Fish Parasites: A Handbook of Protocols for their Isolation, Culture and Transmission 178918133X, 9781789181333

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FISH PARASITES

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FISH PARASITES

A Handbook of Protocols for their

Isolation, Culture and Transmission

European Association of Fish Pathologists (EAFP) / 5m Books Series

Edited by

Prof. Ariadna Sitjà-Bobadilla

Prof. James. E. Bron

Prof. Geert Wiegertjes

Dr. M. Carla Piazzon

Series Editor

Sean Monaghan

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First published 2021 Copyright © 5m Books 2021 Chapters 4, 21 and 23 are published under a cc-by license. All rights reserved. No part of this publication may be reproduced, stored in

a retrieval system, or transmitted, in any form or by any means, electronic,

mechanical, photocopying, recording or otherwise, without prior permission

of the copyright holder.

Published by

5M Books Ltd,

Lings, Great Easton,

Essex CM6 2HH, UK,

Tel: +44 (0)330 1333 580

www.5mbooks.com

A Catalogue record for this book is available from the Briwtish Library

ISBN 9781789181333

eISBN 9781789181531

DOI 10.52517.9781789181531

Book layout by Servis Filmsetting Ltd, Stockport, Cheshire

Printed by Replika Press Pvt Ltd, India

Photos by the authors unless otherwise indicated

Front cover images: Sea Lice Research Centre, University of Bergen, Norway;

Barthlomew Lab; Ivona Mladineo; Joana Pimentel-Santos

Back cover images: David B. Vaughan; Paola Beraldo; Csaba Székely

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Contents

Contributors Preface and acknowledgements 1 Cryptocaryon irritans David B. Vaughan and Kate S. Hutson 2 Ichthyophthirius multifiliis Kurt Buchmann and Louise von Gersdorff Jørgensen 3 Philasterides dicentrarchi Jesús Lamas and José Manuel Leiro 4 Amyloodinium ocellatum Paola Beraldo, Michela Massimo and Marco Galeotti 5 Trypanosoma and Trypanoplasma Geert F. Wiegertjes, Maria Forlenza and Simon R. M. Jones 6 Neoparamoeba perurans James E. Bron, Irene Cano, Carolina Fernández, Sophie Fridman, Richard Paley and Jadwiga Sokolowska 7 Loma salmonae Simon R. M. Jones and Sarah McConnachie 8 Ceratonova shasta Jerri L. Bartholomew, Stephen D. Atkinson, Sascha L. Hallett and Julie D. Alexander

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x xvii 1 17 33 49 63

78 95 110

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vi ♦  Fish Parasites

9 Enteromyxum leei Ariadna Sitjà-Bobadilla, Itziar Estensoro and Oswaldo Palenzuela 10 Enteromyxum scophthalmi Oswaldo Palenzuela, M. Carla Piazzon and Ariadna Sitjà-Bobadilla 11 Myxobolus cerebralis Jerri L. Bartholomew, Stephen D. Atkinson, Sascha L. Hallett and Julie D. Alexander 12 Sphaerospora molnari Astrid S. Holzer and Joana Pimentel-Santos 13 Tetracapsuloides bryosalmonae Bartolomeo Gorgoglione and Mohamed H. Kotob 14 Leptorhynchoides species (Acanthocephala) Michael A. Barger and Florian B. Reyda 15 Ligula intestinalis David Hoole 16 Zeylanicobdella arugamensis David B. Vaughan and Kate S. Hutson 17 Neobenedenia girellae Kate S. Hutson, David B. Vaughan, Alexander K. Brazenor and Alejandro Trujillo-González 18 Zeuxapta seriolae Kate S. Hutson, Allan J. Mooney and David B. Vaughan 19 Anguillicoloides crassus Csaba Székely, Olga Haenen and Kálmán Molnár 20 Anisakis simplex Ivona Mladineo 21 Hysterothylacium aduncum Francisco Javier Adroher-Auroux and Rocío Benítez­ Rodríguez 22 Caligus rogercresseyi and Caligid sea lice species Sandra L. Marín and Mark D. Fast 23 Lepeophtheirus salmonis Lars Are Hamre and Linda Andersen

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127 143 157 174 189 210 227 242 258 272 286 298 311 330 351

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Contents ♦  vii

24 Ceratothoa oestroides Ivona Mladineo 25 Gnathia aureamaculosa and Gnathia marleyi Alexandra S. Grutter, William E. Feeney, Eva C. McClure, Pauline Narvaez, Nico J. Smit, Derek Sun, Paul C. Sikkel and Kate S. Hutson 26 Saprolegnia parasitica Kostis Apostolakis, Christopher J. Secombes and Pieter van West Index

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365

375 390 402

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Aquatic Animal Diseases Series EAFP book series editor Dr Sean Monaghan, EAFP Publications Officer, University of Stirling, Scotland, UK [email protected]

5m Books Nigel Balmforth, Publisher and Director, Aquatic Sciences, 5m Books, UK [email protected]

When culturing aquatic species, the proper maintenance of health is essential to the sustainability of production. Disease epidemics, for example outbreaks of infectious salmon anaemia (ISA) in Chile, and early mortality syndrome (EMS) in shrimp in South-East Asia have caused widespread and devastating effects on the aquaculture industries in the affected countries. In order for aquatic animal diseases to be properly prevented, identified, monitored, treated and managed, there is a constant need for high quality and authoritative books to inform the range of personnel working in the aquaculture industry. The new EAFP/5m Books Series encompasses professional-level reference books and practical guides aimed at fish farm operatives and technicians, fish biologists and pathologists, veterinarians, students who are studying aquaculture and fish health, and the many personnel involved in supplying the aquaculture industry with pharmaceuticals, nutraceuticals, feed and equipment.

Aquatic Animal Diseases Series ♦  ix

The first tiles in the series include: Ariadna Sitjà-Bobadilla, James E. Bron, Geert F. Wiegertjes and M. Carla Piazzon (eds) (2021) Fish Parasites: A Handbook of Protocols for their Isolation, Culture and Transmission Albert Tacon (2022) A Handbook of Nutritional Disorders and Diseases of Fish and Shrimp Bartolomeo Gorgoglione and Christyn Bailey (eds) (2022) Proliferative Kidney Disease

The European Association of Fish Pathologists (EAFP) exists to promote the exchange of knowledge and coordination of research related to fish and shellfish pathology. Membership of the EAFP is open to all fish and shellfish pathologists, students of fish pathology and organisations, agencies or other associations which have an interest in maintaining the objectives of the EAFP. https://eafp.org 5m Books publishes professional-level books for people and businesses in aqua­ culture, fisheries, agriculture, veterinary sciences and the global food chain. www.5mbooks.com

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Contributors

Francisco Javier Adroher-Auroux – Departamento de Parasitología, Facultad de Farmacia, Universidad de Granada, 18071 Granada, Spain; e-mail: [email protected] Julie D. Alexander – Department of Microbiology, Oregon State University, Nash Hall 226, Corvallis, OR, USA; e-mail: julie.alex [email protected] Linda Andersen – The Aquatic and Industrial Laboratory (ILAB), Thormøhlensgate 55, 5006 Bergen, Norway; e-mail: linda.ander [email protected] Kostis Apostolakis – International Centre for Aquaculture Research and Development, Aberdeen Oomycete Laboratory, Institute of Medical sciences, University of Aberdeen, Aberdeen AB25 2ZD, Scotland, UK; Scottish Fish Immunology Research Centre, School of Biological Sciences, University of Aberdeen, Aberdeen AB24 2TZ, Scotland, UK Stephen D. Atkinson – Department of Microbiology, Oregon State University, Nash Hall 226, Corvallis, OR, USA; e-mail: stephen. [email protected] Michael A. Barger – Department of Biology and Health Sciences, Stephens College, Columbia, MO 65215, USA; e-mail: mallen [email protected]

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Contributors ♦  xi

Jerri L. Bartholomew – Department of Microbiology, Oregon State University, Nash Hall 226, Corvallis, OR, USA; e-mail: jerri. [email protected] Rocío Benítez-Rodríguez – Departamento de Parasitología, Facultad de Farmacia, Universidad de Granada, 18071 Granada, Spain; e-mail: [email protected] Paola Beraldo – Department of Agricultural, Food, Environmental and Animal Sciences (Di4A) – Section of Animal and Veterinary Sciences – University of Udine; e-mail: [email protected] Alexander K. Brazenor – Biosecurity Animal Division, Australian Government Department of Agriculture, 18 Marcus Clarke Street Canberra, Australian Capital Territory, Australia; e-mail: Alex. [email protected] James E. Bron – Institute of Aquaculture, University of Stirling, Stirling FK9 4LA, UK; e-mail: [email protected] Kurt Buchmann – University of Copenhagen, Faculty of Health and Medical Sciences, Department of Veterinary and Animal Science, Laboratory of Aquatic Pathobiology, Frederiksberg C., Denmark; e-mail: [email protected] Irene Cano – Centre for Environment, Fisheries and Aquaculture Science (CEFAS), The Nothe, Barrack Road, Weymouth, Dorset DT4 8UB, UK; e-mail: [email protected] Itziar Estensoro – Fish Pathology Group, Instituto de Acuicultura Torre de la Sal, Consejo Superior de Investigaciones Científicas (IATS-CSIC), 12595 Ribera de Cabanes, Castellón, Spain; e-mail: [email protected] Mark D. Fast – Atlantic Veterinary College, University of Prince Edward Island, Charlottetown, PEI, Canada; e-mail: mfast@ upei.ca William E. Feeney – Environmental Futures Research Institute, Griffith University, Nathan, QLD 4111, Australia; e-mail: will [email protected]

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Carolina Fernández – Institute of Aquaculture, University of Stirling, Stirling FK9 4LA, UK; e-mail: [email protected] Sophie Fridman – Institute of Aquaculture, University of Stirling, Stirling FK9 4LA, UK; e-mail: [email protected] Maria Forlenza – Aquaculture and Fisheries group, Department of Animal Sciences, Wageningen University & Research, The Netherlands; e-mail: [email protected] Marco Galeotti – Department of Agricultural, Food, Environmental and Animal Sciences (Di4A) – Section of Animal and Veterinary Sciences – University of Udine; e-mail: [email protected] Bartolomeo Gorgoglione – Aquatic Animal Health Laboratory, Dept. of Pathobiology and Diagnostic Investigation, CVM & Dept. of Fisheries and Wildlife, CANR – Michigan State University, East Lansing, Michigan, USA; e-mail: [email protected] Alexandra S. Grutter – School of Biological Sciences, The University of Queensland, St Lucia, QLD 4072, Australia; e-mails: a.grutter@ uq.edu.au Olga Haenen – Wageningen Bioveterinary Research, Fish Diseases Laboratory, P.O. Box 65, 8200 AB Lelystad, The Netherlands; e-mail: [email protected] Sascha L. Hallett – Department of Microbiology, Oregon State University, Nash Hall 226, Corvallis, OR, USA; e-mail: sascha. [email protected] Lars Are Hamre – SLRC – Sea Lice Research Centre, Department of Biological Sciences, University of Bergen, Postbox 7803, 5020 Bergen, Norway; e-mail: [email protected] Astrid S. Holzer – Institute of Parasitology, Biology Centre, Czech Academy of Science, České Budějovice, Czechia; e-mail: astrid. [email protected] David Hoole – School of Life Sciences, Huxley Building, Keele University, Keele, Staffordshire, ST5 5BG, UK; e-mail d.hoole@ keele.ac.uk

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Contributors ♦  xiii

Kate S. Hutson – Marine Parasitology Laboratory, Centre for Sustainable Fisheries and Aquaculture, College of Science and Engineering, James Cook University, 1 University Drive, Townsville, Queensland 4814, Australia; Cawthron Institute, 98 Halifax Street East, Nelson 7010, New Zealand; e-mail: kate.hutson@cawthron. org.nz Simon R. M. Jones – Fisheries and Oceans, 3190 Hammond Bay Road, V9T 6N7 Nanaimo, British Columbia, Canada; e-mail: [email protected] Mohamed H. Kotob – Dept. of Pharmaceutical Chemistry, University of Vienna, Vienna, Austria; Dept. of Pathology, Faculty of Veterinary Medicine, Assiut University, Assiut, Egypt Jesús Lamas – Department of Functional Biology, Cell Biology, CIBUS, Institute of Aquaculture, Campus Vida, University of Santiago de Compostela, E-15782, Santiago de Compostela, Spain; e-mail: [email protected] José Manuel Leiro – Department of Microbiology and Parasitology, Laboratory of Parasitology, Institute of Research and Food Analysis, Campus Vida, University of Santiago de Compostela, E-15782, Santiago de Compostela, Spain. e-mail: josemanuel. [email protected] Sandra L. Marín – Instituto de Acuicultura, Universidad Austral de Chile, Sede Puerto Montt, Chile; Centro FONDAP de Investigación en Dinámica de Ecosistemas Marinos de Altas Latitudes (IDEAL), Universidad Austral de Chile, Puerto Montt, Chile; e-mail: smarin@ uach.cl Michela Massimo – Department of Agricultural, Food, Environmental and Animal Sciences (Di4A) – Section of Animal and Veterinary Sciences – University of Udine; e-mail: michela.mas [email protected] Eva C. McClure – School of Biological Sciences, The University of Queensland, St Lucia, QLD 4072, Australia; Present address: Australian Rivers Institute, Griffith University, Parklands Dr,

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Southport, Queensland 4215, Australia; e-mail: e.mcclure@griffith. edu.au Sarah McConnachie – Ottawa Animal Emergency and Specialty Hospital 203-1155 Lola St, Ottawa, ON K1K 4C1, Canada Ivona Mladineo – Institute of Oceanography and Fisheries, Split, Croatia; e-mail: [email protected] Kálmán Molnár – Institute for Veterinary Medical Research, Centre for Agricultural Research, 1143. Budapest. Hungária krt. 21, Hungary; e-mail: [email protected] Allan J. Mooney – Independent researcher; e-mail: mooney.allan@ gmail.com Pauline Narvaez – Centre for Sustainable Fisheries and Aquaculture, College of Science and Engineering, James Cook University, 1 University Drive, Townsville, Australia; ARC Centre of Excellence for Coral Reef Studies, James Cook University, 1 James Cook Drive, Townsville, Queensland 4810, Australia; e-mail: pauline.narvaez@ my.jcu.edu.au Oswaldo Palenzuela – Fish Pathology Group, Instituto de Acuicultura Torre de la Sal, Consejo Superior de Investigaciones Científicas (IATS-CSIC), 12595 Ribera de Cabanes, Castellón, Spain; e-mail: [email protected] Richard Paley – Centre for Environment, Fisheries and Aquaculture  Science (CEFAS), The Nothe, Barrack Road, Weymouth, Dorset DT4 8UB, UK; e-mail: richard.paley@cefas. co.uk M. Carla Piazzon – Fish Pathology Group, Department of Marine Species Biology, Culture and Pathology, Instituto de Acuicultura Torre de la Sal, Consejo Superior de Investigaciones Científicas (IATS-CSIC), Spain; e-mail: [email protected] Joana Pimentel-Santos – Institute of Parasitology, Biology Centre, Czech Academy of Science, České Budějovice, Czechia

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Contributors ♦  xv

Florian B. Reyda – Biology Department & Biological Field Station, State University of New York, College at Oneonta, Ravine Parkway Oneonta, NY, USA; e-mail: [email protected] Christopher J. Secombes – Scottish Fish Immunology Research Centre, School of Biological Sciences, University of Aberdeen, Aberdeen AB24 2TZ, Scotland, UK Paul C. Sikkel – Water Research Group, Unit for Environmental Sciences and Management, North-West University, Potchefstroom 2520, South Africa; Department of Biological Sciences and Environmental Sciences Program, Arkansas State University, State University, AR 72467, USA; e-mail: [email protected] Ariadna Sitjà-Bobadilla – Fish Pathology Group, Instituto de Acuicultura Torre de la Sal, Consejo Superior de Investigaciones Científicas (IATS-CSIC), 12595 Ribera de Cabanes, Castellón, Spain; e-mail: [email protected] Nico J. Smit – Water Research Group, Unit for Environmental Sciences and Management, North-West University, Potchefstroom 2520, South Africa; e-mail: [email protected] Jadwiga Sokolowska – Institute of Aquaculture, University of Stirling, Stirling FK9 4LA, UK; e-mail: [email protected] Derek Sun – School of Biological Sciences, The University of Queensland, St Lucia, Queensland 4072, Australia; e-mail: derek. [email protected] Csaba Székely – Institute for Veterinary Medical Research, Centre for Agricultural Research, 1143. Budapest. Hungária krt. 21. Hungary; e-mail: [email protected] Alejandro Trujillo-González – Institute for Applied Ecology, University of Canberra, 11 Kirinari street, Bruce, Canberra, Australian Capital Territory, Australia; e-mail: alejandro.trujillo [email protected] David B. Vaughan – School of Access Education, Central Queensland University, 554–700 Yaamba Road, Rockhampton,

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xvi ♦  Fish Parasites

Queensland 4701, Australia; Marine Parasitology Laboratory, College of Science and Engineering, James Cook University, 1 University Drive, Townsville, Queensland 4814, Australia; e-mail: [email protected] Pieter van West – International Centre for Aquaculture Research and Development, Aberdeen Oomycete Laboratory, Institute of Medical sciences, University of Aberdeen, Aberdeen AB25 2ZD, Scotland, UK; e-mail: [email protected] Louise von Gersdorff Jørgensen – University of Copenhagen, Faculty of Health and Medical Sciences, Department of Veterinary and Animal Science, Laboratory of Aquatic Pathobiology, Frederiksberg C., Denmark Geert F. Wiegertjes – Aquaculture and Fisheries group, Department of Animal Sciences, Wageningen University & Research, The Netherlands; e-mail: [email protected]

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Preface and acknowledgements

As the field of parasitology continues to expand its horizons and capabilities, so does global aquaculture. Finfish aquaculture now surpasses wild fisheries in its capacity to provide food for an expand­ ing human population and also serves to support a flourishing aquarium trade and conserve wild populations. Aquaculture intensi­ fication, coupled with changing climatic conditions, has exacerbated some parasitic diseases, generating new challenges for diagnosis and treatment, and causing considerable economic losses. The prevailing situation demands a better understanding of para­ sitic disease in aquaculture and requires more tools for management and control than currently exist. However, this is a difficult task, one that starts with three nominally simple, yet challenging, steps: the isolation of the infectious agent, the development of suitable in vitro culture procedures and the experimental transmission of the para­ site. These three steps are indispensable for further characterization and investigation of targeted parasites, ranging from the morpho­ logical or molecular level, to the level of host–parasite interactions. These steps are also increasingly important to the provision of pre­ cious transcriptomic and genomic data, allowing the development of new research tools and treatments and the identification of novel vaccine targets. In addition, these three steps are profoundly inter­ connected. While establishing a transmission model in the labora­ tory is often an intricate process, most of the difficulties relate back to the initial isolation of parasites and their long-term maintenance, either in vitro or in vivo.

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xviii ♦  Fish Parasites

This book offers step-by-step procedures for isolating these para­ sites and reproducing parasitic infections under laboratory condi­ tions. It covers different endo- and ectoparasites drawn from diverse taxonomic groups, both protists and metazoans (ciliates, amoebae, microsporidians, flagellates, myxozoans, helminths, crustaceans and oomycetes). Most of the parasites we describe can be transmitted directly from fish to fish, with no need for intermediates or vectors, however, others require specific interactions with several hosts. For some parasites in vitro culture has been achieved, for others in vivo continuous passage in fish is needed. With this book, we wanted readers to find information quickly and easily, so that for each para­ site chapter we have provided an initial summary, including details of the life cycle and the disease invoked, followed by information on the facilities, equipment and reagents needed for parasite iso­ lation, culture and transmission and a set of standard operating procedures (SOPs) with summary templates. The target audience for this volume comprises not only scholars and students, but also researchers from a range of backgrounds who wish to work with fish parasites, but have been reluctant to do so, due to the inherent difficulties involved in starting the process. The inspiration leading to this book’s inception was provided by the extensive work, involving 28 institutions from 13 coun­ tries, conducted over a period of 5 years within ParaFishControl, a research project funded by the European Union’s Horizon 2020 framework. European specialists in fish parasitology conceived the content of this book, but its remit soon broadened and was greatly enhanced by the contributions of wider international collaborators. This book therefore embodies many of the best aspects of the inter­ disciplinary, collaborative nature of parasitology, a science that gath­ ers the expertise of veterinarians, biologists, pharmacists, chemists, epidemiologists and many more. Our blended, interconnected work was concluded during the COVID-19 pandemic, a stark reminder of the capacity of infectious disease to cause extensive harm. Despite the challenging times, we succeeded in completing this book in order to increase interest in, sate inherent scientific curiosity about and advance research relating to parasitic diseases worldwide. It took us many years, often decades,

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Preface and acknowledgements ♦  xix

to develop and master some of the procedures gathered in this book and we therefore hope that this compilation empowers a new gen­ eration of researchers to tackle the many fascinating problems in fish parasitology that remain unsolved. We wish to thank 5m Books for believing in our idea and to thank all the authors that have followed it and contributed to make this book a reality. We would like to extend our special thanks to Miranda Maybank for editing the text. Finally, we would like to express our gratitude to the international volunteer external reviewers of the individual chapters for their expert and constructive criticism: Eugene M. Burreson (Virginia Institute of Marine Science, USA); Javier Diéguez (Real Jardín Botánico, CSIC, Spain); Edit Eszterbauer (Hungarian Academy of Science, Hungary); Stephen Feist (Centre for Environmental, Fisheries and Aquaculture Science, UK); Raul Iglesias (Universidad de Vigo, Spain); Egil Karlsbakk (University of Bergen, Norway); Arne Levsen (Institute of Marine Research, Norway); Francisco Montero (University of Valencia, Spain); Hiroshi Yokoyama (Okoyama University of Science, Japan); and Carlos Zarza (Skretting Aquaculture Research Centre, Norway).

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Chapter 1

Cryptocaryon irritans David B. Vaughan1,2 and Kate S. Hutson2,3 1

School of Access Education, Central Queensland University,

554–700 Yaamba Road, Rockhampton, Queensland 4701, Australia;

e-mail: [email protected]

2 Marine Parasitology Laboratory, College of Science and Engineering, James

Cook University, 1 University Drive, Townsville, Queensland 4814, Australia

3 Cawthron Institute, 98 Halifax Street East, Nelson 7010, New Zealand;

e-mail: [email protected]

1. Background

C

ryptocaryon irritans is a unicellular ciliated protozoan that is an obligate parasite of marine bony fishes. Globally, this species is responsible for marine white spot disease in fishes in aquaculture, and in home and public aquaria. Clinical signs of the disease include visible white nodules on the skin and fins of infected host fishes (Fig. 1.1), increased or laboured breathing, inappetence and general

Fig. 1.1 Juvenile barramundi (Lates calcarifer) infected with Cryptocaryon irritans. Mature trophonts are visible as white nodules. Credit: Pauline Narvaez, James Cook University.

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2 ♦  Fish Parasites

lethargy. Marine white spot disease of fishes is similar to freshwa­ ter white spot disease, caused by a separate ciliated protozoan para­ site, Ichthyophthirius multifiliis. Neither of these infections should be confused with white spot disease of crustaceans, which is caused by the white spot syndrome virus. Cryptocaryon irritans exhibits a very low host specificity, which makes it particularly problematic for fish farmers. It can kill juvenile fishes and decrease the growth performance of larger fishes in culture. The parasite has a direct life cycle (Fig. 1.2), requiring only a single host, and typically includes

Fig. 1.2 Life cycle of Cryptocaryon irritans. Host fish with mature ciliated trophonts (a) present under the epidermis of the body, fins and gill lamellae. After vacating the host fish, pro-tomonts (b) enter the environment and form an encysted tomont (c). Free-swimming theronts (d) actively seek out a new host. Credit: Eden Cartwright, Bird Circus.

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Cryptocaryon irritans ♦  3

four stages: the parasitic stage or trophont (Fig. 1.2a) that infects the host and lives beneath the epidermal skin layer; the pro-tomont (vacated from host; Fig. 1.2b); the reproductive tomont stage, which undergoes asexual reproduction in the immediate environment (Fig. 1.2c); and the theront stage (Fig. 1.2d), which actively seeks out a new host to infect in the water column (Colorni and Burgess, 1997). Large numbers of theronts can be produced per tomont, with larger tomonts producing the highest numbers (Colorni and Burgess, 1997). Under normal host conditions, mature trophonts emerge through the epidermis during darkness (Burgess and Matthews, 1994) and vacate the host. If the host dies, the trophonts will also be triggered to vacate host tissue and will often be smaller, although many will have the capacity to survive. Trophonts that have vacated the host, sometimes referred to as pro-tomonts, fall to the benthos where they continue to move around for a short period of time, seeking out crevices in the substrate or between small stones or sand, where they encyst to form true tomonts. This encystment provides considerable protection as the cells undergo asexual reproduction though binary fission, forming multiple tomite divisions. Under laboratory condi­ tions the tomonts often clump together to form large aggregations (Colorni and Burgess, 1997; Fig. 1.3). Theronts are released from the cysts in the early hours of the morning (Diggles and Lester, 1996; Skilton et al., 2020), before sunrise. Using a dissection microscope, theronts can be observed within the tomonts becoming increasingly active just prior to excystment. This increased activity is likely to

Fig. 1.3 Aggregation of Cryptocaryon irritans tomonts in culture. Scale bar, 400 µm. Credit: David B. Vaughan.

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4 ♦  Fish Parasites

cause the cyst wall to break open, releasing the theronts into the water column. Theronts are active swimmers and have a limited time in which to find a susceptible host. Usually theronts can survive without a host for 24 h, but successful infectivity decreases 6–8 h after excystment (Colorni and Burgess, 1997). Once theronts locate the skin of a suitable host, they can invade the epidermis in as little as 30 seconds (Diggles and Lester, 1996). It is important for prospective aqua­ culturists to understand that the influence of temperature (on the completion of the life cycle), as well as the size and number of ther­ onts produced per tomont, can vary between isolates, geographic localities and host species cultured. Theronts also exhibit a level of emergence asynchrony. It is therefore advisable to observe all stages carefully under a dissection microscope when preparing a culture for controlled laboratory experiments. Culture isolate identity should be confirmed, and a representative molecular sequence deposited into a public database, such as GenBank.

2. General facilities and fish All continuous cultures of C. irritans should be preceded by the appropriate institutional animal ethics approvals and necessary per­ mits, according to country-specific legislation. Laboratory facilities housing cultures of C. irritans should be strictly access controlled, with dedicated staff, and should follow strict biosecurity protocols to reduce the risk of transmission or parasite escape through waste seawater. Chlorination of equipment and waste seawater, using 50 mg/l sodium hypochlorite, is simple and effective (Vaughan et al., 2018). All culture systems must be strictly isolated from any other seawater source or aquaculture facility, as the parasite is easily transmitted in the water and with untreated fomites. 2.1. In vivo culture Fish species used for culture purposes should ideally be euryhaline, to facilitate salinity manipulations of the host’s captive conditions

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Cryptocaryon irritans ♦  5

by acclimation, both within and outside of the parasite’s tolerance range. This accommodates the use of fish for culture of the parasite in seawater, with a host recovery period in freshwater. Cryptocaryon irritans cannot survive and reproduce at salinities below 10‰. Some mullet species, mollies (Poecilia sphenops) and barramundi (Lates calcarifer) are likely host candidates, the latter being a particularly effective host species which can be managed as a small population (e.g. Skilton et al., 2020; Vaughan et al., 2018). While other non­ euryhaline marine species can be used, unless their post-infection care includes a rigorous active treatment regime and a quarantine isolation period, the most humane end-point is euthanasia by anaes­ thetic overdose. Host fish may acquire limited immunity after an initial infection with C. irritans. This immunity does not confer complete resistance, but reduces the intensity of subsequent infections and may last for 6 months (see Burgess and Matthews, 1995). Therefore, the host population used for a C. irritans culture should be managed care­ fully to accommodate this potential immunity, and be large enough to provide a rest period for individual fish of at least 6 months prior to reuse. A low-intensity infection can be maintained in a set of two iden­ tical glass aquaria containing rudimentary life support and without any substrate, both controlled to the optimum temperature for the parasite. Two tanks are always maintained simultaneously, to avoid losing the infection in case of any technical failure in one of them. The bottom of the tanks should be painted black on the outside to facilitate easy detection of settled tomonts. Optimum temperature may vary per isolate, but is considered to be 24–26°C. Salinity should be maintained at 35‰. Cryptocaryon irritans is capable of producing in excess of 1000 theronts per tomont (Diggles and Lester, 1996). However, normal invasion success can range between 5 and 20% (Burgess and Matthews, 1995). Therefore, depending on the host size, we have concluded in our own continuous culture that the use of 10–20 viable tomonts is adequate for three individ­ ual fish in a single 45-l aquarium. This allows for the maintenance of an effective culture without adversely compromising the host fish.

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6 ♦  Fish Parasites

3. Equipment and reagents Basic: • • • • • • • • • • • • •

Standard PPE; disposable gloves and laboratory coat Standard plastic Pasteur pipettes (single use) Reusable glass Petri dishes and matching lids (6–12 cm) Standard glass microscope slides Dissection needles Automatic pipettes and pipette tips Haemocytometer Dissecting (light) microscope Hand tally counter Thermometer Refractometer 25-l buckets Permanent marker.

Aquaria (45-l glass with secure lids; base painted black on the outside): • Basic entry-level externally mounted power filter with standard media • Air tubes and stones • Air pumps • Submersible thermostatically controlled heaters • Total ammonia, nitrite and nitrate test kits. Reagents: • • • •

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Supply of fresh, pre-filtered seawater Distilled water Sodium hypochlorite 10% formalin.

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Cryptocaryon irritans ♦  7

4. Standard operating procedures (SOPs) for C. irritans in vivo culture 4.1. SOP for initiating in vivo culture Set up both culture aquaria at least a week prior to collecting para­ site material. Allow aquaria to operate without livestock at optimum temperature (24–26°C) and salinity (35‰). When initiating the culture, seek out an infected fish from a local ornamental fish sup­ plier or local fish farm with a known infection. If the fish is large enough, a simple gill biopsy is often sufficient to collect individual live trophonts in situ, the largest of which are visible to the naked eye (~450 µm), contrasted against the red pigment of the lamellae. Trophonts removed in this way can be transported back to the lab in a vial of fresh seawater. On arrival of this tissue in the laboratory: 1. 2. 3.

4.

5. 6. 7.

8. 9.

Place gill biopsy sample into a glass Petri dish containing fresh filtered seawater. Place the Petri dish under the dissection microscope and observe the area of tissue containing live trophonts. Using dissecting needles, carefully tease apart epidermal tissue surrounding each trophont, allowing them to release them­ selves from the surrounding tissue independently. Using a standard plastic Pasteur pipette or automatic pipette, aspirate individual trophonts to a separate 12-cm glass Petri dish containing fresh, filtered seawater and cover with match­ ing lid. Insert label with the contents, noting contents and date using waterproof paper and pencil labelling. Incubate at room temperature under natural lighting. After 24 h observe encysted tomonts in the Petri dish under the dissection microscope and remove 50% of the seawater with a plastic Pasteur pipette. Gently replace the removed seawater volume with fresh, fil­ tered seawater; repeat daily. Monitor for tomite divisions, which become visible after several days within each tomont. Under the dissection microscope,

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8 ♦  Fish Parasites

carefully dislodge 10–20 tomonts from the surface of the Petri dish using the blunt mouth of a Pasteur pipette and introduce to each awaiting culture aquarium directly, allowing them to settle on the benthic surface of the aquarium. 10. Introduce three pre-acclimated host fish to each aquarium. 4.2. SOP for perpetuating in vivo culture Within days of successful initial infection, host fish will display clin­ ical signs of infection that include visible white nodules on the skin and fins. When these signs become visible: 1. 2. 3.

4. 5. 6.

7.

8. 9.

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Cover the benthic surface of each aquarium with glass micro­ scope slides or small (6-cm) glass Petri dishes. Once visible tomonts are observed attached to microscope slides or Petri dishes, remove from each tank. Under the dissection microscope remove and collect about 100 of the largest tomonts to a 12-cm glass Petri dish containing fresh, filtered seawater. Cover with matching lid and label with contents and date using a permanent marker; incubate at room temperature and under natural lighting. Carefully remove host fish from culture tanks and acclimate to freshwater to recover. Turn off power to each aquarium, remove and clean heaters and filters. Use a sponge scourer to systematically remove any biofilm and any remaining tomonts from the aquarium glass surface area. Drain aquarium water to two 25-l buckets and treat waste sea­ water and cleaning equipment with 50 mg/l sodium hypochlo­ rite for 30 minutes, before disposing of treated water into an approved waste water system. Re-fill aquaria with fresh, filtered seawater and restore filters and heaters to normal functioning operation. Observe the development of the tomonts daily under the dis­ secting microscope.

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Cryptocaryon irritans ♦  9

Remove 10–20 mature tomonts with notable tomite divisions to each aquarium. 11. Introduce three naïve host fish to each tank. 12. Excess tomonts can be maintained as a back-up in labelled glass Petri dishes, replacing 50% of their seawater volume daily and treating any waste seawater as described above. They can also be utilized for study, or disposed of using the same sodium hypochlorite solution and exposure time. Glassware can be cleaned thereafter for reuse. Live tomonts should not be refrig­ erated, as refrigeration significantly reduces their viability. 13. Repeat the above steps to maintain a continuous, low-intensity culture. 10.

4.3. SOP for metered volumes of infective theronts for experimentation Some experiments require a more accurate determination of infec­ tive theronts for specific transmission values. This can be facilitated by collecting large numbers of tomonts (tomont aggregations; Fig. 1.3) from the culture, and incubating them together in a single 12-cm Petri dish. The cells are incubated at 24–26°C and observed daily under a dissection microscope for development and maturity, usually over 5–8 days. In the daylight hours preceding excystment (during the following period of darkness), the tomite divisions become slightly darker in colour and are individually well defined. In the early hours of the following morning (02:00–06:00; see Diggles and Lester, 1996) theronts become active inside the tomont and break free. These theronts are positively phototactic and will quickly congregate en masse in the Petri dish towards the strong­ est light source (Burgess and Matthews, 1994). Theronts 50, dilute the sample in PBS and count again. If the number is 2 g), parasite dose or intended outcome of the infection]. • Allow fish to recover in flow-through tanks with specific-patho gen-free water and held until signs of disease develop (2–4 weeks depending on temperature). Note: if part of an experimental trial, a control group should be inoculated with the same volume of PBS.

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Ceratonova shasta ♦  117

4.1.5. Monitoring fish for infection • Euthanize fish showing signs of clinical disease with anaesthetic. • Prepare a wet mount from an intestinal scraping or from ascites, if present. • Presumptive diagnosis is based on identification of characteristic trophozoite stages and confirmed by either parasite-specific PCR or detection of mature parasite spores (Fig. 8.4; Bartholomew, 2012). 4.2. Infection of fish in mesocosms Note: depending on the purpose of the study, it may be important to characterize the parasite genotype and maintain infections sepa rately according to genotype. This involves conducting a genotyp ing assay (Atkinson et al., 2018) on myxospores used originally for infection of the annelid cultures, then also on actinospores in water samples collected from the mesocosms. 4.2.1. Infection of donor fish • Infection of donor fish is as described in section 4.1; however, because the source material for infection of annelid cultures is mature myxospores, sterile conditions are not required and either whole intestine or an intestinal scraping is generally a good source of this material. 4.2.2. Collection of myxospores • Collect dead or moribund fish (euthanize the latter with an overdose of anaesthetic) to assure that the proportion of mature myxospores is high. • Necropsy fish and remove the entire intestine. Examine a scrap ing from the inner wall of the intestine under a microscope at 200× to confirm that spores are mature (isolated, bean-shaped myxospores) (Fig. 8.5). • Collect parasites by cutting open the intestine and using a

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118 ♦  Fish Parasites

Fig. 8.5 Necropsy of infected fish for collection of myxospores from the intestine (A). Mature myxospore stages collected for infection of annelids (B). Credit: Bartholomew Lab.

scalpel to scrape the epithelial layer into a tube containing water or PBS. • If information on genotype is necessary, sub-sample tissue from each tube for sequencing (Atkinson et al., 2018). Material from each fish should be stored separately in tubes in the refrigerator until the genotype assay is complete. 4.2.3. Infection of annelids • Add infected tissues directly to annelid mesocosms (section 2). Turn off incoming flow to the cultures for 24 h, while maintain­ ing aeration, to allow the myxospores to sink and be consumed by the worms. Turn water flow back on after 24 h. 4.2.4. Monitoring of annelid cultures for actinospore release • Beginning at 2 weeks post-exposure, collect 3 × 1-l water sam­ ples weekly to assay for parasite density. • Use a vacuum pump and filtration flask (Fig. 8.6) to draw the water sample through a 5-µm filter disc. Rinse the empty sample bottle with water from the wash bottle, pour the washings into the funnel and proceed to rinse the sides of the funnel. When all the liquid has passed through the filter, unclamp the funnel and use forceps to fold the filter disc in half 3–4 times, then place the folded filter in a 2-ml labelled microfuge tube. Hold on ice,

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Ceratonova shasta ♦  119

Fig. 8.6 Apparatus used for filtration of water samples (A). Filter with collected sample (B). Folded filter disc inserted in microfuge tube for molecular assay (C). Credit: Bartholomew Lab.

then freeze until qPCR assay (Hallett and Bartholomew, 2006; Hallett et al., 2012). • Alternately, if only small numbers of actinospores are needed (to infect a few fish or highly susceptible fish), annelids can be removed from the culture beginning 3–4 weeks after infection and examined individually for development of mature actino­ spores (Fig. 8.7). When mature spores are observed in the body wall of the worm, slight pressure applied to the coverslip will cause their release. Actinospores can be collected using a pipette and counted directly on a coverslip. 4.2.5. Infection of fish • Place fish directly in the flow-through effluent of the mesocosms for continuous exposure. Calculate the approximate exposure

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120 ♦  Fish Parasites

Fig. 8.7 Manayunkia occidentalis isolated in a culture dish (A). Mature actinospore stages collected for infection of fish (B). Credit: Bartholomew Lab.

dose using the qPCR assay results (Hallett et al., 2012) if the flow rate is known. Collect water samples at intervals during exposure and average the spore dose (e.g. three samples per day) for the duration of exposure. • Alternately, a volume of effluent may be collected and added to a tank of fish for a controlled exposure. In this case, the dose would be calculated directly using the qPCR assay result and volume of effluent collected. 4.2.6. Monitoring fish for infection • Euthanize fish showing signs of clinical disease with anaesthetic. • Prepare a wet mount from an intestinal scraping or from ascitic fluid, if present. • Presumptive diagnosis is based on identification of characteris­ tic trophozoite stages and confirmed by either parasite-specific PCR or detection of mature parasite myxospores (Bartholomew, 2012).

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Ceratonova shasta ♦  121

5. Summary templates Ceratonova shasta Noble, 1950 Parasite isolation from fish host Fish host

Rainbow/steelhead trout (O. mykiss), Chinook salmon (O. tshawytscha), coho salmon (O. kisutch); other salmonid species are less commonly used

Susceptible host characteristics

No sex or age/size distinctions

Environment of infected hosts

Fresh, specific-pathogen-free water, at 13–18°C is optimal for maintaining infections. Infection occurs, but progresses more slowly at lower temperatures; higher temperatures result in stress for fish and more rapid disease progression

Identification of infected individuals

Clinical disease signs include anorexia, lethargy, darkening, distention of the abdomen with ascites, exopthalmia and a swollen and haemorrhagic vent

Parasite stage(s) collected

Trophozoite (pre-sporogonic) stages for infection of fish; myxospores for infection of annelids

Size (mm) of parasite stages

~4–20 µm depending on stage

Site of infection, including target tissues if relevant

Ascitic fluid collects in the peritoneal cavity; intestine is the target tissue

Isolation technique(s)

Aspiration of ascites from peritoneal cavity of anaesthetized fish using a sterile needle and syringe

Additional isolation requirements

Microscope and slides; haemocytometer if quantifying inoculum

Post-processing

Not required, but parasites can be concentrated by gentle centrifugation and suspension in a small volume of PBS or by use of a plankton centrifuge

Collection medium

Can use ascites directly or wash cells and suspend in PBS

Transport/maintenance conditions

Hold in refrigerator or on ice until used for injection, within 24 h

Monitoring requirements

Visual, by microscopic examination at 200×

Optimal fixative(s) if using

Freeze or place in 100% ethanol if preserving sample for DNA extraction

Other relevant information

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122 ♦  Fish Parasites In vitro culture Axenic

N/A

With co-culture

Invertebrate annelid host M. occidentalis (Atkinson et al., 2020); collected from the river in sediment and tubes

Source of inoculum

Infected salmonid

Quantification of infective stages

Myxospores can be quantified using a haemocytometer, but this is generally not done to maintain an infected population

Dose of inoculum

Depends on size of annelid population, availability of myxospores and intent of experiment. Multiple dosing is often necessary to maintain infection

Culture conditions

Mesocosms maintain a river sediment habitat type as closely as possible

Monitoring of culture

Water samples are collected weekly beginning 2 weeks* after infection of annelid cultures, filtered and assayed by qPCR using parasite-specific primers to assess actinospore abundance. *Prior to this, myxospores will dominate

Renewal of culture medium, timing and harvesting

Actinospores may be harvested by collection of effluent or examination of worms for infected individuals. Infected worms must be examined immediately after removal from the mesocosm using a microscope at low magnification. Actinospores will be released from the body wall of the worm with light pressure on a coverslip

Parasite stages cultured

Actinospores

Storage

Hold on ice or refrigerate until use, within 24 h

Attenuation of virulence

There has been no research to demonstrate this

Other relevant information

Handling stress on individual, isolated worms should be minimized. The highest prevalence of infection (~80%) in mesocosm worms is in spring, but populations can produce spores year round.

In vivo infection challenge: injection of ascites Host characteristics: age, sex, weight if are relevant for the transmission or condition

Species of salmon and trout; no sex or age/size distinctions

Water conditions

Fresh, flow-through water at temperature permissive for the fish host and for infection to develop (~13–18°C)

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Ceratonova shasta ♦  123 Challenge dose

Dependent on host susceptibility; for susceptible trout strains, 1–10 parasites are sufficient to cause mortality; for resistant strains the dose may be thousands of parasites

Parasite stage/age/state

Trophozoite (presporogonic) stages; typically harvested from donor host 2–4 weeks following infection

Challenge method

Intra-abdominal injection

Intermediates/vectors involved

N/A

Timing of the infection: when it is expected to have a specific prevalence after exposure, and whether intermediate samplings have to be done before completion (duration of the experiment)

Dependent on dose, host susceptibility and water temperature. Susceptible fish receiving a heavy inoculum may die within days of injection. At lower temperatures and with a low infection dose, infection may develop over a few weeks, but the duration is generally shorter than a natural exposure

Containment/disinfection procedures of the facilities, water, used material, etc.

Surfaces and equipment may be cleaned with routine disinfectants; infected tissues are generally sterilized by autoclaving or incineration; effluent disinfection with chlorine or ozone

Type of facilities required

Tanks with flow-through specific-pathogen-free water at 13–18°C to support salmonid culture

Diagnosis of infection

Wet mounts prepared from an intestinal scraping or from ascitic fluid, if present. Presumptive diagnosis based on identification of characteristic trophozoite stages and confirmed by either parasite-specific PCR or detection of mature parasite spores.

In vivo infection challenge: Exposure to actinospores from mesocosms Host characteristics

Species of salmon and trout; no sex or age/size distinctions

Water conditions

Fresh, flow-through water at temperature permissive for the fish host and for infection to develop (~13–18°C)

Challenge dose

Dependent on host susceptibility; for susceptible trout strains, 1–10 parasites are sufficient to cause mortality; for resistant strains the dose may be thousands of parasites Dose is quantified by qPCR of 1 l of effluent collected on a 5-µm filter. Assay three samples for each day of exposure and average to obtain dose per litre; use flow rate to calculate total exposure

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124 ♦  Fish Parasites If worms are examined individually, actinospores may be directly counted on a coverslip and washed into an exposure vessel. This method is suitable for infection of only a small number of fish, or of highly susceptible fish Parasite stage/age/state

Actinospores

Challenge method

Fish are typically exposed in mesocosm effluent

Intermediates/vectors involved

Invertebrate host M. occidentalis is the actinospore source

Timing of infection: when it is expected to have a specific prevalence after exposure, and whether intermediate samplings have to be done before completion (duration of the experiment)

Dependent on temperature, dose and host susceptibility; at 13°C mortality will begin later and total mortality will be lower than at higher temperatures. Generally, mortality begins 2–3 weeks following exposure and continues for 4–6 weeks

Containment/disinfection procedures of the facilities, water, used material, etc.

Surfaces and equipment may be cleaned with routine disinfectants; infected tissues are generally sterilized by autoclaving or incineration; effluent disinfection with chlorine or ozone

Type of facilities required

Tanks with flow-through specific-pathogen-free water at temperatures of 13–18°C to support salmonid culture

Diagnosis of infection

Wet mounts prepared from either an intestinal scraping or ascitic fluid, if present. Presumptive diagnosis based on identification of characteristic trophozoite stages and confirmed by either parasitespecific PCR (Palenzuela et al., 1999) or detection of mature parasite spores.

6. Glossary Actinospore: Waterborne spore stage of a myxosporean parasite that develops in the annelid host and is infectious to fish. Ascites: Extracellular fluid that accumulates in body cavities in response to a disease process; for myxozoans the fluid contains developing parasites. Enzootic: Native to a specific geographic location. Myxospore: Waterborne spore stage of a myxosporean parasite that devel­ ops in the fish host and is infectious to the annelid.

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Ceratonova shasta ♦  125 Myxozoan: Member of a group of parasitic Cnidaria: microscopic, reduced body organization and genomes compared with free-living cnidarians; responsible for significant diseases in both wild and cultured fish. Nematocyst: Specialized organelle with evertable tubule, found in all Cnidaria; used in myxosporeans to attach spores to the host for initia­ tion of infection. Trophozoite: One of the growing stages of a parasite.

7. References Atkinson, S.D. and Bartholomew, J.L. (2010) Disparate infection patterns of Ceratomyxa shasta (Myxozoa) in rainbow trout Oncorhynchus mykiss and Chinook salmon Oncorhynchus tshawytscha correlate with ITS-1 sequence variation in the parasite. International Journal for Parasitology 40, 599–604. Atkinson, S.D., Hallett, S.L. and Bartholomew, J.L. (2018) Genotyping of individual Ceratonova shasta (Cnidaria: Myxosporea) myxospores reveals intra-spore ITS-1 variation and invalidates the distinction of genotypes II and III. Parasitology 145, 1588–1593. Atkinson, S.D., Bartholomew J.L. and Rouse G.W. (2020) The inverte­ brate host of salmonid fish parasites Ceratonova shasta and Parvicapsula minibicornis (Cnidaria: Myxozoa), is a novel fabriciid annelid, Manayunkia occidentalis sp. nov. (Sabellida: Fabriciidae). Zootaxa 4751, zootaxa.4751.2.6. Bartholomew, J.L. (2012) Salmonid ceratomyxosis. In: AFS-FHS (American Fisheries Society-Fish Health Section). FHS Blue Book: Suggested Procedures for the Detection and Identification of Certain Finfish and Shellfish Pathogens, 2010 edn. AFS-FHS, Bethesda, Maryland, USA, 14 pp. Bartholomew, J.L., Whipple, M.J., Stevens, D.G. and Fryer, J.L. (1997) The life cycle of Ceratomyxa shasta, a myxosporean parasite of salmo­ nids, requires a freshwater polychaete as an alternate host. American Journal of Parasitology 83, 859–868. Bjork, S.J. and Bartholomew, J.L. (2009) The effects of water velocity on the Ceratomyxa shasta infectious cycle. Journal of Fish Diseases 32, 131–142. Bjork, S.J. and Bartholomew, J.L. (2010) Invasion of Ceratomyxa shasta (Myxozoa) and comparison of migration to the intestine between

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126 ♦  Fish Parasites susceptible and resistant fish hosts. International Journal for Parasitology 40, 1087–1095. Hallett, S.L. and Bartholomew, J.L. (2006) Application of a real-time PCR assay to detect and quantify the myxozoan parasite Ceratomyxa shasta in river water samples. Diseases of Aquatic Organisms 71, 109–118. Hallett, S.L., Ray, R.A., Hurst, C.N., Holt, R.A., Buckles, G.R., Atkinson, S.D. and Bartholomew, J.L. (2012) Density of the waterborne para­ site, Ceratomyxa shasta, and its biological effects on salmon. Applied and Environmental Microbiology 78, 3724–3731. Hurst, C.N., Alexander, J.D., Jia L., Dolan B. and Bartholomew, J.L. (2019) Outcome of within-host competition demonstrates that para­ site virulence doesn’t equal success in a myxozoan model system. International Journal for Parasitology 9, 25–35. Hurst, C.N. and Bartholomew, J.L. (2012) Ceratomyxa shasta genotypes cause differential mortality in their salmonid hosts. Journal of Fish Diseases 35, 725–732. Palenzuela, O., Trobridge, G. and J.L. Bartholomew (1999) Development of a polymerase chain reaction diagnostic assay for Ceratomyxa shasta, a myxosporean parasite of salmonid fish. Diseases of Aquatic Organisms 36, 45–51. Ray, A.R, Holt, R.A. and Bartholomew, J.L. (2012) Relationship between temperature and Ceratomyxa shasta-induced mortality in Klamath River salmonids. Journal of Parasitology 98, 520–526.

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Chapter 9

Enteromyxum leei Ariadna Sitjà-Bobadilla1, Itziar Estensoro1 and

Oswaldo Palenzuela1

1

Fish Pathology Group, Instituto de Acuicultura Torre de la Sal (IATS), Consejo Superior de Investigaciones Científicas (CSIC), 12595 Ribera de Cabanes, Castellón, Spain; e-mails: [email protected]; [email protected]; [email protected].

1. Background

E

nteromyxum leei is a microscopic myxozoan parasite (Cnidaria: Myxozoa: Myxosporea) that infects the intestinal tract of fish and sometimes associated organs, such as the gall bladder and liver. The life cycle of myxosporeans generally involves two alternating hosts: fish and annelids. Myxospores are ingested by annelids, infect­ ing their gut epithelium or the epidermis and subsequently produc­ ing actinospores. The actinospores released from the annelid remain in the water or sediments until they reach a fish surface (skin or gills) and penetrate through the epithelium. Once in the fish host, the developmental stages migrate until they reach the final site of infec­ tion and develop into myxospores. Although these diheteroxenous cycles have been described for about 50 myxozoan species (of more than 2200 described taxa), laboratory studies covering the whole cycle in both invertebrate and vertebrate hosts have been completed for five species only (Eszterbauer et al., 2015). This gives an idea of the intrinsic difficulty in setting up this type of experimental model. In contrast to this complex life cycle, direct spontaneous fish-to-fish transmission has been demonstrated only for species belonging to the genus Enteromyxum, in various marine fish (SitjàBobadilla and Palenzuela, 2012). E. leei developmental stages (also

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128 ♦  Fish Parasites

named trophozoites) and myxospores are released into the water with faeces, or from dead fish. Released trophozoites are infective to other fish, and thus responsible for horizontal transmission. Although it is possible that this myxozoan also has an indirect life cycle involving invertebrate hosts in the wild, fish-to-fish transmis­ sion by cohabitation, contaminated water and oral and anal intu­ bation has been experimentally achieved (Table 9.1), but the exact protocol has never been described in detail. Transmission between different fish species has also been demonstrated. This unique mode of horizontal transmission favours the spread of enteromyxoses in cultured fish stocks and allows for its in vivo maintenance under laboratory conditions (Fig. 9.1). In addition, continuous in vitro cultivation of myxozoans has not been achieved and, in particular, various attempts to culture Enteromyxum species have not been fully successful (Redondo et al., 2003). Thus, only in vivo standard operating procedures (SOPs) will be included for this parasite. There are different ways to achieve E. leei transmission, but both procedures described here have been repeated and improved successfully for gilthead sea bream for more than 10 years: anal intubation and effluent exposure. Figure 9.2 diagrammatically shows these two procedures. Fig. 9.1 Life cycle of Enteromyxum leei. Proliferative (a–c) and sporogonic (d–f) development in the intestinal epithelium of the fish. Stages a–e are responsible for invasion and dispersion within the fish, as well as for transmission to other fish, when released through faeces, or when dead fish are eaten. It is currently unknown whether mature spores (f) start an alternate cycle infecting an invertebrate host. Credit: Itziar Estensoro, Fish Pathology group, IATS.

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Enteromyxum leei ♦  129 Table 9.1. Available data in the literature on experimental infections achieved for E. leei by feeding (F) or inoculating orally (O) or anally (A) with infected gut scrapings, by cohabitation (CH) or by contact with effluent (EF) water from infected donor tanks. D, donor fish species; R, recipient fish species; A, allogeneic donor; X, xenogeneic donor; p.e., post-exposure. Mode of D infection

R

Days p.e.

Prevalence (%)*

Reference(s)

F, O

A

Sparus aurata

X X

13 0–93.3 80 90

Diamant, 1997 Sitjà-Bobadilla et al., 2007 Diamant et al., 2006 Diamant et al., 2006

X X A

Puntius tetrazona1,2 Oreochromis mossambicus1,2 Astronotus ocellatus1,2 Danio rerio1,2 Takifugu rubripes

35 28–29 35 43 35 40 42

85 53 100

F A CH

X A X X X X A A A

Pagrus major Paralichthys olivaceus T. rubripes T. nipholes1 Amphiprion clarki1 Amphiprion ocellaris1 Epinephelus malabaricus Sparus aurata S. aurata

CH

X X A

Sciaenops ocellatum Diplodus puntazzo D. puntazzo

A X A A

T. rubripes Pagrus major E. malabaricus S. aurata

X X A X

Sciaenops ocellatum Dicentrarchus labrax T. rubripes Pagrus major

42 32 23 N/A 28 28 33 60–216 63 56 71–113 43 10 19 19 128 70 33 63 56 77–84 43 139 41 70

57.1 100 40 N/A 50 67 80 46.1–100 31.6 50 10–92 45.8 100 100 100 28.5 42.8 100 33.3 40 2.4–78 35 53.3 40 25

Diamant et al., 2006 Diamant et al., 2006 Yasuda et al., 2002 Yanagida et al., 2006 Yanagida et al., 2004 Yasuda et al., 2005 Yasuda et al., 2005 Yokoyama and Shirakashi, 2007 Yokoyama and Shirakashi, 2007 Yokoyama and Shirakashi, 2007 China et al., 2013 Estensoro et al., 2010 Diamant, 1997 Diamant and Wajsbrot 1997 Sitjà-Bobadilla et al., 2007 Diamant ,1998 Álvarez-Pellitero et al., 2008 Golomazou et al., 2006 Muñoz et al., 2007 Yasuda et al., 2002 Yanagida et al., 2008 China et al., 2013 Diamant, 1997 Diamant and Wajsbrot, 1997 Sitjà-Bobadilla et al., 2007 Diamant, 1998 Sitjà-Bobadilla et al., 2007 Yasuda et al., 2002 Yanagida et al., 2008

F

F

CH CH EF

EF

*Highest value achieved, when different sampling times p.e. were reported from a single experiment.

A maximum–minimum range is provided for different experiments.

1 This infection was either not detected naturally or no data were available under the particular farming

conditions. 2 Freshwater species; the remainder are marine fish.

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130 ♦  Fish Parasites

Fig. 9.2 Schematic drawing of Enteromyxum leei SOPs. D, donor; R, recipient fish. Credit: Ariadna Sitjà-Bobadilla, Fish Pathology group IATS.

The other two possible procedures, oral (intubation with intestinal scrapings or feeding with chopped or minced infected intestines) and cohabitation, are also feasible. The oral method is more similar to the natural infection route, but its reproducibility under labora­ tory conditions is low due to the action of digestive enzymes that can inactivate trophozoites if they are not protected by fish host cells/mucus due to the isolation procedure, and may also yield dif­ ferent results between agastric and gastric fish. The latter method would be very similar to effluent transmission, with the cohabita­ tion of infected and naïve fish in the same tank (either together or separated by a screen, to deter aggressive behaviour and can­ nibalism when size difference is wide). Anal intubation is a more homogeneous and simultaneous way to infect all recipient fish in a rapid and reproducible way, with many replicates, whereas efflu­ ent exposure is similar to what may occur under farm conditions but it is slower and not homogeneous. The choice of transmission method depends on the purpose of the study.

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Enteromyxum leei ♦  131

2. General facilities and fish For both transmission procedures a stock of infected gilthead sea bream (Sparus aurata) is necessary; these fish are called donors (D) throughout the document. Donor fish can come from either a previ­ ously infected stock under laboratory conditions or farmed stocks. In the latter case, fish should be allocated in a quarantine area and their health status checked to exclude other parasites or pathogens that could be transmitted to the experimental stock. Fish to be used as recipient (R) of the infection should be purchased from hatcherynursery facilities with no previous records of E. leei infection. Upon arrival a representative number of fish should be analysed for the absence of E. leei. Depending on the purpose of the transmission experiment and planned downstream analyses, it is advised to avoid using R fish with other concomitant gastrointestinal infections, such as Enterospora nucleophila or Cryptosporidium molnari. Recipient fish should be acclimated to the experimental condi­ tions for at least 2 weeks before the beginning of the experiment, or grown on until the desired size. Water oxygen content should be kept >85% saturation, and unionized ammonia below toxic levels (18°C for non-latent infections and thus heaters may be needed, depending on latitude and season. Water effluent from infected fish stocks (D and R) needs disinfec­ tion treatment (chlorination, ozone, etc.) before being discharged, and disinfection of all material (tanks, buckets, nets, aerators, etc.) will be required after contact with infected fish or water.

3. Equipment and reagents Basic: • Plastic sterile Pasteur pipettes, with blunt end cut with a sterile scalpel • Plastic sterile Petri dishes

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• • • •

Dissection tools Automatic pipettes and pipette tips Glass slides and coverslips PBS or HBSS with antibiotics (penicillin/streptomycin or genta­ mycin commercial solutions) • Anaesthetic (clove oil, MS-222, etc.) • Syringes • Light microscope. Optional: • • • •

PIT-tag gun and detector Thermocycler Sterile cotton/alginate swabs for rectal biopsies. Lysis buffer/DNA preserving medium for PCR testing of swabs and inocula • Centrifuge (50 and 15 ml) sterile tubes • Microcentrifuge (1.5 ml) sterile tubes.

4. Enteromyxum leei transmission 4.1. Transmission by anal intubation Obtaining the inoculum: PIT-tag donor fish if they need to be selected and tracked indi­ vidually afterwards on the basis of their infective status. 2. Starve both D and R fish two days before they are going to be used. 3. Lightly anaesthetize the fish with clove oil for easy and stressfree handling. 4. Check the E. leei infective status of D fish non-lethally (NL). This can be done by either: a. Rectal probing (see Fox et al., 2000; Palenzuela and Bartholomew, 2002) and subsequent PCR detection of the 18S ribosomal RNA gene. NL-qPCR will take several hours 1.

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5.

6.

7.

8.

9.

to obtain the results and fish should not be used the same day that they have been sampled. b. Anal cannulation of D fish with a Pasteur pipette and direct microscopic observation of the fresh smear with a drop of PSB. Only fish with smears in which the parasite can easily be detected at low magnification (not higher than 200×) and in high amounts are considered good candidates to be used as donors for the anal inoculum. Annotate the type of stages detected (trophozoites, sporoblasts, spores according to the stages described in Álvarez-Pellitero et al., 2008) (Fig. 9.3) and the intensity of infection following a semi-quanti­ tatively conventional scale from 1+ to 6+, with the following ranges: 1+ = 1–5; 2+ = 6–10; 3+ = 11–25; 4+ = 26–50; 5+ = 51–100; 6+ >100 parasite stages per microscope field obser­ vation at 120×. This procedure, though less accurate, allows a quick and easy selection of infected fish just prior to their use as donors. The same day of inoculation, sacrifice D fish with an overdose of anaesthetic (e.g. 3-aminobenzoic acid ethyl ester, MS-222, at 0.1 g/l) and bleed them from the caudal vessels with a syringe, to avoid blood congestion in the visceral cavity. At necropsy, remove the intestine as a single piece and place it in a sterile Petri dish. Keep the tissues cool at all times (e.g. over a cold bench or close to ice blocks). Take a small sample of the three segments of the intestine (ante­ rior, middle and posterior) and scrape the epithelium to obtain a fresh smear. Observe immediately under the microscope and evaluate using the same criteria as described in step 4.b) in order to select tissues with high infection intensity (≥4). Avoid unnec­ essary dilution of the inoculum with less infected segments and avoid intestines with bacterial invasion. The selected intestinal segments are then cut open and lightly scraped with the non-cutting side of a surgical blade to peel off the epithelial layer where the parasite is harbouring. Dilute with sterile saline containing antibiotics (PBS/HBSS) as needed to facilitate the pooling and homogenization of scrap­ ings in a single 50- or 15-ml tube and keep cool until used.

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Fig. 9.3 Fresh smear of an intestinal scraping with abundant proliferative stages of Enteromyxum leei. Credit: Ariadna SitjàBobadilla, Fish Pathology group, IATS.

Take an aliquot of the inoculum for further observation with a light microscope or for qPCR, if quantification of the parasites is needed. Dilute as needed to obtain the desired volume and concentration for the infection of all R fish (see challenge dose in summary templates). Anal intubation: Gently anesthetise R fish with clove oil and smoothly introduce the pipette tip (with wide bore or previously cut at the bevel, if necessary) into the anal pore (Fig. 9.4). The size of the pipette or pipette tip will depend on fish size. Introduce an adequate volume of inoculum, depending on fish size. It is very impor­ tant to perform this step rapidly (if necessary, two persons at the same time) to avoid decrease in parasite viability. Keep the tubes containing the inoculum cool, but avoid direct contact with ice and gently mix them every time an aliquot is taken to ensure its homogeneity. 2. Place the fish back in the water container with aeration to recover from the anaesthetic, and then back into their corresponding tank. 1.

Note 1: If part of an experimental trial and not for the normal main­ tenance of the infection, a control group should be inoculated with the same volume of PBS.

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Fig. 9.4 Anal intubation of a small juvenile with automatic pipette. Credit: Ariadna SitjàBobadilla, Fish Pathology group, IATS.

Note 2: This inoculum could also be used for oral intubation of R fish. We do not describe the associated SOP, as the results thus far obtained with this procedure are less reliable and repeatable. 4.2. Enteromyxum leei transmission by water effluent exposure Set-up of D/R fish and time of exposure: Two or more tanks holding naïve R fish are connected to receive the effluent water from another tank containing D fish until the end of the experiment. The D tank is the only one that receives inlet water and all tanks must be supplied with aeration (rec­ ommended) or oxygen. Water flow to R tanks should be equal if a constant infective pressure on R groups is desired. Water flow to the D tank needs to be set to support all the biomass in the experimental set-up, but low enough to ensure that parasite stages are not rapidly removed. 2. D fish are previously selected by non-lethal methods (as explained in section 4.1). The infection status of D fish must be high and relatively recent, as we have observed that fish with old infec­ tions tend to be less infective. 3. D/R biomass ratio and water flow largely influence the infective pressure on R fish, so it is recommended to keep this within a relatively stable range and, if possible, substitute dead D fish with other infected fish in order to obtain predictable results. A 1.

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D/R biomass ratio of 1.0 is recommended, though a range from 1.8 to 0.7 has proven to be satisfactory; however, other ranges could possibly work. Time of exposure to the effluent is relevant to achieving high infection levels in R fish. For gilthead sea bream, one week is enough to achieve 100% prevalence of infection under high water tempera­ ture (Picard-Sánchez et al., 2020).

5. Summary templates Enteromyxum leei Diamant, Lom & Dyková, 1994 In vivo experimental transmission: general Fish or intermediate host species Gilthead sea bream (S. aurata), other fish species are required for parasite collection susceptible Susceptible host characteristics

No sex distinctions. No effect of R size on the prevalence of infection has been detected

Environment of infected hosts

Temperature: never 28°C) for extended periods. Recommended to use 1-µm filtration and UV irradiation to avoid the entrance of other pathogens. Natural sea water. Salinity is not relevant

Identification of infected individuals

18 S rRNA PCR detection (lethal from intestinal tissue sample or non-lethal from rectal probe). Light microscope examination (lethal from intestinal scraping or non-lethal from anal cannulation)

Parasite stage(s) collected

Trophozoites: primary, secondary and tertiary cells. Disporoblasts may be present

Size of parasite stages

10 µm single cell – 100 µm multicellular

Site of infection, including target tissues if relevant

Target tissue is the intestine, preferably the posterior segment. Associated organs may be affected (liver, gall bladder)

Isolation technique(s)

N/A

Additional isolation requirements

Light microscope or thermocycler for determination of infection intensity

Post-processing

N/A

Collection media

PBS or HBSS with antibiotics (penicillin/streptomycin or gentamycin commercial solutions)

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Enteromyxum leei ♦  137 Transport/maintenance conditions

N/A

Monitoring requirements

N/A

Optimal fixative(s) if using

N/A

Disinfection procedures

Effluent disinfection of water with chlorine, ozone or electrolysis systems, as implemented in the facilities. Fish handling nets and tools, aerators, pipes, etc., disinfected with routine surface disinfectants as used in the facilities. Allow physical separation of tanks to exclude water splashes or cross-contamination of tanks due to malpractice

Type of facilities required

Separate tanks from general tanks in the facilities, with a different water system, either in recirculation or open flow. Information has been obtained from open flow systems

Other relevant information

Termination of infection is decided depending on the prevalence and intensity of infection achieved. Infected R fish can then be used as D for other infections and maintain the infection in vivo at the laboratory. Alternatively, part of the infected fish can be used to obtain histological or molecular samples for other purposes. If trials go on for longer periods, the prevalence of infection tends to decrease, especially under high water temperatures

In vivo experimental transmission: Anal intubation Host characteristics of R fish

Prevalence values between 66 and 100% have been achieved using R fish from 17 to 315 g

Feeding

R fish can be fed with automatic feeders, but it is recommended to feed manually ad libitum if feed intake has to be registered to monitor anorexia caused by the disease. Use dry pellet commercial diet at about 1–1.5% of body weight daily

Challenge dose

qPCR Ct values of the inoculum between 18 and 32.4 are capable of inducing 90–100% prevalence of infection. The inoculated volume depends on fish size, from 0.15 to 1.0 ml. As a guide: fish smaller 200 g: up to 1 ml

Parasite stage

Trophozoites: primary, secondary and tertiary cells

Challenge method

Anal intubation with intestinal scrapings (see further details in section 4.1)

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None known thus far

Timing of infection

Depends on water temperature. A good end point at water temperature >20°C is about 60–70 days post challenge. It can take longer (about 90 days) at 18°C

Diagnosis of infection

Disease signs and daily mortalities have to be recorded and the parasitic status of dead fish checked by microscopic observation of intestinal scrapings or qPCR. Progression of infection is checked at the mid­ point (at 35 or 40 days post inoculum) by NL-qPCR to determine the best endpoint

In vivo experimental transmission: Effluent exposure Host characteristics: age, sex, weight

Infection has been achieved using fish from 23.3 to >300 g

Water conditions (salinity, oxygen, temperature, open/ recirculating system, etc.)

For D tanks: Water flow should be high enough to support all the biomass in the set-up. Supplementary oxygen or aeration can be provided to R tanks. Together with water flow and D/R ratio, temperature modulates the infective pressure and rate of transmission

Feeding

Avoid immunostimulant-containing diets and feed as usual in the facilities. Feed intake will be reduced in infected fish: it is advised to feed manually ad libitum to monitor anorexia caused by the disease. Use commercial, dry pellet diet at about 1–1.5% of body weight daily, depending on fish size and temperature. Use the same type of food for D and R fish

Challenge dose

D/R biomass ratio of 0.7–2.8 can achieve prevalence of 50–100%. D infection status (intensity and time since first exposure) and D/R ratio are relevant

Parasite stage

Trophozoites surrounded by mucus casts are released by D with faeces into the water, and float on the surface of the tank. They are transported passively with the water to neighbouring tanks

Challenge method

Exposure to the effluent water of tanks holding D fish

Intermediates/vectors involved

None known thus far

Timing of infection

Depends on water temperature. A good end point at water temperature >20°C is about 90 days postexposure. It can take longer (about 120 days) at 18°C

Diagnosis of infection

The infection is checked before the end point (at 60–70 days post-exposure) by NL-qPCR to determine the appropriate endpoint

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6. Glossary Actinospores: Spore stage developed during the myxosporean sexual pro­ cess in the definitive host (the invertebrate). These are infective for the vertebrate host. Diheteroxenous cycle: A heteroxenous life cycle, with complex or indi­ rect parasite life cycles involving more than one host. A diheteroxenous cycle alternates between two hosts. Horizontal transmission: Direct transmission of a parasite between hosts. There is no need, as in vertical transmission, for host reproduction in order to transmit the parasite to the hosts’ progeny. Myxospores: Spore stage developed in the intermediate host (the verte­ brate, usually fish), for parasite multiplication. They are infective for the invertebrate host. Myxozoa: Phylum of microscopic cnidarian parasites, with >2,200 described species. They are mostly aquatic and have complex life cycles involving alternate hosts. Trophozoites: proliferative vegetative myxozoan stages that invade the host tissue prior to sporogony. Their main role is multiplication, migra­ tion and nutrition in the host.

7. References Álvarez-Pellitero, P., Palenzuela, O. and Sitjà-Bobadilla, A. (2008) Histopathology and cellular response in Enteromyxum leei (Myxozoa) infections of Diplodus puntazzo (Teleostei). Parasitology International 57, 110–120. China, M., Nakamura, H., Hamakawa, K., Tamaki, E., Miwa, S., Meng, F. and Yokoyama, H. (2013) Occurrence of the myxosporean emacia­ tion disease caused by Enteromyxum leei in cultured malabar grouper Epinephelus malabaricus. Fish Pathology 48, 88–96. Diamant, A. (1997) Fish-to-fish transmission of a marine myxosporean. Diseases of Aquatic Organisms 30, 99–105. Diamant, A. (1998) Red drum Sciaenops ocellatus (Sciaenidae), a recent introduction to Mediterranean mariculture, is susceptible to Myxidium leei (Myxosporea). Aquaculture 162, 33–39. Diamant, A. and Wajsbrot, N. (1997) Experimental transmission of Myxidium leei in gilthead sea bream Sparus aurata. Bulletin of the European Association of Fish Pathologists 17, 99–103.

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140 ♦  Fish Parasites Diamant, A., Ram, S. and Paperna, I. (2006) Experimental transmission of Enteromyxum leei to freshwater fish. Diseases of Aquatic Organisms 72, 171–178. Estensoro, I., Redondo, M.J., Álvarez-Pellitero, P. and Sitjà-Bobadilla, A. (2010) Novel horizontal transmission route for Enteromyxum leei (Myxozoa) by anal intubation of gilthead sea bream Sparus aurata. Diseases of Aquatic Organisms 92, 51–58. Eszterbauer, E., Atkinson, S., Diamant, A., Morris, D., El-Matbouli, M. and Hartikainen, H. (2015) Myxozoan life cycles: Practical approaches and insights. In: Okamura, B. et al. (eds), Myxozoan Evolution, Ecology and Development. Springer International Publishing, Switzerland. Fox, M.D., Palenzuela, O. and Bartholomew, J.L. (2000) Strategies for the diagnosis of Ceratomyxa shasta using the polymerase chain reaction: comparison of lethal and nonlethal sampling with microscopic exami­ nation. Journal of Aquatic Animal Health 12, 100–106. Golomazou, E., Athanassopoulou, F., Karagouni, E., Tsagozis, P., Tsantilas, H. and Vagianou S. (2006) Experimental transmission of Enteromyxum leei Diamant, Lom and Dyková, 1994 in sharpsnout sea bream, Diplodus puntazzo C. and the effect on some innate immune parameters. Aquaculture 260, 44–53. Muñoz, P., Cuesta, A., Athanassopoulou, F., Golomazou, H., Crespo, S., Padrós, F., Sitjà-Bobadilla, A., Albiñana, G., Esteban, M.A., ÁlvarezPellitero, P. and Meseguer, J. (2007). Sharpsnout sea bream (Diplodus puntazzo) humoral immune response against the parasite Enteromyxum leei (Myxozoa). Fish and Shellfish Immunology 23, 636–645. Palenzuela, O. and Bartholomew, J.L. (2002) Molecular tools for the diagnosis of Ceratomyxa shasta (Myxozoa). In: Cunningham, C.O. (ed.). Molecular Diagnosis of Salmon Diseases. Springer Netherlands, Dordrecht, the Netherlands, pp. 285–298. Picard-Sánchez, A., Estensoro, I., Del Pozo, R., Palenzuela, O.R., Piazzon, M.C. and Sitjà-Bobadilla, A. (2020) Water temperature, time of expo­ sure and population density are key parameters in Enteromyxum leei fish­ to-fish experimental transmission. Journal of Fish Diseases 43, 491–502. Redondo, M.J., Palenzuela, O. and Álvarez-Pellitero, P. (2003) In vitro studies on viability and proliferation of Enteromyxum scophthalmi (Myxozoa), an enteric parasite of cultured turbot Scophthalmus maxi­ mus. Diseases of Aquatic Organisms 55, 133–144. Sitjà-Bobadilla, A. and Palenzuela, O. (2012) Enteromyxum species. In:

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Enteromyxum leei ♦  141 Woo, P.T.K. and Buchmann, K. (eds). Parasites: Pathobiology and Protection. CAB International, Oxford, UK, pp. 163–176. Sitjà-Bobadilla, A., Diamant, A., Palenzuela, O. and Álvarez-Pellitero, P. (2007) Effect of host factors and experimental conditions on the horizontal transmission of Enteromyxum leei (Myxozoa) to gilthead sea bream, Sparus aurata L., and European sea bass, Dicentrarchus labrax (L.). Journal of Fish Diseases 30, 243–250. Yanagida, T., Nomura, Y., Kimura, T. Fukuda, Y., Yokoyama, H. and Ogawa, K. (2004) Molecular and morphological redescriptions of enteric myxozoans, Enteromyxum leei (formerly Myxidium sp. TP) and Enteromyxum fugu comb. n. (syn. Myxidium fugu) from cultured tiger puffer. Fish Pathology 39, 137–143. Yanagida, T., Palenzuela, O., Hirae, T. Tanaka, S., Yokoyama, H. and Ogawa, K. (2008) Myxosporean emaciation disease of cultured red sea bream Pagrus major and spotted knifejaw Oplegnathus punctatus. Fish Pathology 43, 45–48. Yanagida, T., Sameshima, M., Nasu, H., Yokoyama, H. and Ogawa, K. (2006) Temperature effects on the development of Enteromyxum spp. (Myxozoa) in experimentally infected tiger puffer, Takifugu rubripes (Temminck & Schlegel). Journal of Fish Diseases 29, 561–567. Yasuda, H., Ooyama, T., Iwata, K. Tun, T., Yokoyama, H. and Ogawa, K. (2002) Fish-to-fish transmission of Myxidium spp. (Myxozoa) in cultured tiger puffer suffering from emaciation disease. Fish Pathology 37, 29–33. Yasuda, H., Ooyama, T., Nakamura, A., Iwata, K., Palenzuela, O. and Yokoyama, H. (2005) Occurrence of the myxosporean emaciation disease caused by Enteromyxum leei in cultured Japanese flounder Paralichthys olivaceus. Fish Pathology 40, 175–180. Yokohama, H. and Shirakashi, S. (2007) Evaluation of hyposaliniy treat­ ment on infection with Enteromyxum leei (Myxozoa) using anemonefish Amphiprion spp. as experimental host. Bulletin of the European Association of Fish Pathologists 27, 74–78.

8. Acknowledgements This SOP is the result of more than a decade of continuous research on this parasite model, with funding in recent years from Spanish (AGL2013–48560-C2–2-R) and EU (ParaFishControl, 634429)

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projects. The authors are indebted to Professor Pilar Álvarez-Pellitero, for her tenacity and efforts in starting this research line. She coordi­ nated the first European project (MyxFishControl) to work on this disease along the Mediterranean basin (QLRT-2001–00722).

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Chapter 10

Enteromyxum scophthalmi Oswaldo Palenzuela1, M. Carla Piazzon1 and Ariadna

Sitjà-Bobadilla1

1

Fish Pathology Group, Department of Marine Species Biology, Culture and Pathology, Instituto de Acuicultura Torre de la Sal (IATS), Consejo Superior de Investigaciones Científicas (CSIC), Spain; e-mails: [email protected]; [email protected]; [email protected]

1. Background

E

nteromyxum scophthalmi is the causative agent of a parasitic ema­ ciative disease that affects turbot (Scopthalmus maximus), known as sunken-head syndrome or turbot enteromyxosis. This organism is a myxozoan, a group of microscopic endoparasites that resemble protozoans but which are in fact metazoans, related to free-living cnidarians. E. scophthalmi is akin to E. leei, a different species which causes similar syndromes in various fish species in the Mediterranean and worldwide (reviewed in Sitjà-Bobadilla and Palenzuela, 2012). E. scophthalmi develops in the intestinal epithelium of turbot, caus­ ing severe catarrhal enteritis and impairing intestinal function. The infection is usually lethal just a few weeks after infection. Although typical myxozoan life cycles usually alternate between a fish and an invertebrate host (annelids or bryozoans), Enteromyxum spp. are unique in their ability to be transmitted directly between fish. Thus, E. scophthalmi has been demonstrated to be experimen­ tally transmissible to naïve turbot from infected fish through three routes: (1) cohabitation in the same tank; (2) receiving contami­ nated effluent water from tanks holding the infected fish; and (3) the oral route (Redondo et al., 2002, 2004). In aquaculture set­ tings this transmission occurs readily and spontaneously, completely

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apart from the putative natural occurrence of an alternating life cycle (which is currently unknown). It is believed that these routes of contagion of E. scophthalmi involve parasite cells (throphozoites) present throughout the entire developmental cycle in the fish, but no spores or other resistant stages. Indeed, mature spore develop­ ment in turbot is usually scarce. The catarrhal enteritis induced by the parasite favours their release into the water within epithelium remnants and mucus casts, which confer short-term protection. Developmental stages of E. scophthalmi do not survive in sea water more than 24 h, and continuous in vitro cultivation has not been fully achieved (Redondo et al., 2003). Therefore, the infection can be maintained in the laboratory only by in vivo passage, which is described in this chapter.

2. Facilities and fish For the laboratory transmission of enteromyxosis, a stock of natu­ rally or laboratory-infected turbot is necessary, and these fish are termed donors (D). Since other parasites or pathogens could be transmitted to the experimental stock, general quarantine precau­ tions and health checks are advised before incorporating farm stocks for use as donors. Fish to be used as recipients (R) of the infection should be purchased from known hatchery-nursery facilities (see recommended sizes in section 5). Upon arrival, a representative number of fish should be analysed for the absence of E. scophthalmi and other pathogens. Depending on the purpose of the transmission experiment and planned downstream analyses, it is advised to avoid using R fish with other concomitant gastrointestinal infections, such as Cryptosporidium scophthalmi. Turbot should be maintained in flat-bottom tanks using standard conditions of water flow and quality, aeration/oxygen, density and feeding regimen, as recommended elsewhere (e.g. Person-Le Ruyet, 2010). Temperature should be below 20°C. In Mediterranean facili­ ties, cooling of water is necessary to maintain turbot year round, by the use of recirculation units. In such cases, control and infected stocks must be kept on separate circuits.

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3. Equipment and reagents Basic: • • • • • • • • • •

Plastic disposable Pasteur pipettes, wide bore Plastic sterile Petri dishes (60–90 mm) Dissecting tools Automatic pipettes (1.0 and 0.2 ml) and pipette tips (wide bore or pre-cut) Sterile 50- and 15-ml centrifuge tubes Sterile 1.5-ml Eppendorf tubes Glass slides and coverslips Sterile saline (PBS or HBSS) containing antibiotics (penicillin/ streptomycin or gentamycin commercial solutions) Anaesthetic (clove oil, MS-222, 3-aminobenzoic acid ethyl ester, etc.) Light microscope.

Optional: • Tagging methods (see notes on tagging in section 4.3) • Sterile cotton swabs for rectal biopsies • Lysis buffer/DNA preserving media for PCR testing of swabs and inocula • Thermal cycler and PCR consumables.

4. Procedure for transmission A schematic representation of the different transmission methods described below is illustrated in Fig. 10.1. 4.1. Transmission by oral intubation PIT-tag D fish if they need to be selected and tracked individu­ ally afterwards, based on their infective status. 2. Starve both D and R fish for two days before the procedure. 1.

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Fig. 10.1 Schematic drawing of Enteromyxum scophthalmi transmission methods. Simultaneous cohabitation and effluent exposure (A) and oral transmission (B) are summarized. Donor fish (D) are represented larger and darker than recipient fish (R) in A. Credit: Fish Pathology group IATS.

Lightly anaesthetize R fish with clove oil for easy and stress-free handling. Turbot are quite tolerant to handling and minimum anaesthesia is needed. 4. Check the E. scophthalmi infective status of D fish non-lethally (NL). This can be done by: a. Rectal probing (see Fox et al., 2000; Palenzuela and Bartholomew, 2002) and subsequent PCR detection of the 18S ribosomal RNA gene. Because NL-qPCR will take sev­ eral hours to produce results, fish cannot be used the same day they have been sampled. b. Anal cannulation with a pipette and direct microscopic observation of the fresh smear with a drop of saline (Fig. 10.2). Usually, severely infected fish discharge a clear liquid towards the rectum following light abdominal mas­ sage. When present, a drop of this liquid can be smeared on a glass slide rather than cannulation. Fish yielding smears in which the parasite is readily detected in high numbers at low magnification (not higher than 200×) are considered good candidates for use as donors for the oral inoculum. Register the types of stages detected (trophozo­ ites, sporoblasts, spores) according to the stages described in Redondo et al. (2004), and the intensity, following a 3.

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Fig. 10.2 Fresh smear of an intestinal scraping with abundant proliferative stages of Enteromyxum scophthalmi. Credit: Fish Pathology group, IATS.

5.

6.

7.

8.

semi-quantitative conventional scale from 1+ to 6+, with the following ranges: 1+ = 1–5; 2+ = 6–10; 3+ = 11–25; 4+ = 26–50; 5+ = 51–100; 6+ >100 parasite stages per micro­ scope field observation at 120×. Although this is not an accurate procedure for evaluation of infection intensity, it allows a fast and convenient selection of donors just prior to their use. On the same day as inoculation, sacrifice D fish by deep anaesthesia (e.g. 3-aminobenzoic acid ethyl ester, MS-222, at 0.1 g/l) and exsanguinate with a syringe from the caudal vein, to avoid blood congestion in the visceral cavity. At necropsy, if the intestine is bloated with clear liquid, aspirate this by syringe and save in a tube on ice. Remove the intestine as a whole piece by cutting it from the end of the stomach to the rectum, and place it in a sterile Petri dish. Keep the tissues cool all times (e.g. over a cold bench or close to ice blocks), but avoid freezing or placing directly within ice. Make fresh smears of the intestine aspirate and scraping of small pieces of the intestinal mucosa taken at the pyloric caeca/ anterior intestine, the rectum and the middle intestine. Observe immediately under the microscope and evaluate infection status using the criteria described in step 4b. This allows selec­ tion of tissues with high parasite numbers and avoids unneces­ sary dilution of inoculum with non-infected segments. Usually the anterior intestine, pyloric caeca and rectum contain the highest numbers of parasites and yield the best inocula. The selected intestinal segments are opened in a disposable

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148 ♦  Fish Parasites

Petri dish, and the mucosal layer gently scraped with the blunt edge of a surgical blade to peel off the epithelial layer where the parasite is harboured. Dilute with sterile saline containing antibiotics as needed to facilitate scraping of epithelia, and add the liquid intestine aspirate collected in step 6 (if it contains parasites on examination in step 7). 9. Pool scrapings in a single 50- or 15-ml tube and keep cool until used. Take an aliquot of the inoculum for further observation under light microscopy or for qPCR if quantification of para­ sites is needed. Dilute as required to obtain the desired volume and concentration for the infection of all R fish (see challenge dose in section 5 and step 11 below). 10. Very gently anaesthetise R fish with clove oil and carefully introduce the pipette tip (with wide bore or previously cut at the bevel, if necessary, to dispense the viscous inoculum) into the mouth of the turbot (Fig. 10.1B). Penetrate gently per­ pendicular to the face as far as the oesophagus/stomach, where some resistance starts to be felt, to deliver the inoculum. The size of the pipette or pipette tip depends on the size of the fish. Usually fish >50 g can be intubated with a disposable plastic Pasteur pipette, whereas smaller fish can be inoculated directly with an automatic pipette with a pre-cut 200-µl tip. Larger fish (>250 g) can be intubated with an automatic dispenser pipette with a 1000-µl tip (wide bore). 11. Deliver an adequate volume of inoculum, depending on fish size: 0.5–1 ml is optimum in a wide range of R sizes, but smaller fish (20°C very well. Avoid very low water temperatures (e.g.