The apicomplexan -tubulins


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also have a conserved site for polyglycylation at the carboxy terminus (Zhu and Keithly, 1996). The addition of oligoglycyl side chains is particularly associated with stable mt structures, such as axonemes. In Tetrahymena polyglycylation of -tubulin was essential for growth, motility and cytokinesis; however it is not a requirement for all single cell organisms, since in Trypanosoma brucei this particular ptm was not detected (reviewed by Gull, 2001). The typical eukaryotic promoter regions are absent from the apicomplexan -tubulin genes; in fact Cryptosporidium appears not to have any conserved promoter motifs (Zhu and Keithly, 1996; Caccio et al., 1997).

4.9. Post-translational Modifications of a-tubulin The two -tubulin specific ptms have been studied in the apicomplexa. In the vast majority of -tubulin genes, a lysine at position 40 can be acetylated. At the 30 end the terminal tyrosine can be removed to expose glutamic acid. Both these processes are mediated by specific enzymes and are reversible. These two ptms are interesting because they produce isotypes with distinct cellular locations, and the role each plays may be dependent on the presence of the other.

4.9.1. Acetylation -Tubulin becomes acetylated after mt have been assembled. It is a highly specific ptm, which has been found in a number of organisms. The level of acetylation is thought to reflect the exposure time to tubulin acetyl transferase, and is generally found in mt which form stable or cross linked organelles (reviewed by McKean et al., 2001). For example, there is more acetylated than non-acetylated tubulin in the subpellicular mt of trypanosomes (Gull et al., 1986). A variety of other proteins are known to be acetylated, particularly nuclear proteins. In this context, acetylation is thought to regulate DNA


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recognition, protein–protein interactions, protein stability and may also have a role in signalling. The function of tubulin acetylation remains unclear, however, since the substitution of one form for the other had no discernible phenotype (reviewed by Kouzarides, 2000). Takemura et al. (1992) suggest that it may be involved in the binding of crosslinking MAPs. Read et al. (1993) studied the acetylation potential of P. falciparum -tubulins. A lysine is found at position 40 in the -tubulin I gene, but not the sexual-stage specific -tubulin II gene (Holloway et al., 1990). In fact, the sequence between positions 40 and 45 represents the major difference between -tubulin I and -tubulin II genes. It is difficult to assess the significance of this, since an antibody to acetylated tubulin showed no reactivity with the tubulin of asexual malaria parasites (Read et al., 1993). The authors point out that this may be because of motif differences downstream of the position 40 lysine (Holloway et al., 1989). The C. parvum -tubulin gene has a methionine insertion at position 38 (Bonafonte et al., 1999) which may also affect its acetylation potential.

4.9.2.

Tyrosination

Most -tubulin genes encode a C-terminal tyrosine, which can be removed to expose glutamic acid. Detyrosination of -tubulin is thought to be mediated by a specific, mt associated, carboxy peptidase, as the mt is generated. When the mt depolymerise, the reaction is reversed by tubulin tyrosine ligase in the cytoplasm (reviewed by Gull et al., 1986). Several essential processes, such as growth, differentiation and motility, require -tubulin detyrosination (reviewed by Eiserich et al., 1999), so many studies have searched for a role for the tyrosination cycle in the kinetics of mt assembly, mt dynamics, mt stability and distribution (reviewed by Idriss, 2000). More recently it has been suggested that detryrosination may act as a signal that segregates the two functional activities of mt: the dynamics of polymerisation vs. mt-organelle interactions (Kreitzer et al., 1999).


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In several of the apicomplexan -tubulin genes, the carboxyterminal tyrosine is missing. In C. parvum the -tubulin gene has a Phe instead of a Tyr (Bonafonte et al., 1999), similar to several chick -tubulin genes (reviewed by Idriss, 2000). Although the P. falciparum -tubulin I gene has the site for detyrosination, AspTyr, and -tubulin I reacts with an antibody to tyrosinated tubulin (Read et al., 1993), the P. falciparum tub II gene has a terminal Glu (Holloway et al., 1990). A similar dichotomy exists in P. yoellii -tubulin genes (Akella et al., 1988). This raises the possibility that the Glu is removed later to reveal the penultimate Tyr, a process which would require a novel enzyme, or that Tyr is added as a ptm (Holloway et al., 1990), as seen in the mammalian M4 tubulin gene (Gu et al., 1988). Another possibility is that the -tubulin II is excluded from the tyrosination cycle. Opting out from the cycle would not be unique (Alfa and Hyams, 1991; Schneider et al., 1999). Detyrosinated mts were shown to have enhanced stability against cold-initiated depolymerisation (reviewed by Idriss, 2000). This may be relevant: a drop in temperature is one of the cues for exflagellation in Plasmodium. Equally, detyrosination may be a consequence of stabilisation, possibly by mt capping (reviewed by Idriss, 2000). Another interesting observation is that the molecular motor kinesin binds with higher affinity to detyrosinated mt (Liao and Gundersen, 1998; Kreitzer et al., 1999) and the interaction of cytoplasmic-dynein with mt is disrupted when mt are irreversibly modified with 3-nitrotyrosine (Eiserich et al., 1999). A possible distinction of detyrosinated mt, therefore, is that they may provide special tracks for motor proteins. The flagellar axoneme mt are driven by dynein, so the detyrosinated state of the Plasmodium -tubulin may facilitate flagellar assembly.

4.10. Microtubule Associated Motor Proteins Mt and their associated motor proteins, the kinesin family and cytoplasmic dynein, are essential for mitosis (Vaisberg et al., 1993; Barton and Goldstein, 1996). The recent genome projects on

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Plasmodium, Cryptosporidium and the expressed sequence tag (EST) projects for Toxoplasma, Eimeria and Neospora have sequences which have been identified as encoding putative dyneins, kinesins and kinesin-like proteins (reviewed by Morrissette and Sibley, 2002a). With the exception of mitosis, however, their relevance to apicomplexan biology is unclear. In higher eukaryotes they are constitutive components of the intracellular transport system, responsible for vesicle, Golgi tubule and organelle trafficking (Hirokawa et al., 1998; Lippincott-Schwartz, 1998). In flagella and cilia, dynein arms are the ATP-driven motors that generate motility (Preston et al., 1993b). In the asexual stages of P. falciparum one can distinguish between mitotic and post-mitotic events (Pouvelle et al., 1994; Fowler et al., 1998). However, dissecting roles for motor proteins in merozoite motility and/or merozoite assembly is more difficult. We have demonstrated the presence of both kinesin and dynein in P. falciparum merozoites by immunofluorescence. Neither co-localised with the subpellicular mt, but appeared as strong, peripheral arcs of fluorescence in the anterior third of the cells during the latter stages of merogony. This late appearance suggests either merozoite component emplacement or a mustering for action during host-cell escape and invasion. The inhibition of either could explain a reduction in invasion rates observed when post-mitotic parasites were incubated with dynein ATPase inhibitors (Fowler et al., 2001). The role of dynein and kinesin requires further investigation, with more specific ways of inhibiting their motor activity and ultrastructural localisation in merozoites, other stages of P. falciparum and other apicomplexans.

5. MODELS OF MOTILITY Building on observations first made with Gregarina King (1981, 1988) put forward a convincing model of apicomplexan motility, proposing a cell-surface linear motor, which could drive three types of locomotion. The first is gliding motility, which has been observed in


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the sporozoites and ookinetes of many apicomplexans, tachyzoites of Toxoplasma and gregarine trophozoites. Gliding is the ability to traverse substrates, without any apparent body shape changes, or employment of locomotory organelles, such as cilia or flagella (Preston et al., 1993a). Plasmodium and Eimeria sporozoites glide over glass slide substrates, often in a clockwise circular motion (Vanderberg, 1974; Russell and Sinden, 1981). Toxoplasma tachyzoites show a similar pattern of motion, if they are found positioned on their ‘right’ surface; on their ‘left’ surface they traverse substrates in a helical rotation (Endo and Yagita, 1990; Frixione et al., 1996; Hakansson et al., 1999). Plasmodium ookinetes require the presence of insect cells to be actively motile (Speer et al., 1975; Siden-Kiamos et al., 2000). Fluctuations in force generation and changes in parasite orientation mean that the gliding motion is not continuous over short time scales (Preston et al., 1993a; Hakansson et al., 1999). Many apicomplexan parasites leave a trail of surface proteins as they glide (e.g. Russell and Sinden, 1981; Stewart and Vanderberg, 1988; Arrowood et al., 1991; Dobrowolski et al., 1997a), and this trail also includes lipids from the plasma membrane as shown in T. gondii (Hakansson et al., 1999) and Plasmodium ookinetes (Shahabuddin et al., 1998). The second of the types of locomotion is the capping of crosslinked surface molecules from the anterior to the posterior of parasites (Russell and Sinden, 1981; Dubremetz et al., 1985). The capping of the surface protein of Eimeria tenella sporozoites is an example of this, initiated by specific monoclonal antibodies, crosslinked by anti-IgG, followed by the translocation of these complexes to the basal pole at the rear of the sporozoite (Speer et al., 1985). Polystyrene beads have been used to study the capping process in Gregarina trophozoites, Eimeria sporozoites and Plasmodium sporozoites and showed that capping occurs in a linear fashion, and occurred primarily from the anterior to the posterior, although Plasmodium sporozoites demonstrated bi-directionality of bead movement (reviewed by King, 1988). These experiments also suggested that the capping of beads was analogous to gliding across a substrate, the beads acting as a mobile substrate. Both of these

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processes can be inhibited by cytochalasin B (CB), which indicates that filamentous actin is required (Russell and Sinden, 1981; Gut and Nelson, 1984; Stewart and Vanderberg, 1988; Dobrowolski and Sibley, 1996). The third type of locomotion, invasion of host cells, can also be inhibited by CB (Jensen and Edgar, 1976; Ryning and Remington, 1978; Miller et al., 1979). For some apicomplexans, such as T. parva and C. parvum sporozoites, invasion is dependent on subverting the actin cytoskeleton of the host cell (Shaw and Tilney, 1999; Chen and LaRusso, 2000). Experiments treating either host cells or parasites with selected chemical inhibitors, indicated that Eimeria and Toxoplasma actively invaded their host cells (Jensen and Edgar, 1976; Werk and Bommer, 1980; Dobrowolski and Sibley, 1996). Dobrowolski and Sibley (1996) confirmed that Toxoplasma invasion depended on parasite actin, rather than host cell actin, by using CB resistant parasites and host cells, respectively. In the presence of CB, resistant parasites could enter sensitive host cells, but not vice versa. The classical coccidian invasion process is initiated when a parasite either attaches to a host cell in an apically directed manner, or attaches and orientates to bring the apex in contact with the host cell membrane (Sinden, 1985; Bannister and Dluzewski, 1990; Morisaki et al., 1995). Host cell entry requires the locomotion of the parasite (in many cases into a parasite-induced vacuole), through an aperture bounded by an electron dense structure called the tight or close junction—so called because of the close apposition of the parasite plasma membrane with the host cell membrane (Aikawa et al., 1978; Michel et al., 1980; Bannister and Dluzewski, 1990). In T. gondii and Plasmodium ookinetes the tight junction can be observed as constriction upon parasite entry. Malaria merozoite invasion is inhibited by CB at the point where the apical contact has been established. Although some evidence of vacuole formation is apparent, the merozoite fails to enter the cell (Miller et al., 1979; Dluzewski et al., 1989). The mechanism responsible for gliding and capping is also thought to propel the invading parasite through the aperture into the lumen of the host cell, by using the tight junction


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as the ‘substrate’ (Russell, 1983; King, 1988). It is important to note that ookinetes and sporozoites (invading into salivary glands) appear not to produce a parasitophorous vacuole as they traverse through cells in their migration through the host, and there may be several differences in the biology of cell invasion in these instances (Sinden, 1985). The cell surface linear model is thought to account for all three forms of motility. It proposes, as illustrated in Figure 1, that an actomyosin motor is located directly under the plasma membrane, connected to the exterior via transmembrane proteins which adhere to the substrate. The force generated by the lever action of the myosin head against the actin filament is brought to bear against the substrate by the interaction of either myosin or actin with the cytoplasmic tails of these non-compliant proteins. It is difficult to resolve which component of the acto-myosin motor interacts with the proteins that transduce the force, since the spatial resolution of immunogold localisation of myosin has been limited by the size of the detection antibodies. Binding studies, such as yeast two-hybrid systems (Bergman et al., 2003), and immunoprecipitation of myosincomplexes (Herm-Gotz et al., 2002) have provided some answers by revealing what interacts with the myosin tails (see also review by Opitz and Soldati, 2002). Although the TSP related family of adhesive proteins have short cytoplasmic domains, and as such, present good candidates as the transducing agents, there may well be other proteins which form the link between the adhesive/receptor proteins and the motor. It also seems likely that different molecules are involved in different modes of motility. Russell (1983) suggested that the availability and specificity of the parasite adhesive/receptor molecules would determine when the parasite was motile, and what it could use as a substrate. It has also been noted that the number of micronemes (the organelles thought to extrude these molecules) found in different apicomplexan lifecycle stages is related to the level of parasite motility (Blackman and Bannister, 2001). The discovery of actin binding proteins (ABPs) in malaria parasites (Tardieux et al., 1998a,b; Forero and Wasserman, 2000) and T. gondii (Allen et al., 1997; Poupel et al., 2000) has indicated that actin

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dynamics are extremely important, and highly regulated during invasion. This was also implied by the sequestration of monomer actin in T. gondii (Dobrowolski et al., 1997b) and the pattern of actin polymerisation induced by jasplakinolide (Shaw and Tilney, 1999; Wetzel et al., 2003). It seems likely that in response to cell signalling molecules, short, capped, actin filaments form when required for motility and in the appropriate location, i.e. at the apex and under the plasma membrane. If the membrane bound myosin is situated on the outer cisternal membrane, then these filaments could be shuttled backwards on the myosin molecules. The intramembranous proteins would interact with the actin, and be moved to the posterior of the cell, thus propelling the parasite forwards. Although the fluid nature of the lipid bilayer forming the plasma membrane is necessary for the flow of the ligand proteins, the converse is true for the cisternal membranes. To achieve this forward propulsion the myosin molecules would have to be securely anchored into the cisternal membrane, which in turn would need to be firmly supported. This might be achieved by the binding of myosin to a framework of proteins within the cisternal membranes. In addition, a level of rigidity is probably provided by the subpellicular mt and network (Mann and Beckers, 2001). Alternatively, the actin filaments may polymerise, possibly on short filaments already present in the pellicle, and bind to the cisternal membranes or proteins in the framework. In this scenario, the myosin tails would be bound to the plasma membrane proteins, presumably within the membrane, and would pull them through the plasmalemma as they progressed along the actin filaments. The alternatives are shown schematically in Figure 1. The movements of some of the apicomplexa are difficult to ascribe to a simple gliding motion driven by a linear motor. Eimeria sporozoites move with a helical gliding motion, and bend and flex (Russell and Sinden, 1981). The most detailed studies have been on T. gondii tachyzoites, which also move forward by a biphasic clockwise helical gliding motion. After moving one body length whilst rotating 180 the tachyzoite is left in an upright position and


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must re-orientate itself by touching its apical end and twisting so that ts convex side is also in contact with the substratum and its helical progression can continue (Hakansson et al., 1999). Tachyzoites go through a complex range of movements which contribute to this motility, including turning along their longitudinal axis, torsion of the anterior region, pivoting on the basal pole, flexing (or bowing) and retraction. A combination of the last two resulted in the end-toend vertical distance being reduced by approximately 50% (Frixione et al., 1996). An anticlockwise torsion has also been noted in invading T. gondii tachyzoites, and it seems likely that many of these movements are also involved in invasion (Chiappino et al., 1984). Helical gliding was inhibited by both cytochalasin D (CD) and myosin inhibitors, demonstrating dependence on an acto-myosin motor (Hakansson et al., 1999). Since Toxoplasma tachyzoite and Eimeria sporozoite subpellicular mt have an anticlockwise twist (as seen from the apex), and the orientation of the IMC plates follow the mt, it is logical to assume that they dictate the direction of the linear motor, and thus the clockwise forward motion of the parasite. The observations of torsion, pivoting, flexing and retraction are not so easy to explain in terms of the linear motor. Frixione et al. (1996) suggest that the interaction of the two cytoskeletal systems in the parasite might provide a mechanism, if suitably orientated forces, such as a contractile acto-myosin motor, were brought to bear upon the helical subpellicular mt. Yasuda et al. (1988) showed that immunogold anti-actin labelling appeared next to subpellicular mt and the authors speculated that actin and myosin might interact with the mt. However, if the two systems are separated by the IMC, as current evidence would suggest, (Morrissette and Sibley, 2002a) any interaction would require membrane spanning proteins. The conoid might form part of such an interactive system, since it can extend, retract, rotate and tilt (Pulvertaft et al., 1954). Previous electron microscopy studies have shown that the position of the polar ring in relation to the conoid determines whether the latter is extended or retracted. It is possible that the polar ring rotates on the spiral elements of the conoid, resulting in torsion of the cell body (Nichols and Chiappino, 1987). The apical rings and the conoid

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appear filamentous in E.M. studies and immunolocalisation studies have shown that antibodies to actin label the conoid area (Cintra and De Souza, 1985a; Nichols and Chiappino, 1987; Yasuda et al., 1988). However, initial suppositions that the conoid consisted of tubulin (Roberts and Hammond, 1970; D’Haese et al., 1977) have proved correct (Hu et al., 2002), as discussed above. Structures resembling dynein arms have also been noted within the conoid, and between the subpellicular mt and cisternal membranes (Nichols and Chiappino, 1987; Hu et al., 2002). The involvement of microtubule based motility systems has been largely ignored since apicomplexan gliding motility was inhibited by CD (King, 1988; Stewart and Vanderberg, 1988; Forney et al., 1998) and the microtubule inhibitors colchicine, nocadazole and vinblastine were shown to have no effect (Jensen and Edgar, 1976; Russell and Sinden, 1981). However, these toxins did not depolymerise the subpellicular mt of Sarcocystis, Besnoitia or Eimeria zoites (D’Haese et al., 1977; Russell and Sinden, 1981), possibly because they are heavily decorated with microtubule associated proteins (Morrissette et al., 1997) which often have a stabilising effect (Drewes et al., 1998). Gregarines of the genus Selenidium have a similar arrangement of subpellicular mt, but these were shown to be susceptible to destruction by colchicine or urea, and as a consequence the parasites became motionless (Schrevel et al., 1974). Incubation with dinitroaniline compounds, which may prevent mt polymerisation (Stokkermans et al., 1996; Shaw et al., 2000; Morrissette and Sibley, 2002b) or may depolymerise mt in plants and protozoa (Chan and Fong, 1994; Kaidoh et al., 1995), reduced the invasion rates of both C. parvum sporozoites and P. falciparum merozoites (Wiest et al., 1993; Fowler et al., 1998). The effect of mt disruption on invasion may have been indirect by preventing migration of micronemes to the apical tip of the parasite (Morrissette and Sibley, 2002b; Bannister et al., 2003; G. Margos and colleagues, unpublished observations). It should be noted that in Cryptosporidium mitotic spindles and centrioles have not been detected ultrastructurally, and care was taken in the P. falciparum experiments to temporally exclude any effects of the dinitroanilines on mitosis.


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Recently we have demonstrated the presence of kinesins and dynein, which are microtubule dependent motors, at the apex of mature P. falciparum merozoites. We have also shown that the dynein inhibitors, sodium ortho-vanadate and erythro-hydroxy-nonyl adenine (EHNA), had inhibitory effects in invasion assays (Schliwa et al., 1984; Shimizu, 1995) and the role of dynein in P. falciparum invasion requires further investigation (Fowler et al., 2001). Nevertheless, taken together, these studies suggest that a gliding motility generated by an acto-myosin motor may not be the only cytoskeletal mechanism important for apicomplexan motility. ACKNOWLEDGEMENTS The authors are greatly indebted to Conrad King (very much the father of apicomplexan motility studies), Jeni Fordham, Walter Gratzer and to our colleagues Lawrie Bannister and Anton Dluzewski for innumerable useful discussions; to Megan Morris for her patient compliance with the evolving requirements of the illustration; and to the Wellcome Trust for continuing generous and far-sighted support (currently Grant numbers 059566 and 069515). GHM is grateful to George Hinchliffe for musico-bibliographic advice. REFERENCES Adams, R.J. and Pollard, T.D. (1989). Binding of myosin I to membrane lipids. Nature 340, 565–568. Aikawa, M., Miller, L.H., Johnson, J. and Rabbege, J. (1978). Erythrocyte entry by malarial parasites. A moving junction between erythrocyte and parasite. Journal of Cell Biology 77, 72–82. Akella, R., Arasu, P. and Vaidya, A.B. (1988). Molecular clones of alphatubulin genes of Plasmodium yoelii reveal an unusual feature of the carboxy terminus. Molecular and Biochemical Parasitology 30, 165–174. Alfa, C.E. and Hyams, J.S. (1991). Microtubules in the fission yeast Schizosaccharomyces pombei contain only the tyrosinated form of alphatubulin. Cell Motility and the Cytoskeleton 18, 86–93.

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Index

Acanthamoeba 229 N-acetyl-O-trimethylsilyl derivatives of sphinganine 21 N-acetyl-O-trimethylsilyl derivatives of sphingosine 21 actin binding proteins (ABPs) 224–226, 245 actin depolymerizing factor (ADF) 224–225 actinosporean phase, characters of 60– 62 ageing 148–149 Albinaria coerulea 139 Alcyonella fungosa 64 AMA1 220–221 aminoacyl-tRNA synthetase 146 2-aminoethyl phosphonate (AEP) 19 Ancylostoma 176 Ancylostoma caninum 176, 184 Ancylostoma duodenale 158–159, 161, 165, 176, 179–180, 184, 186 anti-AMA1 220 Apicomplexa 214 apicomplexan actins 221–223 apicomplexan invasion, motility in 213–263 apicomplexan myosins 226–233 cellular localization and function 231–233 descriptive data and associated accession numbers 228 membrane binding properties 232


models of 242–249 structure 226–231 apicomplexan tubulin genes 237–238 apicomplexan zoite basic description 215–219 generalized body plan 216 apoptosis 148–149 Arthropoda 139 Ascaris lumbricoides 172 Ascaris suum 137, 151, 159, 161, 163, 172, 179 Aspergillus 229 Aspergillus fumigatus 26–27 Babesia 226 Babesia bovis 227, 231 Bathnostomum 176 Bbmyo-B 231 Bbmyo-C 231 benzimidazole (BZ) 170–171 Besnoita jellisoni 215 Brugia 169 Brugia malayi 151 Bryozoa morphology and development of Tetracapsuloides bryosalmonae in 87–89 prevalence of Tetracapsuloides bryosalmonae in 90 Buddenbrockia 44, 65, 96, 99, 103, 115 Buddenbrockia plumatellae 50–51, 59, 64–81, 89, 98–103, 113 behaviour 77–79

265

266 differentiation of sac-like stages 94 ecology 77–79 history 64–65 morphology and development of sac stage 74–76 morphology and development of vermiform stage 65–74 pathogenicity 77–79 timing of developmental sequence 74 Bunostomum 176 butanedione monoxime (BDM) 226 Caenorhabditis elegans 153, 159, 163, 179–180, 186–187 calmodulin-like domain protein kinase (CDPK1) 234 Candida albicans 26 Ceratomyxa 58 Ceratomyxa shasta 51 Chagas disease, Trypanosoma cruzi 2 chemical ionization 9 chemical ionization/mass spectrometry 23 Chloromyxum 52, 58 Chloromyxum diploxys 51 class VI myosins 229 Cnox genes 100–101 Crassostrea gigas 139, 142 Cristatella mucedo 51, 66, 72–79, 82, 87, 90 Cryptococcus neoformans 26–27 cryptomonad flagellates 107 Cryptosporidium 214, 239, 242, 248 Cryptosporidiun parvum 221, 223, 231, 237–238, 240–241 Cyprinus carpio 58, 63 malacosporean stages 79–80 cytochalasin B (CB) 244 cytochalasin D (CD) 247–248 cytoskeletal and motor proteins 221–242 cytoskeleton 213–263 organization of 215–219

INDEX DALAK motif 229 deaminated glycoinositolphospholipid 24 dendritic cells (DCs) 30 Dictyocaulus viviparus 158, 177–179 Dictyostelium 229 Dictyostelium discoidum 224 dihydrosphingosine (DHS) 26 diploblasts 111 Dirofilaria 169 Dirofilaria immitis 159, 161, 163, 179–180 Dirofilaria vivparus 184 DNA denaturation 156 DNA fragments 157 DNA molecules 174 DNA polymerase 156 DNA protein interaction(s) 147 DNA repair system 148 DNA replication and transcriptional processes 141 DNA sequence 147 DNA strand 175 DNA synthesis 141 DNA transfer 103 Drosophila melanogaster 143 EAFGNAKT 231 Echiichthys vipera 56 Echinococcus 164 Echinococcus granulosus 151 Echinostoma 164 Eimeria 214, 217, 242–243, 246, 248 Eimeria tenella 231 electron ionization 9 electron ionization/mass spectrometry 21, 23 Enteromyxum scophthalmi 54 erythro-hydroxy-nonyl adenine (EHNA) 249 Escherichia coli 146, 186 Esox lucius 82 ethanolamine phosphate (EtNP) 19

INDEX expressed sequence tag (EST) 242 extrusible organelles 104 F-actin 225–226 Fabespora sp. 52 Fabespora vermicola 52 Fasciola 164 fast atom bombardment (FAB) 9 fast atom bombardment/mass spectrometry 13–15 fatty acid methyl esters (FAMEs) 22 flatworms mitochondrial genomes 150–151 see also individual species Fredericella indica 82 Fredericella sultana 64–69, 72, 82, 87, 89–92 G-actin-binding/F-actin-severing proteins 225 gametogenesis 116 gas chromatography/mass spectrometry (GC/MS) 8 gene databases 138 genetic diseases 148 genetic markers, sequences and gene order 149–150 GESGAGKT 231 Globocephalus 176 Globodera pallida 153 glycoinositolphospholipids (GIPLs) 2–3, 14–16 biosynthesis 5 ceramide-linked 10 deamination 12 from Trypanosoma cruzi 1–41 function of 5 structure 4 see also Trypanosoma cruzi GIPL glycosidic bond cleavages 9 glycosylphosphatidylinositol (GPI) anchors 2–3 biosynthesis 5

267 glycosylphosphatidylinositol (GPI) biosynthesis in Trypanosoma cruzi 24–28 Grammocephalus 176 Gregarina 242–243 GTP 236 Haemonchus contortus 158, 165, 170 Haemonchus placei 165–166, 170 Halocynthia roretzi 142–144 Henneguya 60 Henneguya striolata 56 Heterorhabditis 168 1-O-hexadecyl-2,3-di-Otrimethylsilylglycerol 23 high performance liquid chromatography (HPLC) 14 Histoplasma capsulatum 26 Hoferellus gilsoni 55 Hox gene analysis 100–101, 108, 114 H-strand transcription (HSP) 141 human immunodeficiency virus (HIV) 52 Hyalinella punctata 64, 67–70, 73, 78, 99–103 Hydroides norvegica 51 Hypodontus macropi 185 IFN- 30 IgM 30 IL-2 29–30 IL-2R 29 IL-4 30 IL-5 30 IL-10 31 IL-12 31 inner membrane complex (IMC) 217, 247 inner membrane complex protein 1 (IMC1) and 2 (IMC2) 235 inositolphosphorylceramide (IPC) 26 internal transcribed spacers (ITS-1 and ITS-2) 164, 167–168, 185

268 intramembranous particles (IMPs) 217–218 Isospora belli 53 jasplakinolide 246 karyorelictid ciliates 107 Katharina tunicata 139–140 KT5926 234 Kudoa 58 Leishmania 3, 236 Leishmania adleri 16 Leptomonas samueli 29 Leptotheca 58 Leuciscus cephalus 51 lipopeptidophosphoglycan (LPPG) 4 lipophosphoglycan (LPG) 3–4 lipopolysaccharide (LPS) 30–31 Litomosoides sigmodontis 169 Loligo bleekeri 139 Lophopodella carteri 64 L-strand transcription (LSP) 141 Lumbricus terrstris 139 macrophages 30 Macropus fuliginosus 186 Macropus giganteus 186 Malacosporea 46, 115 diagnostic characters 62–94 Manayunkia speciosa 51 mass spectrometry (MS) 7 Mazmastrongylus odocoilei 170 Meloidogyne 166 Meloidogyne arenaria 155, 166 Meloidogyne chitwoodi 166 Meloidogyne hapla 155, 166 Meloidogyne hispanica 166 Meloidogyne incognita 155, 166 Meloidogyne javanica 153, 155, 166, 181 Metastrongylus pudendotectus 158 metazoa, occurrence of extrusible organelles 104 methionine 143

INDEX methylation analysis 14 microneme protein 2 (MIC2) 219–220 microtubule-associated motor proteins 241–242 microtubules 235–236 mitochondria 136 mitochondrial associated proteins (MAPs) 235, 240 mitochondrial DNA analysis 166, 174 mitochondrial DNA-encoded subunits 149 mitochondrial DNA sequence data 168 mitochondrial DNA sequences 149 mitochondrial DNA turnover 148 mitochondrial gene markers 175–179 for molecular systematics and population genetics of parasitic nematodes 163–179 mitochondrial gene sequences, systematics studies using 166–169 mitochondrial genomes flatworms 150–151 nematodes 151–155 parasitic helminths 150–163 significance of studying 147–150 mitochondrial genomics 133–211 animals 136–150 evolutionary implications 159–163 gene arrangement 138–140 general and historical perspective 136–138 genetic and genomic studies 147–148 genetic code 142–144 non-coding regions 146–147 protein genes 142–144 replication and transcription processes 140–142 sequencing approaches and recent technological improvements 155–159 structural and functional aspects 138–147 transfer and ribosomal RNA genes 144–146

INDEX mitochondrial organizing centres (MTOCs) 217, 236 mitochondrial transfer RNA 145 motility, external proteins implicated in 219–221 motor and cytoskeletal proteins 221–242 MREI 238 mutation scanning analysis, population genetics by 175–179 mutations-disease 148–149 myoA tail domain interacting protein (MTIP) 233–234 myosin binding proteins 233–235 myosins, class VI 229 Mytilus edulis 139–140, 142, 144 Myxidium 60, 99 Myxidium fugu 55 Myxidium leei 54 Myxidium lieberkuehni 100 Myxidium trachinorum 55–56 Myxobolus cerebralis 45, 50–51, 58, 61, 80 Myxobolus pendula 57, 116 Myxobolus sp. 52 Myxosporea 115–116 diagnostic characters 54–62 life cycle 54–62 myxosporean phase, characters of 55–60 Myxozoa 16S rDNA sequences 105, 114 18S rDNA phylogeny 95–99 ancestral body form 101 biodiversity and evolution 43–134 biology 47–49 candidate symbionts for nematocysts and polar capsules 105–108 classification 46–47 comparison of spores 51 convergent evolution 108 diagnostic characters 47–54 early classifications 94–95

269 endosymbiont acquisition of polar capsules and nematocysts 102–108 gene transfer 109 homology of polar capsules and nematocysts 109–111 hosts invertebrates 49–52 vertebrates 52–54 inferred higher level phylogeny 94–101 life cycles 46–49 lineages and evolution of parasitism 112–117 morphological simplification due to parasitism 113–117 multicellularity of spores 45 multiple acquisition and incorporation of endosymbionts 102–103 non-symbiotic origins of polar capsules and nematocysts 108–111 overview 44–46 phylogenetic relationships 94–111 polar capsules and Cnidarian nematocysts 102–111 Necator 176 Necator americanus 158–159, 161–162, 165, 171, 176–177, 179–180, 184, 186 Necator caninum 231 negative fast ion fast atom bombardment/mass spectrometry 24 Nematoda, overview 134–136 nematodes evolutionary implications 159–163 mitochondrial gene markers for molecular systematics and population genetics 163–179 mitochondrial genomes 151–155

270 sequencing approaches of mitochondrial genomes from 155–159 species for which complete, nearcomplete or partial mitochondrial genome sequences are available 152 species identification and differentiation 164–166 Neospora 242 Nephrostoma 51 Nereis diversicolor 51 NFAT-1 29 NK cells 30 nuclear genome of animals 143 nuclear magnetic resonance (NMR) 7 nuclear magnetic resonance spectroscopy 17–19 nuclear overhauser enhancements (NOEs) 17 nuclear small subunit ribosomal gene (nrSSU) sequence data 161 Onchocerca 169 Onchocerca volvulus 151, 159, 161, 163, 171–172, 179–180 Oncorhynchus mykiss 85 Ostertagia ostertagi 169–170 Paragonimus 164 Paramacropostrongylus iugalus 186 Paramacropostrongylus typicus 186 Parascaris equorum 158 parasitic helminths, mitochondrial genomes 150–163 Pectinatella magnifica 82, 87, 90 Pfmyo-A 230–232 Pfmyo-B 230 Pfmyo-C 230–231 Pfmyo-D 230–231 phosphatidylinositol 4,5-biphosphate 224 phosphatidylinositol glycans see PI-glycans

INDEX phosphatidylinositol-specific phospholipase C (PIPLC) 11 phosphoinositol glycans 19 Photorhabdus 168 PI-glycans 12–13, 15–18 structure of 6 PKD see proliferative kidney disease (PKD) PKX organism 80–94 Placoconus 176 Plasmodium 214, 217–218, 220–221, 227, 242–244 Plasmodium berghei 219 Plasmodium falciparum 103, 215, 218–219, 221–224, 226, 230–231, 237, 240–242, 248–249 Plasmodium yoellii 233–234, 241 Platichthys flesus 181 Platynereis dumerilii 139 Plumatella emarginata 82, 85–87, 89 Plumatella fungosa 65–66, 72–74, 78 Plumatella repens 65–67, 72, 92 Plumatella rugosa 82, 87 Podocoryne carneae 110 polyclonal B lymphocyte 30 Polykrikos 105 polymerase chain reaction (PCR) 91–92, 156–158, 165–166, 168, 172–174, 184, 231 Polypodium 96 Polypodium hydriforme 96–100 population genetics 135, 169–173 by mutation scanning analysis 175–179 technological considerations 173–175 positive ion fast atom bombardment/ mass spectrometry 16 proliferative kidney disease (PKD) 44, 80–94 Pupa strigosa 139 regulatory proteins 233–235 restriction fragment length polymorphism (RFLP) 157, 165, 169–170, 172–173, 184

INDEX Rhabdonema 114 ribosomal DNA (rDNA) 164 ribosomal RNA (rRNA) 137–138, 144–146 RNA components 136, 146 RNA editing 148 RNA genes 136 RNA synthesis 141–142 Romanomermis culicivorax 153, 155 Saccharomyces cerevisae 26–28 Saccosporidae 46 Salmo gairdneri 84 Sarcocystis 217, 248 Sarcocystis muris 231 Schistosoma 164 Schistosoma japonicum 151, 175 Schistosoma mansoni 140, 151 Scopthalmus maximus 55 Selenidium 248 Semotilus atromaculatus 57 serine-palmitoyl transferase (SPT) 26 Serrasalmus striolatus 56 single-strand conformation polymorphism (SSCP) 174–175, 177, 184 sodium ortho-vanadate 249 Sparus auratus 54 Sphaerospora 48, 58 Sphaerospora renicola 58, 79, 116 sphinganine, N-acetyl-O-trimethylsilyl derivatives of 21 sphingosine, N-acetyl-O-trimethylsilyl derivatives of 21 Spirorbis spirorbis 51 Stolella evelinae 64 Strongyloides stercoralis 154, 158–159, 161, 163, 179–180 Strongylus vulgaris 158 systematics studies using mitochondrial gene sequences 166–169 Taenia 164 Takifugu rubripes 55

271 Talpa europaea 52–53 TEDS rule 229–230 Teladorsagia boreoarcticus 166 Teladorsagia circumcincta 166, 170 Teladorsagia trifurcata 166 Tetracapsula 46 Tetracapsula bryosalmonae 62, 89, 115 Tetracapsula bryozoides 62, 66, 72–74, 79–81 Tetracapsula renicola 63, 81 Tetracapsuloides bryosalmonae 51, 62–63, 65–67, 73, 80–95, 112, 117 differentiation of sac-like stages 94 geographical distribution 82–83 hosts 82 morphology and development in Bryozoa 87–89 pathogenicity and development in fish 83–86 prevalence in Bryozoa 90 taxonomic history 80–94 transmission of 90–93 Tetractinomyxon intermedium 51 Tetractinomyxon irregulare 51 Tetrahymena 222, 239 Tetraspora discoidea 61 Tetraspora rotundum 61 TgIMC1 235 TgMLC1 234 Tgmyo-A 227, 231–234 Tgmyo-B 229, 232 Tgmyo-C 229, 232 Tgmyo-D 232 Tgmyo-E 232 Theileria 214–215 thrombospondin (TSP) 220, 245 TNF- 31 Tortrix viridana 51 Toxoplasma 214, 217, 219, 221, 223–224, 226–227, 230–235, 242–243, 247 Toxoplasma gondii 215, 218–219, 222, 224–225, 227, 231, 233, 237–238, 244–247 transfer RNA genes 137

272 Trichinella 165 Trichinella spiralis 151, 153–155, 159, 162, 179 Trichinella zimbabwensis 165 triploblasts 111 Trypanosoma brucei 27, 239 Trypanosoma cruzi epimastigotes 27 glycoinositolphospolipid from 1–41 GIPL 3 and host immune responses 28–31 characterization of lipid moiety 20–24 characterization of PI-glycan 12–20 chemical composition 9–10 GC/MS characterization of lipid components 20–22 lipid composition 11 MS characterization of intact lipid components 22–24 sample preparation 11–12 structural characterization of 7–24 GPI biosynthesis in 24–28 ICP synthase 27

INDEX Tubifex sp. 51 Tubifex tubifex 61 tubulin 235–236 -tubulin acetylation 239–240 autoregulation 238 post-translational modification 239–241 tyrosination 240–241 -tubulins 238–239 Uncinaria 176 Unicapsula 48, 58 Venerupis philippinarum 139, 142 Wolbachia 168–169 Wuchereria 169 yeast 229 Zell-Linean Z 69 Zschokkella 60 Zschokkella pleomorpha 55

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